CPL
CHEMISTRY AND
ELSEVIER
Chemistry and Physics of Lipids
81 11996) 215 227
PHYSICS OF LIPIDS
Modulation of the activities of enzymes of membrane lipid
metabolism by non-bilayer-forming lipids I
Rosemary B. Cornell*, Rebecca S. Arnold
In.~titute ~1 Molecular Biology and Biochemistry and the Department of Chemistry. Simon Fraser University, Burm~h)', B.C.
V5A 1S6, Camuk~
Abstract
The activities of several enzymes which catalyze the synthesis or degradation of important lipid structural
components of membranes are influenced by non-bilayer-forming lipids. We present and critique the hypothesis that
enzymes that catalyze interconversions between bilayer- and non-bilayer-forming lipids are regulated by the lipid
packing perturbations of the non-bilayer-formingcomponent. The consequence of this regulation is lipid polymorphic
homeostasis. The enzymes which are modulated by non-bilayer components of membranes include a diglucosyl
diacylglycerol synthase from mycoplasma, secretory and cell-associated phospholipase A2s, phospholipase C,
phosphocholine cytidylyltransferase, phosphatidate phosphohydrolase and diacylglycerol kinase. The effect of the
non-lamellar lipids, both reversed phase-forming and miceIIar-forming, are discused in terms of bilayer packing strain,
which can effect enzyme-membrane associations, and lateral domain formation, which can modulate the effective
concentration of lipid activators.
Ahhreviations: DAG, diacylglycerol; PA, phosphatidic acid; PC, PE, PG, PS, PI, PIP2. (phosphatidyl)-choline. ethanolamine,
glycerol, serine, inositol, inositol bis phosphate; MGlcDAG, monoglucosyl diacylglycerol; DGIcDAG. diglucosyl diacylglycerol:
DSC, differential scanning calorimetry; L~, lamellar liquid crystalline phase; L/C, lamellar gel phase; tq~, hexagonal 1 phase; H~,
reversed hexagonal II phase; MLV, multilamellar vesicles: SUV, small unilamellar vesicles: LUV, large unilamellar vesicles: PLA2,
phospholipase A2; PLC, phospholipase C; CT, CTP, phosphocholine cytidylyltransferase; PAP, phosphatidate phosphohydrolase.
* Corresponding author. Institute of Molecular Biology and Biochemistry, and the Department of Chemistry, Simon Fraser
University, Burnaby, B.C. V5AIS6, Canada. Fax: + 604 291 5583, email: [email protected].
Research in the authors' lab on the cytidylyltransferase was supported by the Medical Research Council of Canada.
I)009-3084/96,'$15.00 :~ 1996 Elsevier Science Ireland Ltd. All rights reserved
Pll S0009- 3084196)02584-4
216
R.B. Cornell, R.S. Arnoht / C/wmisuT aml Ph3:~'ic,~O~ kipi~£ SI (1996)215 227
1. Introduction
Enzymes of membrane lipid metabolism should
be responsible for maintaining a lipid composition
compatible with an optimally functioning bilayer.
Therefore they could be sensitive to fluctuations in
the ratio of bilayer/non-bilayer-forming lipids, and
their activities could be modulated in ways that
would maintain a critical minimum content of
non-bilayer lipids, but prevent excessive accu- mulation. For example, to prevent excessive accumulation of non-bilayer lipids, a phospholipase that
degrades PC to D A G could be subject to inhibition
by the nonbilayer-forming product. Conversely, an
enzyme that catalyzes the conversion of D A G to a
bilayer-forming phospholipid could be stimulated
by lipids that prefer the reversed hexagonal (H,)
phase. Moreover, the mechanism of the regulation
could be a direct response by the enzyme to the lipid
packing perturbations associated with increases in
the non-bilayer-forming lipid. This kind of regulatory mechanism could be referred to as lipid
polymorphic homeostasis.
There is only one well-studied example among the
enzymes of lipid metabolism for which there is
evidence of in vivo regulation by the proportion of
non-lamellar lipids in the membrane. This is a
glucosyltransferase from a mycoplasma, which catalyses the synthesis of diglucosyl DAG. We will
begin this review describing the work on this system.
Secondly, we will describe in vitro studies on PC
phospholipases which suggest modulation by nonlamellar-forming lipids. Lastly, we will describe
work on a group of enzymes of phospholipid metabolism, the activities of which can be modulated by
non-bilayer-forming lipids, but for which no separate criteria for a mode of action involving non-bilayer phases have been documented. We will critique the lipid polymorphic homeostasis hypothesis.
2. Diglucosy! DAG synthase
The mycoplasma, Acholeplasma laidlaw& appears to have a mechanism for regulating the ratio
of reversed hexagonal (H.) vs. bilayer-forming
lipids. Enrichment of the cytoplasmic membrane
with unsaturated or saturated fatty acids (via
medium supplementation in the presence of avidin
or in the absence of pantetheine) resulted in marked
changes in the relative amounts of two glucolipids,
mono- and diglucosyl diacylglycerol (MGIcDAG
and DGlcDAG: [I]). MGlcDAG prefers the H u
phase, while DGlcDAG prefers lamellar phases.
This assignment is based on analysis of the pure
lipids or analysis in combination with Hu-forming
PE using eH-NMR, polarizing light microscopy, or
DSC [2 4]. The ratio of mono- to diglucosyl DAG
was v 2.4 in membranes enriched with palmitate
chains (80% of total fatty acyl chains), and ~ 0.5
in membranes enriched with oleate chains ( ~ 100%
of total fatty acyl chains) [1]. Addition of cholesterol
(H.-forming) to oleate-enriched membranes lowered this ratio to ~ 0.3. Lipid unsaturation also led
to increased PG content from ~ 13% to ~ 40% of
total lipid [1]. Since unsaturated fatty awl chains
and cholesterol promote H , phases in PE [5], it was
reasoned that the cell has evolved mechanisms to
prevent the accumulation of non-bilayer lipids by
stimulating the synthesis of a larger polar head
group to match the cross-sectional area occupied by
the hydrocarbon component [3].
Recently there has been some progress toward
understanding the regulation of the enzymes responsible for the changes in the ratio of the glycolipids in response to changes in the proportion of
non-lamellar components. The synthesis of the glucolipids occurs in two steps: glucose is transferred
from UDP glucose to DAG to form MGlcDAG,
followed by a second transfer to form DGIcDAG.
The two glucosyl transfers are catalyzed by separate
enzymes [6]. Activation of the DGlcDAG synthase
by non-lamellar forming lipids would constitute a
simple mechanism for maintaining a balance between non-lamellar and lamellar lipids.
Neither of these membrane-bound enzymes have
been purified, however their activities have been
studied following delipidation of whole cells and
reconstitution with lipid dispersions of various
compositions, which contain the lipid substrates,
DAG or MGIcDAG [7,8]. Reactivation of either
enzyme was dependent on anionic lipids. The best
activator was PG; activation saturated at 40-60
tool% depending oll the 'matrix' bilayer-forming
lipid used (PC or DGIcDAG; [7]). The PG requirement was not simply related to its negative charge
R.B. Cornell, R.S. Arnold / Chemisto' and PIlysics O! LipMs 81 (1996) 215 227
since other phospholipids with a single negative
charge were much less effective. PG is the predominant anionic phospholipid in A. laidlawii membranes. Activation of the glucolipid synthases by
PG could reflect a regulatory mechanism for maintaining a balance between the content of PG and
the neutral lipid, and thus the negative surface
charge density of the cells.
The relationship between non-lamellar forming
components and the activities of the glucolipid
synthases were explored using reconstituted crude
enzymes [8]. The activity of the DGIcDAG synthase, but not the MGIcDAG synthase was stimulated by Hu-forming lipids, such as 1,3-DAG,
MGIcDAG, a branched chain PC, and cholesterol
derivatives. The stimulation of DGlcDAG synthase
by 5% 1,3-DAG was blocked by >_ 10% C~6E8, a
nonionic micelle-forming amphiphile. This finding
links the stimulation by DAG to its H.-forming
tendencies. Comparisons between the changes in in
vitro rates of mono vs. diglucosyl DAG synthesis
and the changes in mass ratio in cells upon addition
of steroids suggested that the magnitude of the
enzymatic changes could account for the changes
in cellular membrane lipid composition [8]. These
findings support a homeostatic mechanism for
controlling the lipid composition at the level of the
DGlcDAG synthase.
The non-lamellar additives potentiated the activation of the DGIcDAG synthase by PG [8]. For
example, 15 tool% 1,3-DAG lowered the concentration of PG required for maximum stimulation
from > 50% to ~ 20%. That is, the negative charge
density requirement was lowered as the proportion
of H.-forming lipid increased. Possibly, the requirement for anionic phospholipids like PG might
be related to the generation of unilamellar vesicles,
which are spontaneously formed at high tool percent in vortexed lipid dispersions. However even
with unilamellar vesicles an absolute requirement
for PG was observed for the activation of
DGIcDAG synthase (Wieslander, personal communication). The mechanism of the reduction in
the PG tool% requirement by non-lamellar additives has not yet been determined. One possibility
would be the induction of domains of PG in
membranes enriched in H,-forming lipids. Here is
an example of a membrane enzyme in which the
activity is stabilized by both an H.-forming lipid
217
which promotes negative curvature, and a charged
phospholipid which promotes positive curvature.
Their effects appear to be cooperative. Two other
examples of cooperation between HI (micellar) and
Hu-forming lipid modulators will be discussed
below (Section 3.1 and Section 4).
The use of the MGDAG/DGlcDAG ratio as an
index of the membrane lipid phase preference has
been recently criticized from work on Acholeplasma
laidlawii strain B [9]. In this strain there are two
other prominent lipids: acylpolyprenyl glucoside,
which is very strongly HH-forming, and glycerylphosphoryl-diglucosyl DAG, which is strongly
H~-forming. Both of these varied as the fatty acid
degree of unsaturation was varied, and the changes
in MGIcDAG/DGlcDAG ratio alone was not a
good indicator of the non-lamellar tendencies of the
total membrane components [9]. These authors
stressed the need for more comprehensive analysis
of lipid compositional manipulations [4,9].
Two other procaryotic systems have been described for which there is evidence for regulation
of membrane lipid composition by the intrinsic
phase preferences of the component lipids: in
ClostrMium species, the ratio of PE to a glycerol
acetal derivative of PE decreased in response to
increases in the degree of fatty acid unsaturation
[10]. Since the unsaturated PE is H,-forming, and
the glycerol acetal species is bilayer-forming, these
changes can be rationalized in terms of lipid
polymorphic control [10,11]. Secondly, an E. coil
PE auxotroph is non-viable unless high concentrations of divalent cations such as Ca 2 + or Mg 2 + are
included in the medium [12], It has been proposed
that the combination of the divalent cation and the
high membrane contents of cardiolipin in these
strains allows cation-cardiolipin complexes that
functionally replace the PE as the non-bilayerforming component. Substitution of Ba~+ for
Mg 2* o r C a 2 + does not allow growth, yet it does
decrease the negative surface potential, and has
equivalent effects on acyl chain order: thus it was
argued that the important parameter for survival
of the bacteria was not related to the surface
potential or chain packing but rather to a certain
percentage of lipids with non-bilayer tbrming tendencies [12]. The enzyme sites responsive to lipid
polymorphism have not been identified in either of
these procaryotic systems.
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R.B. Cornell, R.S. Arnold / Chemistry and Physics o[' Lipids 81 (1996) 215 227
3. Phospholipases
The action of phospholipases on bilayer-forming phospholipids such as PC or PIP2 generates
nonbilayer forming lipid products. PLCs produce
DAG, a strong inverted phase (Hu) promotor,
and PLA2 produces lysophospholipids and fatty
acids, both of which prefer the H~ or micellar
phase. In the context of lipid polymorphic homeostatic control, one could predict that PLC would
be inhibited by Hn-forming lipids, whereas PLA2
would be stimulated. The effects of nonbilayer
lipids on the activities of several members of these
phospholipase families have been investigated in
conjunction with the effects on lipid phase behavior or other physical properties related to lipid
packing. Secretory phospholipase A2s have been
most thoroughly studied. Before discussing this
data, we will briefly review the catalytic mechanism, membrane interactions, and functional roles
of PLA2s.
3. I. Phospholipase A2
There are several classes of phospholipases that
cleave the sn-2 ester linkage of phospholipids. The
secreted PLA2s (sPLA2) from snake or bee
venom, from pancreas, or certain blood cells such
as macrophages or platelets are related in sequence and have a highly conserved structure [13].
Whereas the former enzymes function in digestion
and toxicity, the cell-associated sPLA2s may function in cell membrane phospholipid turnover.
Members of the sPLA2 group of 13-18 kDa
proteins have been crystallized, in some cases with
bound monomeric lipid substrate (or inhibitor) in
the active site [14-19]. The enzymes are soluble,
but catalysis is enhanced when the substrate is
provided in a lipid aggregate (micelle or bilayer)
rather than as a monomer in solution. The catalytic mechanism is well understood. Catalysis involves two principle and sequential binding steps:
(i) enzyme binds to a lipid interface; (ii) surfacebound enzyme binds substrate monomer [20,21].
The initial step enhances catalysis via reduction in
dimensionality in the search for substrate and via
small changes in enzyme conformation (which can
be detected by effects on the fluorescence of Trp3
[22]). After binding to the lipid surface the enzyme
undergoes a conformational change which can be
detected by changes in nOes of protons in the
N-terminal helix, the active site, and the C-terminal basic region of pancreatic PLA2 [23]. The
NMR-derived solution structures of pancreatic
PLA2 in the presence and absence of an inhibitor
reveal conformational changes at the active site
and reduced motion of the N-terminal z-helix
[24], which is involved in interfacial binding. The
activated enzyme draws in a substrate monomer
from the lipid surface into the active site so that
9 carbons of the sn-2 chain interact with the
enzyme [15,24,25]. The transition state is stabilized by a bound Ca 2+ ion [13,14]. After hydrolytic cleavage the products are released back
into the lipid layer [25].
The portion of the protein that binds to the
lipid surface is believed to be a surface surrounding the active site cleft that includes Trp-3 and
other aliphatic side chains of an N-terminal helix,
as well as Tyr-69, near the active site of the
pancreatic PLA2 [14,25 27]. The binding of pancreatic PLA2 to anionic lipid bilayers is much
more stable than to zwitterionic vesicles, and its
selectivity for anionic surfaces is decreased by
deletion of a surface loop, which is not present in
the less-selective venom PLA2s [28]. Lipid packing properties could affect either the initial surface
binding step, interaction with substrate monomer,
or both. The activity of pancreatic and venom
PLA2s is clearly enhanced at the lipid phase
transition temperature [22,29], and is enhanced by
lateral domain formation [30,31]. Venom PLA2s
bind optimally to DPPC bilayers in the gel phase,
so the binding per se does not require packing
defects. There is a delay between addition of the
enzyme to PL bilayers and hydrolysis, which is
related to the accumulation of the hydrolysis
products, free fatty acid and lyso PC [30,32]. The
resulting changes in packing perturbations increase access of the lipid-bound enzyme to its
phospholipid substrates. Thus, it is the substrate
accessibility that is most influenced by lateral
packing stress and/or domain formation.
The effects of various types of bilayer packing
stress on the activity of sPLA2 have been examined. Sen et al. [33] measured the rate of phospho-
R.B. Cornell, R.S. Arnold / Chemistry and Physics of Lit~ids 81 (1996~ 215- 227
lipid hydrolysis of pancreatic PLA2 as a function
of the tool% dil 8:2 PE with or without cholesterol
in MLVs or LUVs that were stabilized by high
pH. They correlated changes in activity with the
appearance of non-lamellar phases. The activity
dropped upon transformation to the H H phase.
The largest enhancements occurred at or just
prior to the L~ ~ H . transitions. Rate enhancements were directly related to the calculated bilayer curvature strain accumulating in the LUVs
prior to the H , transition. The packing stress was
suggested to perturb the interactions between the
phospholipids in the bilayer, thereby facilitating
binding of substrate monomers to enzyme.
Zidovetzki et al. [34] examined the response of
cobra venom, bee venom, and pancreatic PLA2s
to four different types of packing stress induced in
PC MLVs by various DAGs. As in the previous
study the effects were explained in terms of facilitation of the access of enzyme to phospholipid
substrate, specifically the release of substrate
monomers from the bulk lipid layer. (i) Transverse perturbation by short chain DAGs ( < 8
carbons) in DPPC inhibited activity, due to increases in acyl chain order near the bilayer surface
[35]. (ii) Lateral phase separation into regions of
differing composition and fluidity induced by
longer chain (C12-C16) saturated DAGs stimulated activity. The induction of lateral phase separation [35,36] would promote structural defects at
the gel ~ fluid phase boundaries. This would
enhance enzyme binding to the interface or substrate accessibility. (iii) Negative curvature stress
induced by unsaturated DAG in unsaturated PC
membranes [36] stimulated activity. Diolein activated PLA2 at temperatures below that required
for induction of H . and isotropic phases, thus it
was argued that the enzyme was sensitive to bilayer packing stress rather than actual non-lamellar structures. Conversion to an H , organization
is accompanied by an inhibition of PLA2 [33]. (iv)
Ordering of phospholipid side chains by addition
of diolein to DPPC [36] inhibited the PLA2s. Of
the three phospholipases, the pancreatic enzyme
was the most dependent on bilayer packing disruptions. It was argued that this is due to its poor
bilayer-penetrating power compared to the venom
PLA2s [37,38].
219
The above studies showed that Hn-promoting
lipids activate PLA2. On the surface this result
makes sense in the context of lipid polymorphic
homeostatic control. PLA2 activity reduces the
membrane content of a bilayer-forming lipid (PC)
and replaces it with two H~-forming lipids, fatty
acid and lysoPC. The effects of an H,-former
would be counterbalanced by Hcformers, and
homeostasis with respect to curvature would be
maintained. On the other hand, there is much
evidence that lysoPC (Hrformer) activates PLA2.
In a series of studies on snake venom PLA2,
Bell and colleagues demonstrated that the time
delay between initial binding to DPPC LUVs and
detection of enzyme activity was related to the
production of lysoPC and fatty acid. Addition of
lysoPC to the vesicles prior to introduction of
enzyme reduced this delay time [39]. The fatty
acid produced in these systems segregates into
domains [30,32], resulting in increased effective
lysoPC concentrations and lowered requirement
for exogenous lysoPC to activate PLA2 [31,32].
The requirement for lysoPC accumulation was
reduced by addition of di- and triacylglycerols,
but not by mono-acylglycerols [29]. The acylglycerols did not affect the PLA2-membrane surface
interaction [29]. The effects of -~ 3-7 mol% acylglycerols on PLA2 activity was dependent on their
ability to form lateral domains, rather than HH
phase [29]. These studies suggest that positive,
rather than negative curvature is a more physiologically relevant parameter for the activation of
PLA2, and that the effects of these near physiological levels of diacylglycerols are less related to
their H,-promoting tendencies than to domain
formation. At concentrations as low as 7.5 tool%,
dil6:0 DAG can phase separate from liquid crystalline DPPC as 1/1 DAG: PC complexes [40].
The lysoPC requirement for activation was also
reduced at the L/~ --. L~ phase transition temperature. The phase transition was postulated to enhance substrate accessibility by promoting
packing defects, and in this way could substitute
for the lysoPC.
Sheffield et al. [39] showed a temporal correlation between endogenous lysoPC accumulation
(generated via PLA2-catalyzed hydrolysis) and increases in the polarity of the interfacial region of
220
R.B. Cornell, R.S. Arnold ./Chemistry and Physics q[' Lipids" 81 (1996) 215 227
DPPC bilayers, as monitored by fluorescence
shifts of two interfacially located probes, Prodan
and Laurdan. Similar concentrations of exogenously added lysoPC produced similar shifts in
the spectra of the fluorescent probes. A probe
monitoring the order of the interior of the bilayer was not affected by endogenous or exogenous lysoPC. The sudden burst in enzyme
activity upon reaching ~ 18 mol% lysoPC was
ascribed to the generation of a critical positive
surface curvature. Positive bending strain would
be expected to weaken lipid-lipid interactions
near the membrane surface and thus would improve access of the enzyme to the sn-2 ester
bond, whereas negative curvature would tend to
be associated with tighter packing at the surface.
Is there any relation of this work on the secretory PLA2s to the cell-associated PLA2 which
would have a larger role in maintenance of PC
homeostasis? Several reports document stimulation of cellular PLA2s by diacylglycerol. The hydrolysis of phospholipids by crude PLA2 from
platelets [41,42], or intestinal mucosa [43] was
enhanced by various DAGs. Only small rate enhancements ( < 2-fold) were observed with low
(2-5 mol%) D A G [42]. None of these studies
employed pure enzymes, thus the mechanism of
the D A G effects could be very indirect. In other
studies with neutrophils, in which PLA2 mediated arachidonate release is stimulated by the
chemotactic peptide fMLP, 1,2-diC8 and 1,3diC8 DAGs stimulated arachidonate release, but
D A G treatment alone (i.e. without fMLP) did
not activate PL hydrolysis [44]. Thus the effects
of these DAGs on the membrane alone are not
sufficient to activate the phospholipase, but may
cooperate with other signalling mechanisms.
However, these effects do agree in principle with
the more rigorously studied effects of DAGs on
the secretory PLA2s, and thus suggest that the in
vitro effects may have some physiological relevance.
The 85 kDa arachidonate-specific cPLA2 (class
III) translocates from cytosol to nuclear membranes in response to increased cytosolic Ca 2÷
[45,46]. The membrane-binding or interfacial
recognition site is an N-terminally located calcium binding CaL B domain, homologous to the
C 2 domain of PLQq or the classical protein kinase C isoforms. Partially purified enzyme was
stimulated by acidic phospholipids, DAG, or unsaturated PE included in sonicated vesicles containing an arachidonoyl PC [47]. The negatively
charged phospholipids lowered the Ca 2 ÷ requirement for activation. From the effects of DAG
and PE, the authors concluded that lipid packing
density is important for the activity of this enzyme. Burke et al. [48] studied the effects of
arachidonoyl-containing substrate, PAPC, using
pure recombinant 85 KDa cPLA2 and SUVs
composed of dimyristoyl phosphatidylmethanol
(DMPM) to promote lipid binding. PAPC had
cooperative effects on activity between 0 10
mol%, and the cooperativity was specific for the
sn-2 arachidonoyl chain. PAPC caused an increase in the mean molecular area in the compressed (gel) state of monolayers from 31 + 3
A 2 for D M P M to 40 + 3 fik 2 for D M P M / P A P C
(9/1). It was suggested that the expansion due to
DAPC could cooperatively promote abstraction
of substrate monomers into the active site. The
possibility was also raised that lateral phase separation of the PAPC substrate could generate an
interfacial recognition site comprised of a PAPC
cluster [48]. However, the CaL B domain is distinct from the catalytic site (site of PAPC binding), and is selective for acidic lipids. Additional
studies with pure cPLA2 and fully characterized
lipid phases would be of interest to compare the
mechanism of this enzyme with the structurally
divergent secreted PLA2s.
3.2. Phospholipase C
Phospholipase Cs catalyze cleavage of the
phosphate ester to generate D A G and the phosphorylated head group of the phospholipid.
PLCs specific for PIP2 are perhaps the most interesting from a metabolic standpoint, however
less-specific PLC from bacterial sources have
been studied more intensely due to ready access
to the enzyme. In the context of lipid polymorphic regulation one would predict that an enzyme that catalyses increases in an H,-former,
DAG, would be subject to negative regulation by
Hu-forming components, Rao and Sundaram
R.B. Cornell, R.S. Arnold/Chemist O, and Physics ~! LipMs 81 (1996) 215 227
[49] investigated the response of PLC from B.
cereus
to perturbations in lipid packing. They
monitored both the phase behavior and enzyme
activities using SUVs and MLVs containing
mixtures of DOPC with DOPE, cholesterol,
lysoPC or gramcidin. DOPE, cholesterol, and
gramcidin, the Hn-formers, were negatively correlated with enzyme activity, whereas lysoPC
was positively correlated with activity. For each
system there was a direct correlation between
the activity and the percent bilayer component
acquired from 3~P-NMR spectra. Interestingly
the activation by lysoPC peaked at ~ 20 mol%,
which is close to the lysoPC content which produced optimal rates of PLA2 "activity [39]. These
results were interpreted in a straight-forward
way: i.e. negative curvature interferes with PLC
access to the substrate's phosphoester bond, and
positive curvature enhances it.
These effects of PLC differ from those of the
secretory PLA2s, which were stimulated by unsaturated PE [33] and by gramcidin [50]. Like
PEA2, PLC has much higher activity towards
micellar vs. monomeric substrate, and there is a
hydrophobic surface near the active site, which
could function in binding to the lipid-water interface. However, unlike PLA2, there is i1o requirement for PLC to draw a phospholipid
monomer into its active site since the bond to
be hydrolyzed is accessible from the surface.
The active site is not especially hydrophobic
[51]. The rate limiting step in the reaction is
probably the release of the hydrophobic
product, DAG, from the active site. The stimulatory effect of lysoPC seen by Rao [50] could
be due in part to a detergent effect, which could
stimulate the release of DAG [52]. D A G inhibits
PLC via product inhibition [53], but in light of
the general inhibition by Hn-promoting components [50], DAG may also effect the activity via
changes in the phase properties of the lipid
layer.
We now consider three enzymes of phospholipid metabolism, whose activities are sensitive to
non-lamellar forming lipids: cytidylyltransferase,
phosphatidate phosphatase, and diacylglycerol kinase. As yet rigorous analyses of lipid phase
dependencies for these activities are lacking.
221
4. Cytidylyltransferase
CTP:phosphocholine cytidylyltransferase (CT)
catalyzes the condensation of CTP and phosphocholine to generate CDP-choline, the head
group donor for PC. In animal cells, this reaction is, under most conditions, the rate-limiting
step in PC synthesis. The enzyme requires lipids
for activity. In cells, the majority of CT is in a
soluble compartment, recently identified as the
nucleosol [54], and is inactive. It can be recruited to the inner nuclear membrane where it
is activated in response to stimulants of PC synthesis [55,56]. Agents or conditions which promote this translocation in cell culture systems
include fatty acids, short chain DAGs, phospholipase C (probably working via generation of
DAG), choline deprivation, and phorbol esters
[55,56]. Although there is evidence that the
phosphorylation state of CT influences membrane-association, its role is secondary to the
influence of lipid compositional changes [57,58].
The lipid polymorphism control hypothesis
would argue that CT, which controls the rate of
synthesis of the primary bilayer-forming component, PC, should be subject to regulation by the
relative proportions of bilayer vs. non-bilayer
lipids. It should be activated by DAG or other
lipids with non-bilayer propensities, and inhibited
by PC or other bilayer-forming lipids. Jamil et al.
[59] suggested that CT translocation to cell membranes is modulated by the ratio of bilayer:nonbilayer forming lipids. They arrived at this
conclusion from correlations between CT binding
to cell membranes and changes in membrane contents of PC, DAG, PE, mono- and dimethyl PE,
and oleic acid. The data were derived mostly from
cell culture experiments, and did not emphasize
the effects of anionic phospholipids, which have
been implicated as activators of CT only using
pure vesicle systems. CT association with membranes increased with a reduction in the relative
PC content, or an increase in DAG, PE, or oleic
acid. For example, in choline-deprived vs. cholinesupplemented hepatocytes, the PC content varied
20 25%, and there was a close inverse correlation
between the membrane PC content and the
amount of membrane-bound, active CT [60,61].
222
R.B. Cornell, R,S. Arnold / Chemistry and Physics o! Lipids 81 (1996) 215 227
Jamil et al. [59] made an interesting discovery:
LysoPC could reverse the effects of DAG on
CT-membrane binding. DAG was generated to
3 mol% in situ by PLC treatment, but the
effective lysoPC content was not quantitated. If
DAG effects were mediated via negative curvature
strain, the addition of the H~-forming lysoPC
would negate the curvature effects of DAG. Since
the membrane contents of DAG, lysoPC, and PC
affected the CT membrane-binding step, this argues for effects on membrane properties rather
than intervention at CT's active site.
Rat liver CT has been purified, cloned, and
expressed [62-64]. Its activity can be modulated
in vitro by pure lipids. There are two classes of
lipid activators: Class I is comprised of anionic
lipids such as fatty acids and acidic phospholipids,
and Class II is comprised of DAG and related
compounds [65,66]. Whereas class I activators can
activate CT as components of a Triton micelle or
PC vesicle, class II activators are only effective as
components of a bilayer system [65]. This suggests
that the effect of DAG on lipid packing in a
bilayer which is recognized by CT cannot be
reproduced in a micellar organization. Induction
of negative curvature strain may be one such
effect. The counteractive effects of lysoPC and
DAG on CT translocation in hepatocytes [59]
were reproduced in pure vesicles. LysoPC reduced
the activation by 20 mol% DAG when included at
equimolar or greater concentrations with respect
to DAG in egg PC vesicles (Arnold and Cornell,
unpublished data). LysoPC did not however reduce the activating potential of oleic acid [67], and
had only minor inhibitory effects on PC alone
(Arnold and Cornell, unpublished data). These
results imply a role for the curvature effects of
DAG, but further testing of this idea is required.
The potency of the class I activators is also
related to curvature; SUVs composed of PG/PC
(1:3) were effective at micromolar concentrations,
whereas MLVs were activating only at millimolar
concentrations [66]. The dependence of CT activity on the lipid phase was determined by variation
of temperature in the presence of PC/PG MLVs
of defined acyl chain length. Activity was low in
the gel state, higher in the liquid crystalline state,
but was anomalously high at temperatures near
the phase transition, which was monitored by
DSC [66]. This suggested that CT activation was
sensitive to lipid packing perturbations. CT can
bind to PG/PC MLVs in the gel state, although
the nature of the binding is electrostatic and the
enzyme is inactive [68]. Thus binding and activation are likely sequential, independent steps.
The mechanism of CT interactions with lipid
vesicles is fairly well understood. The membranebinding domain consists of an amphlpathic u-helix, approximately 60 amino acids long, located
between the catalytic domain and the C-terminal
phosphorylation domain. Abundant evidence
identifies this region (domain M) as the membrane-binding domain. Proteolytic fragments [69]
or genetically engineered mutants [57,70,71] lacking this domain lack membrane-binding capacity.
Synthetic peptides corresponding to portions or
all of domain M bind as a-helices to anionic
vesicles [72,73]. Intercalation of domain M has
been demonstrated using the intact enzyme and
lipid photolabels (Johnson et al., submitted for
publication). Using brominated phospholipids as
depth quenchers of peptide tryptophan groups,
Johnson and Corneli [72,73] showed that the peptides derived from domain M intercalate half-way
into the outer monolayer of the vesicles when they
bind. In water the peptides adopt a random coil
structure. Binding, intercalation, and stabilization
of an a-helical structure is selective for anionic
vesicles. These peptides do not interact with
DAG-containing vesicles; however other data
strongly imply a requirement for this domain in
the interaction of the whole enzyme with DAGcontaining vesicles [69,70] (also Johnson et al.,
submitted for publication). At this time the identity of other CT domain(s) required for the interaction with DAG is not known. The
NMR-derived structure of domain M bound to
SDS micelles reveals an uninterrupted amphipathic helix with a strip of basic residues
aligned at one polar/nonpolar interface (Dunne et
al., submitted for publication). The positively
charged strip is likely the primary reason for the
specificity towards anionic lipids. These and other
data suggest that the mechanism for activation of
CT involves enrichment of membranes with lipids
that facilitate the intercalation of the nonpolar
R.B. Cornell, R.S. Arnold / Chemistry and Physics ~! Lipids 81 (1996) 215 227
face of a long amphipathic helix, which is a
prerequisite for conformational alterations of the
adjacent catalytic domain.
How do class 1 and class II lipids facilitate this
intercalation? In addition to the generation of a
negative surface charge which would attract and
stabilize the positively charged face of the helix,
class I lipids would also decrease the packing
density at the surface due to charge repulsion of
the head groups. This latter feature would facilitate intercalation of the hydrophobic amino acids.
DAG and other class II lipids, on the other hand,
would not provide any charge-charge stabilization, but would create surface packing deformities, i.e. 'holes' in the regular surface of PC.
Moreover, it is possible, from consideration of the
geometry of the CT amphipathic helix [74], that
CT binding would relieve the negative curvature
strain produced by the DAG, such that the free
energy of the system would decrease upon CT
binding to DAG-enriched membranes. Since the
amphipathic CT peptides do not bind to PCDAG vesicles, another portion of the enzyme
must also contribute to the DAG-responsiveness
via a separate mechanism.
The concentrations of the activators required
for substantial activation of CT in both the cell
and in vitro systems are higher than that normally
found in membranes of unstimulated cells. For
example, complete membrane translocation of CT
in HeLa cells treated with oleic acid required
enrichment to approximately 20 tool% of total
phospholipid [75]. Full activation of pure CT by
SUVs required 5 mol% cardiolipin, 10 tool% PA,
or ~20 tool% PG, and 15 mol% DAG [68].
However when combinations of class I and class
II activators were tested, synergism was observed.
Activation was 60% of maximal at 2.5 mol%
PA + 2.5 tool% DAG [68]. These concentrations
are close to those observed in stimulated cells.
The mechanism of the synergistic effects is unknown. It may involve increased disruption of the
regular packing above that achieved by low mol%
of the individual lipids. Alternatively, DAG may
induce lateral segregation and domain formation
of the acidic phospholipid creating a higher effective concentration, as has been proposed for
PLA2 activation by lysoPC [39].
223
5. Phosphatidate phosphohydrolase and
diacylglyceroi kinase
Phosphatidate phosphohydrolase (PAP) and
DAG kinase catalyze interconversions between
PA, a bilayer-forming phospholipid, and DAG,
an Hu-forming lipid. Although both PA and
DAG are minor components of cell membranes
(generally < 2 tool% each), only a few tool%
DAG is sufficient to induce curvature strain in
bilayers [76]. Thus DAG kinase and PAP might
be subject to positive and negative regulation,
respectively, by DAG or other Hu-forming lipids.
PAP catalyzes the dephosphorylation of PA to
DAG. In eucaryotes this reaction occupies a key
branching point in phospholipid synthesis, between acidic and zwitterionic phospholipids [77].
PAP is the committed step for PE and PC synthesis. The dephosphorylation of PA is probably the
major route of DAG production in unstimulated
cells, and in cells stimulated with agonists that
promote PC turnover via phospholipase D
[78,79].
The regulatory properties of PAP enzymes from
yeast, chloroplasts, and mammalian cells have
been explored. Enzymes from yeast, pig thymus,
and rat liver have been purified [80-82], and it is
clear that different enzymes are utilized by the
different sources to catalyze the same reaction.
Although there have been studies suggesting that
PAP enzymes can be modulated by non-bilayer
forming lipids such as PE, DAG, fatty acids, or
sphingosine and related amphiphilic amines
[77,80,81,83 85], none of these studies paid attention to the effects of the various lipids on the
physical properties of the lipid preparations.
The yeast and chloroplast enzymes are inhibited
by 1,2-DAGs [80,83], but the inhibitory effects
were observed using a Triton mixed micelle system to disperse lipid modulators as well as the PA
substrate. Thus, the effects of DAG can not be
restricted to a mechanism involving defects in
bilayer packing. Furthermore, although 1,2DAGs were inhibitory, other similar Hn-forming
species such as 1,3-DAG or monoacylglycerol
were not. The mechanism of inhibition by DAG
appears to be via competitive inhibition of the PA
substrate, i.e. product inhibition [83].
224
R.B. Cornell, R.S. Arnold / Chemistry and Physkw ~?['Lipids 81 (1996) 215 -227
There are at least two distinct mammalian
PAPs: PAP-l, a form that is distributed between
cytosol and microsomes and is considered to be
involved in glycerolipid synthesis and PAP-2, a
predominantly plasma membrane-associated integral enzyme believed to be the signal regulated
form (producing DAG as a second messenger).
The two PAPs are also distinguished by sensitivities to N-ethylmaleimide and Mg 2+ [86]. PAP-1
can be translocated to membranes in response to
treatment of cells with fatty acid, and this translocation can be blocked by amphiphilic amines,
suggesting a role for negative charge in the membrane binding event [84]. An 83 kDa PAP-2 type
enzyme was inhibited by 1,2-DAG, but not by
mono- or triacylglycerol when provided as components of a Triton micelle [81]. The effects of
non-bilayer-forming lipids on mammalian PAP
activity has not been examined using bilayer systems monitored for effects on phase behavior.
DAG kinase catalyzes the phosphorylation of
DAG to produce PA. It functions in the PI cycle
in animal cells, recycling DAG back to PI via PA
and CDP-DAG intermediates [87]. Thus far, three
homologous mammalian isoforms have been
cloned [88-90], as well as a structurally distinct
DAG kinase from E. coli [91]. In the mammalian
enzymes, the DAG binding site is likely a zinc
butterfly motif, very like the domain in PKC
responsible for DAG interaction [88]. It also contains a 33-residue amphipathic helix, which has
been speculated as a membrane-binding site [92].
In addition to interacting with the enzyme as
substrate, DAG could modify the activity of the
kinase via effects on lipid phase properties. As
with PAP, this possibility has not been examined
in suitable model bilayer systems. Translocation
to cell membranes in response to elevated DAG
content has been reported for rat brain DAG
kinase [93]. The E. coli enzyme was activated by
both 1,2- and 1,3-DAG using a mixed micelle
assay system [94]. It could be argued that enzymes
like PAP or DAG kinase are more likely to be
regulated by chemical signals or kinase/phosphatase action, rather than by membrane physical
parameters, especially since they are not involved
in conversions that involve major lipid structural
components of membranes, but rather lipids that
serve regulatory functions. On the other hand,
regulation of metabolic enzymes is multi-layered,
thus regulation linked to lipid polymorphic control merits an investigation.
PE is the major non-bilayer component of
membranes. The enzymes which produce or metabolize PE could be subject to regulation by the
proportion of non-bilayer lipids. These enzymes
would include PS decarboxylase, PE methyltransferase, which have been purified [95,96], and
ethanolamine phosphotransferase which has not
been purified but has been cloned from yeast
[97,98]. However no direct modulation of these
activities by non-bilayer lipids have been reported.
Clearly, this is an area which could benefit from
study of activity modulation of purified enzymes
by changes in lipid phase and/or packing properties.
6. Summary and conclusions
The general prediction of lipid polymorphic
regulation is that enymes which catalyze conversion between bilayer and non-bilayer forming
lipids would be negatively regulated by the phase
preference of the product lipid. The lipid regulation of the diglucosyl DAG synthase, phospholipase C, and CT provide evidence in support of
this idea. The DGlcDAG synthase and CT are
examples of enzymes regulating the formation of
bilayer-forming lipids from non-bilayer lipids, and
they are activated by non-bilayer forming lipids.
PLC catalyzes the conversion of a bilayer-former
to a non-bilayer (H~) former, and is inhibited by
HH forming lipids. Regulation of the sPLA2s
appears more complex. The products of its reaction (Hi-forming) stimulate enzyme activity. Activation by the Hu-forming lipid, DAG, probably
occurs via promotion of lateral domain formation
of the Hrforming activators. The secretory
PLA2s might not be expected to contribute to
bilayer homeostasis. In fact they evolved to destroy their target bilayers.
Several important themes emerge from the
work reviewed. First, there is no evidence for an
enzyme of lipid metabolism that is preferentially
activated by lipids in the Hu phase (PLC is in fact
R.B. Cornell, R.S. Arnohl / Chemist~3' and Physics (~/ Lipids gl (1996) 215 227
i n h i b i t e d by t h e H H phase). R a t h e r , s t i m u l a t i o n
by H n - p r o m o t i n g lipids is d u e to t h e p a c k i n g
s t r a i n o r p h a s e s e p a r a t i o n s i n d u c e d w i t h i n the
bilayer arrangement. Second, a recurring theme
for r e g u l a t i o n o f e n z y m e s t h a t b i n d r e v e r s i b l y to
the m e m b r a n e s u r f a c e ( p h o s p h o l i p a s e s , C T , P A P )
is t h a t lipids w h i c h i n d u c e p a c k i n g d e f e c t s will
stimulate binding and/or activation. This group
c a n be fully a c t i v a t e d by lipids in m i c e l l a r arr a n g e m e n t , a n d by i n c r e a s i n g p o s i t i v e c u r v a t u r e
in bilayers.
Third, activation of these enzymes commonly
i n v o l v e s c o o p e r a t i v e effects o f H i - a n d H ~ - f o r m ing lipids. T h e m e c h a n i s m s o f the c o o p e r a t i v i t y
are n o t well u n d e r s t o o d . D A G m a y c a u s e p h a s e
separations that increase the effective concentration of the lysoPC activator of PLA2 or the
a n i o n i c lipid a c t i v a t o r s o f C T . In the case o f the
D G l c D A G s y n t h a s e , H ~ r f o r m i n g lipids m a y also
p r o m o t e d o m a i n f o r m a t i o n o f the a c t i v a t o r P G .
Lastly, t h e r e are m a n y e n z y m e s w h i c h c a t a l y z e
conversions between lamellar- and non-lamellarf o r m i n g lipids w h i c h h a v e n o t b e e n subject to
e x a m i n a t i o n for t h e effects o f p u r t u r b a t i o n s o f
b i l a y e r o r g a n i z a t i o n . F o r m o s t , c D N A s are available for expression and purification. The work
awaits a curious biophysicist,
References
[I] A. Wieslander, A. Christiansson, L. Rilfors and G. Lindblom (1980) Biochemistry 19, 3650 3655.
[2] /~. Wieslander, L. Rilfors, L. Johansson and G. Lindblom
(1981) Biochemistry 20, 730 735.
[3] ,A. Wieslander, A. Christiansson, L. Rilfors, A. Khan, L.
Johansson and G. Lindblom (1981) FEBS Len. 124,
273 278.
14] P.J. Foht, Q.M. Tran, R.N.A.H. Lewis and R.N. McEIhaney (1995) Biochemistry 34, 13811-13817.
[5] J. Gallay and B. de Kruijff (1982) FEBS Len. 143,
133 136.
[6] A. Dahlqvist. S, Andersson and /~. Wieslander (1992)
Biochim. Biophys. Acta 1105, 131 140.
[7] O.P. Karlsson, A. Dahlqvist and A. Wieslander (1994) J.
Biol. Chem. 269, 23484 23490.
[8] A. Dahlqvist, S. Nordstr6m, O.P. Karlsson, D,A. Mannook, R.N. McElhaney and A,. Wieslander (1995) Biochemistry 34, 13381 13389.
[9] R.N.A.H, Lewis and R.N. McElhaney (1995) Biochemistry 34. 13818 13824.
225
[10] H. Goldfine and N.C. Johnston (1987) Biochemistry 26,
2814 2822.
[I 1] H. Goldfine (1985) Curr. Top. Cell. Regul. 26, 163 174.
[12] A.G. Rietveld, V.V. Chupin, M.C. Koorengevel, H.L.J.
Wienk, W. Dowhan and B. de Kruijff (1994) J. Biol.
Chem. 269, 28670 28675.
[131 E.A. Dennis (1994) J. Biol. Chem. 269. 13057 13060.
[14] S.P. White, D.L. Scott, Z. Otwinowski, M.H. Gelb and
P.B. Sigler (1990) Science 250, 1560 1563.
[15] D.L. Scott. Z. Otwinowski, M.H. Gelb and P.B. Sigler
(1990) Science 250, 1563 1566.
[16] J.P. Wery, R.W. Schevitz. D.K. ('lawson, L.L. Bobbin,
E.R. Dow. G. Gamboa, T. Goodson, R.B. ttermam
R.M. Kramer, D.B, McClure, ED. Mihelich, J.E. Putman. J.D. Sharp, D.H. Stark, C. Teater, M.W. Warrick
and N.D. Jones (1991) Nature 352, 79 82.
[171 D.H. Freemont, D. Anderson, I.A, Wilson, E.A. Dennis
and N-H. Xuong (1993) Proc. Natl. AcaJ. Sci. USA 90,
342 346.
[18] S. Brunie, J. Bolin, D. Gewirth and PB. Sigler (1985) J.
Biol. Chem. 260, 9742 9749.
[19] B.W. Dijkstra, K.H. Kalk, W.G. ttol and J. Drenth
(198l) J. Mol. Biol, 147, 97 I23.
[20} R. Verger (1976)Atom, Rev. Biophys. Bioeng. 5. 77 117.
[21] M.H. Gelb, M.K. Jain, A.M. Hanel and O.G. Berg (1995)
Annu. Re~. Biochem. 64, 653 688.
[221 ,I.D. Bell and R.L. Biltonen (1989)J. Biol. Chem. 264.
225 230,
[23] P.M. Kilby, W.U. Primrose and G.C.K. Roberts (1995)
Biochem. J. 305. 935 944.
[24] B. van den Berg, M. Tessari. R. Boelens, R. Dijkman,
G.H. de Haas, R. Kaptein and H.M. Verheij (1995)
Nature Struct. Biol. 2, 402 406.
[25] D.L. Scott, S.P. White. Z. Otwinowski, W. Yuan, M.H.
(]elb and P,B. Sigler (1990) Science 25(I, 1541 1546.
[26] O.P. Kuipers, M. Vincent. J.-C. Brochon, H.M. Verheij,
G.It. de Haas and J. Gallay (1992) Biochemistry 30,
8771 8785.
[27] A.R. Peters, N. Dekker. L. van den Berg, R. Boelens, R.
Kaptein, A.J. Slotboom and G.tt. de Haas (1992) Biochemistry 31, 10024 10030.
[28] O.P. Kuipers. M.M.G.M, Thunnissen, P. de Geus, B.W.
Dijkstra. J. Drenth, ll.M. Verheij and G.H. de Haas
(1989) Science 244, 82 85.
[29] ,I.D. Bell, M.L. Baker. E.D. Bent, R.W. Ashton, D.,I.B,
Hemming and L,D. Hansen (1995) Biochemistry 34,
11551 11560.
[30] M.K. Jain, B.-Z. Yu and A. Kozubek (1989) Biochim.
Biophys. Acta 980. 23 32.
[31] E.D. Bent and J.D. Bell (19951 Biochim. Biophys. Acta
1254. 349 360.
[32] W.R. Burack, Q. Yuan and R.L. Biltonen (19931 Biochemistr~ 32. 583 589.
[33] A. Sen, T.V. lsac and S.-W. Hui (1991) Biochemistry 30,
4516 4521.
[34] R. Zidovetzki, L. Laptalo and J. Crawford (1992) Biochemistry 31, 7683 7691.
226
R.B. C)*rnell, R.S. Arnold / Chemistry and Physics ~/ Lipids 81 (1996) 215 227
[35] H. De Boeck and R. Zidovetzki (1992) Biochemistry 31,
623 630.
[36] H. De Boeck and R. Zidovetzki (1989) Biochemistry 28,
7439- 7446.
[37] M.K. Jain, M.R. Egmond, H.M. Verheij, R. ApitzCastro, R. Dijkman and G.H. de Haas (1982) Biochim.
Biophys. Acta 688, 341-348.
[38] N. Gheriani-Gruszka, S. Almog, R.L. Biltonen and D.
Lichtenberg (1988) J. Biol. Chem. 263, 11808 11813.
[39] M.J. Sheffield, B.L. Baker, D. Li, N.L. Owen, M.L.
Baker and J.D. Bell (1995) Biochemistry 34, 7796 7806.
[40] P.J. Quinn, H. Takahashi and 1. Hatta (1995) Biophys. J.
68, 1374-1382.
[41] R.M.C. Dawson, N.L. Hemington and R.F. lrvine (1983)
Biochem. Biophys. Res. Commun. 117, 196 201.
[42] R.M. Kramer, G.C. Checani and D. Deykin (1987)
Biochem. J. 248, 779 783.
[43] R.M.C. Dawson, R.F. Irvine, J. Bray and P.J. Quinn
(1984) Biochem. Biophys. Res. Commun. 125, 836 842.
[44] S.A. Bauldry, R.L Wykle and D.A. Bass (1988) J. Biol.
Chem. 263, 16787- 16795.
[45] S. Glover, M. de Carvalbo, T. Bayburt, M. Jonas, E. Chi.
C.C. Leslie and M.H. Gelb (1995) J. Biol. Chem. 270.
15398-15367.
[46] A.R. Schievella, M.K. Regier, W.L. Smith and L.L. Lin
(1995) J. Biol. Chem, 270, 30749 30754.
[47] C.C. Leslie and J.Y. Channon (1990) Biochim. Biophys.
Acta 1045, 261 270.
[48] J.R. Burke, M.R. Witmer, J. Tredup, R. Micanovic, K.R.
Gregor, J. Lahiri, K.M. Tramposch and J.J. Villafranca
(1995) Biochemistry 34, 15165-15174.
[49] N.M. Rao and C.S. Sundaram (1993) Biochemistry 32,
8547 8552.
[50] N.M. Rao (1992) Biochem. Biophys. Res. Commun. 182,
682-688.
[51] S. Hansen, E. Hough, L.A. Svensson, Wong, Y.-L. and
S.F. Martin (1993) J. Mol. Biol. 234, 179-187.
[52] M.Y. El-SaTed and M.F. Roberts (1985) Biochim. Biophys. Acta 831, 133 141.
[531 R.A. Burns, Jr, J.M. Friedman and M.F. Roberts (1981)
Biochemistry 20, 5945 5950.
[54] Y. Wang, T.D. Sweitzer, P.A. Weinhold and C. Kent
(1993) J. Biol. Chem. 268, 5899-5904.
[55] H. Tronch6re, M. Record, F. Terce and H. Chap (1994)
Biochim. Biophys. Acta 1212, 137 151.
[56] R.B. Cornell (1996) in: R.A. Gross (Ed.), Advances in
Lipobiology, Vol. l, JA1 Press (in press).
[57] Y. Wang and C. Kent (1995) J. Biol. Chem. 270, 354360.
[58] M. Houweling, H. Jamil, G. Hatch and D. Vance (1994)
J. Biol. Chem. 269, 7544-7551.
[59] H. Jamil, G.M. Hatch and D.E. Vance (1993) Biochem. J.
285, 815--820.
[60] H. Jamil, Z. Yao and D.E. Vance (1990) J. Biol. Chem.
265, 4332-4339.
[61] P. Weinhold, L. Charles and D. Feldman (1994) Biochim.
Biophys. Acta 1210, 335 347.
[62] P.A. Weinhold, M.E. Rounsifer and D. Feldman (1986) J.
Biol. Chem. 261, 5104 5l 10.
[63] J. MacDonald ~ind C. Ke,mt (1993) Prot. Express. Purif. 4,
l
7.
[64] G. Kalmar, R.J. Kay, A. LaChance, R. Aebersold and
R.B. Cornell (1990) Proc. Natl. Acad. Sci. USA 87,
6029-6033.
[65] R.B. Cornell (1991) Biochemistry 30, 5873 5880.
[66] R.B. Cornell (1991) Biochemistry 30, 5881 5888.
[67] P.S. Sohal and R.B. Cornell (1990) J. Biol. Chem. 265,
11746 11750.
[68] R.S. Arnold and R.B. Cornell (1996) Biochemistry (in
press).
[69] L. Craig, J.E. Johnson and R.B. Cornell (1994) J. Biol.
Chem. 269, 331l 3317.
[70] R. Cornell, G. Kalmar, R. Kay, M. Johnson, J. Sanghera
and S. Pelech (1995) Biochem. J. 310, 699 708.
[71] W. Yang, K. Boggs and S. Jackowski (1995) J. Biol.
Chem. 270, 23951 23958.
[72] J.E. Johnson and R.B. Cornell (1994) Biochemistry 33,
4327 4335.
[73] J.E. Johnson (1995) Ph,D. Thesis, Simon Fraser University.
[74] R.M. Epand, Y. Shai, J.P. Segrest and G.M. Anantharamaiah (1995) Biopolymers 37, 319 338.
[75] R.B. Cornell and D.E. Vance (1987) Biochim. Biophys.
Acta 919, 2 6 36.
[76] R.M. Epand and R.F. Epand (1994) Biophys. J. 66,
1450 1456.
[77] W.I. Wu and G.M. Carman (1996) Biochemistry 35.
3790 3796.
[78] M.M. Billah, S. Eckel, Y.J. Mullmann, R.W. Egan and
M.I. Siegel (1989),I. Biol. Chem. 264, 17069-17077.
[79] M.C. Chabot. L.C. McPhail, R.L. Wykle, D.A. Kennerly
and C.E. McCall {1992) Biochem. J. 286. 693 699.
[80] Y.-P. Lin and G,M. Carman (1989) J. Biol. Chem. 264,
8641 8645.
[81] H. Kanoh, S.-i. Imai, F. Yamada and Sakane (1992) J.
Biol. Chem. 267, 25309 -25314.
[82] D.W. Waggoner, A. Martin, J. Dewald, A. GomezMulSoz and D.N. Brindley (1995) J. Biol. Chem. 270,
19422 19429.
[83] A. Ma[herbe, M.A. Block, J. Joyard and R. Douce (1992)
J. Biol. Chem, 267, 23546 23553.
[84] R. Hopewell, P. Martin-Sanz, A. Martin, J. Saxton and
D.N. Brindley (1985) Biochem. J. 232, 485 491.
[85] A. G6mez-Mufioz, E.H. Hamza and D.N. Brindley
11992) Biochim. Biophys. Acta 1127, 49 56.
[86] Z. Jamal, A. Martin, A. G6mez-Mufioz and D.N. Brindley (1991) J. Biol. Chem. 266, 2988-2996.
[87] H. Kanoh. K. Yamada and F. Sakane (1990) TIBS 15.
47 50.
[88] F. Sakane, K. Yamada, H. Kanoh, C. Yokoyama and T.
Tanabe (1990) Nature 344, 345 348.
[89] K. Goto, M. Funayama and H. Kando (1994) Proc. Natl.
Acad. Sci. USA 91, 13042 13046.
R.B, Cornell, R,S. Arnold / Chemist O, and Physics ~! Lipids 81 (1996) 215- 227
[90] K. Goto and H. Kondo (1993) Proc. Natl. Acad. Sci.
USA 90, 7598-7602.
[91] C.R. Loomis, J.P. Walsh and R.M. Bell (1995) J. Biol.
Chem. 260, 4091-4097.
[92] F. Sakane, K. Yamada, S.-i. Imai and H. Kanoh (1991) J.
Biol, Chem. 266, 7096 7100.
[93] J,M. Besterman, R.S, Pollenz, E.L. Booker, Jr and P.
Cuatrecasas {1986) Proc. Natl. Acad. Sci. USA 83, 9378
9382.
227
[94] J.P. Walsh and R.M. Bell (19861 J. Biol. Chem. 261,
15(/62 15069.
[95] W. Dowhan, W.T. Wickner and E.P. Kennedy (19741 J.
Biol. Chem. 249, 3079.
[96] N.D. Ridgway and D.E. Vance (19871 J. Biol. Chem. 262,
17231- 17239.
[97] R.H. Hjelmstad and R.M. Bell (19881 ]. Biol. Chem. 263,
19748 19757.
[98] R.H. Hjelmstad and R.M. Bell t1991t J. Biol. Chem. 266,
5094 511)3.