From the Laboratory of Cellular and Developmental Biology, NIDDK, National Institutes of Health, Bethesda, Maryland 20892-8028
Received for publication, October 28, 2002, and in revised form, December 6, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Perilipin A coats the lipid storage droplets in
adipocytes and is polyphosphorylated by protein kinase A (PKA); the
fact that PKA activates lipolysis in adipocytes suggests a role for
perilipins in this process. To assess whether perilipins participate
directly in PKA-mediated lipolysis, we have expressed constructs coding for native and mutated forms of the two major splice variants of the
perilipin gene, perilipins A and B, in Chinese hamster ovary
fibroblasts. Perilipins localize to lipid droplet surfaces and displace
the adipose differentiation-related protein that normally coats the
droplets in these cells. Perilipin A inhibits triacylglycerol
hydrolysis by 87% when PKA is quiescent, but activation of PKA and
phosphorylation of perilipin A engenders a 7-fold lipolytic activation.
Mutation of PKA sites within the N-terminal region of perilipin
abrogates the PKA-mediated lipolytic response. In contrast, perilipin B
exerts only minimal protection against lipolysis and is unresponsive to
PKA activation. Since Chinese hamster ovary cells contain no
PKA-activated lipase, we conclude that the expression of perilipin A
alone is sufficient to confer PKA-mediated lipolysis in these cells.
Moreover, the data indicate that the unique C-terminal portion of
perilipin A is responsible for its protection against lipolysis and
that phosphorylation at the N-terminal PKA sites attenuates this
protective effect.
Acute mobilization of adipose triacylglyerol stores for energy is
regulated primarily by the activation state of
cAMP-dependent protein kinase
(PKA)1 (1). Historically,
this stimulation has been attributed to phosphorylation and activation
of hormone-sensitive lipase (HSL), but as noted previously (2, 3), the
meager doubling of HSL activity upon phosphorylation in
vitro falls far short of explaining the 30-100-fold activation of
cellular lipolysis upon elevation of PKA activity in isolated primary
adipocytes. Although some the of differences between the magnitude of
the in vitro and in vivo responses may be
attributed to the PKA-induced translocation of HSL from the cytosol to
the lipid storage droplets within adipocytes (4), it is likely that
additional factors, notably the perilipins, contribute to the cellular response.
The perilipins are a class of proteins found exclusively at the
limiting surface of lipid storage droplets, i.e. at the
lipid/aqueous interface, in adipocytes and in steroidogenic cells
(5-7). These proteins are the most abundant PKA substrates in
adipocytes (5), and both their subcellular location and
polyphosphorylation by PKA suggest a role for the perilipins in
PKA-mediated lipolysis. In adipocytes, alternative mRNA splicing
gives rise to perilipins A and B, the former at much higher levels than
the latter (8). Thus far, functional studies point to a role for the
perilipins in protecting stored TAG from hydrolysis by cellular
lipases. Ectopic expression of perilipin A in 3T3-L1 adipoblasts
results in perilipin A-coated lipid droplets and increases the
half-life of stored TAG deposits by a factor of 4.5 (9). Also,
treatment of 3T3-L1 adipocytes with tumor necrosis factor- None of the above studies address a role for phosphorylation of
perilipin in the lipolytic response. To this end, using CHO cells, we
have examined the functional consequences of introducing perilipin A,
perilipin B, and mutant variants of these species. We report herein
that the expression of perilipin A alone is sufficient to inhibit
lipolysis by 87% and that upon activation of PKA, lipolysis is
stimulated by 7-fold. These results re-create the strong regulatory role for perilipin seen in adipocytes and contrast with results from
another system that detected only a modest, ~50% stimulation by PKA
upon introduction of perilipin A into cells (13). We further show that
such regulation is abrogated upon mutation of the PKA sites that lie
within the N-terminal region of perilipin, as did Souza et
al. (13). In contrast, we report that neither native nor mutated
perilipin B provide protection.
Expression of Perilipins in CHO Cells--
Perilipin functional
studies were conducted with CHO K-1 cells (American Type Culture
Collection, Manassas, VA) into which we introduced murine perilipins A
and B and mutated forms of these proteins in which serine residues
within consensus PKA sites were mutated to alanines. Mutations were
made in sets of three, by grouping the three N-terminal PKA sites
(amino acids 81, 222, and 276). The accession number for murine
perilipin A cDNA is GenBankTM AY161165. The
terminologies for the various constructs are shown in Fig
1. Each mutation was made by PCR using
primers containing the mutation of interest and external primers to
amplify a small cassette, which was then cloned in-frame in mouse
perilipin A cDNA (14). The accuracy of each mutation was confirmed
by sequencing. Both unmodified and mutant full-length cDNAs were
cloned in the retroviral expression vector, pSR MSV-Tk Neo. Retrovirus
was produced by co-transfecting retroviral expression vectors and the
packaging vector, pSV Lipid Loading and Hydrolysis--
Cells infected with
retrovirus-containing perilipin inserts or control cells, which stably
incorporated the retrovirus vector lacking perilipin cDNA,
were plated at a density of 0.5 × 106 cells/35-mm
(diameter) well. After cells became adherent, the medium was changed to
F-12 (Invitrogen) supplemented with 400 µM oleic acid
complexed to 0.4% bovine serum albumin to promote triacylglycerol
deposition. [3H]Oleic acid, at 1 × 106
dpm/well, was included as a tracer. Following a 24-h loading period,
the cells were washed with 4% bovine serum albumin in PBS
to remove unincorporated oleic acid. These conditions were employed for
all experiments in the present studies, and the amount of protein per
well did not differ among cells bearing the different perilipin
species; thus, data are expressed on a per well basis. For lipid
analysis, the cells were extracted with chloroform:methanol (2:1) (18),
and lipids were identified by thin layer chromatography on silica gel
plates (Analtech, Newark, DE). Mobile phase 1 was acetone developed for
4 cm; mobile phase 2 was 80:20:1 petroleum ether:diethyl ether:acetic
acid developed to the top of the plate (19). To measure hydrolysis of
cellular lipids, cells were loaded with radiolabeled oleic acid as
described above and placed in an efflux medium, which was F-12 medium
including 1% defatted bovine serum albumin (ICN Biomedical, Aurora,
OH) as a fatty acid acceptor. The efflux of radioactivity to the medium
was measured over time by scintillation counting. TLC confirmed that
all radioactivity released to the medium was free
[3H]oleic acid. Re-esterification of fatty acids was
prevented by the inclusion of 2.5 µM Triacsin C (Biomol,
Plymouth Meeting, PA), an inhibitor of acyl co-enzyme A synthetase, to
the medium. In preliminary experiments, this level of Triacsin C was
found to effectively block all fatty acid esterification in CHO cells. To stimulate protein kinase A activity, cAMP levels in cells were elevated by the addition of 1 mM isobutylmethylxanthine
(IBMX) and 10 µM forskolin. The efficacy of this
stimulation method was confirmed by immunoblotting, which revealed that
upon stimulation of cells that were expressing perilipin A, the protein
migrated more slowly under SDS-PAGE, characteristic of
PKA-phosphorylated perilipin A (Fig.
2).
As we reported previously (20), lipid droplets in CHO cells
are coated with ADRP (Figs. 2 and 3); this protein and the perilipins share significant sequence homology over their N-terminal
domains (7). Upon expression of
constructs that encode either native or mutated perilipin A, as well as
native or mutated perilipin B, the droplets acquired a coating of
perilipin and the ADRP diminished, as evidenced both by immunostaining
(Fig. 3) and immunoblotting (Figs. 2 and
4). This change in the lipid droplet coat
protein is a reprise of the perilipin-for-ADRP switch that occurs
during differentiation of 3T3-L1 adipocytes (20). Moreover, the amounts of unmodified and mutated perilipins A and B were highly similar. The
only cell lines carried forth to the metabolic studies below were those
in which the perilipin (A or B) expression was sufficient to eliminate
all ADRP expression, as judged both by immunofluorescence and by
immunoblotting. Upon repeated passage of some perilipin A-expressing
cell lines, the perilipin expression waned, and the metabolic phenotype
reverted to that similar to the control cells with ADRP-coated
droplets. In general, a reversal of the perilipin A phenotype occurred
when the perilipin expression declined by ~50%.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
both
reduces expression of perilipin A and increases basal lipolysis. With the use of an adenovirus vector to maintain perilipin expression during
tumor necrosis factor-
treatment, Souza et al. (10) found
that expression of either perilipin A or B was sufficient to maintain a
coating of perilipin on the droplets and to suppress tumor necrosis
factor-
-activated lipolysis. Such findings further support the
conclusion that perilipins can protect the intracellular TAG from
hydrolysis by lipases. Finally, two reports describing the phenotype of
the perilipin null (11, 12) mouse show that these animals have greatly
reduced adipose stores and constitutively high levels of basal
lipolysis in their isolated adipose cells, again consistent with the
loss of the protective effect of perilipin. In one of these reports
(12), it was shown that in the absence of perilipin, the adipose cells
droplets were coated with adipose differentiation-related protein
(ADRP), indicating that this related protein did not substitute for
perilipin in providing a protective barrier against lipolysis in the
absence of PKA activation.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
E-MLV (15), into HEK-293 T fibroblasts using
calcium phosphate. Cells were transfected for 9 h in the presence
of chloramphenicol, following which the medium was changed to fresh
Dulbecco's modified Eagle's medium, and viral production was
continued for 48 h. Prior to infection, CHO-K1 fibroblasts were
treated overnight with tunicamycin (16, 17) to increase the infection
rate. Cells were infected for 24 h using medium containing
4 µg/ml polybrene harvested from the 293 T cells. Immediately
following infection, CHO cells were placed under selection in 600 µg/ml G418.
View larger version (29K):
[in a new window]
Fig. 1.
The various constructs
produced for this study are depicted. Peri A,
full-length perilipin A; 3XN Peri A, perilipin A with
serines in three N-terminal PKA sites at amino acids 81, 222, and 276 mutated to alanines; Peri B, full-length Peri B;
3XN Peri B, perilipin B with Ser-to-Ala mutations at
residues within PKA sites at amino acids 81, 222, and 276.
View larger version (35K):
[in a new window]
Fig. 2.
Shift in perilipin A migration under SDS-PAGE
following stimulation with IBMX and forskolin. Cells expressing
perilipin A and vector control (VC) cells were either
unstimulated (lane 1 ) or stimulated (lane 1+)
with 1 mM IBMX and 10 µM forskolin for 10 min. Homogenates were subjected to SDS-PAGE and immunoblotted with
anti-perilipin and anti-ADRP antisera. The characteristic upward shift
of PKA-phosphorylated perilipin (PA) in stimulated
cells is evident.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (48K):
[in a new window]
Fig. 3.
Immunofluorescence of different cell types
with anti-perilipin and anti-ADRP. Cells were infected with
retroviral constructs bearing the indicated forms of perilipin.
Abbreviations are described in the legend for Fig. 1 except for
BF, which indicates bright field. Control
indicates the empty vector with no perilipin insert. Cells were plated,
grown, and loaded with lipid, fixed, and immunostained for perilipin
and ADRP as described under "Experimental Procedures."
View larger version (17K):
[in a new window]
Fig. 4.
Immunoblot analysis of different types of
cells. Cells from Fig. 3 were homogenized, subjected to SDS-PAGE,
and stained for perilipin and ADRP. Abbreviations are as described in
legend for Fig 1. VC, vector control; PA,
perilipin A; PB, perilipin B.
To assess lipolytic activity, CHO cells were loaded with
[3H]oleic acids as described previously (9), after which
the efflux of [3H]oleic acid to the medium was monitored
for 3 h. Triacsin C was included in all incubations during the
efflux phase to prevent reutilization of fatty acids released from
hydrolyzed TAG (21-23). In control cells (with ADRP-coated droplets),
nearly 30% of the stored [3H]oleic acid was released to
the medium over 3 h, and PKA activation inhibited fatty acid
efflux by about 30%. In contrast, in cells expressing perilipin A, in
the absence of PKA activation, the release of [3H]oleic
acid was suppressed by 87% when compared with that in control cells
with ADRP-coated lipid droplets. Upon activation of PKA with the
combination of forskolin and IBMX, lipolysis was activated by 7-fold
(range of 3-13-fold) over the unstimulated cells. Iin Fig.
5 (Perilipin A panel), note
that the stimulatory effects of PKA activation on oleic acid
release in Peri A cells occurred only after a lag of ~30 min.
During this 30-min period, there were no differences in the rates of
oleic acid release in unstimulated and stimulated cells; thereafter,
lipolysis under the two conditions diverged and remained nearly linear
over the next 2.5 h. There was another highly reproducible break
in the curve at 120 min, indicating that the perilipin-mediated
increase in lipolysis is a slowly evolving process. The lipolytic rates for all constructs tested over the 2.5-h period are depicted in Table
I. The 30-min lag was evident in all
cells infected with perilipin species that were responsive to PKA
activation. The paradoxical inhibition of oleic acid release
upon PKA activation of control cells occurred without a detectable lag,
as shown in Fig. 4 (Vector Control Panel). Perilipin A from
mouse contains six consensus PKA phosphorylation sites (Fig. 1). We
also tested a mutant variant of perilipin A in which the three most
N-terminal serines within PKA sites were mutated to alanines (Peri
3XN). Introduction of this form of perilipin A also prevented the
release of radiolabeled fatty acid stores in the absence of PKA
activation, but unlike native perilipin A, the mutant form was
relatively unresponsive to PKA activation. In contrast to the 7-fold
increase with the perilipin A cells, the activation of the mutated
perilipin was merely 1.7-fold (Table I). In contrast, perilipin B, the shorter splice variant of the perilipin gene failed to shield the TAG
from hydrolysis and the native perilipin B, inhibiting activity by only
13%, when compared with the 90% inhibition by perilipin A. Also,
perilipin B cells were unresponsive to PKA activation. Oddly, the
mutated form of perilipin B (Peri B 3X) was more protective than native
perilipin B, inhibiting lipolysis by 63% but unresponsive to PKA
activation (Table I).
|
|
Under the loading conditions used in these studies, the great majority
of the fatty acids taken up by the cells (>75%) were stored as TAG,
and the distribution of tritiated oleic acid among the phospholipid,
diacylglycerol, cholesteryl ester, and fatty acid pools was independent
of the nature of coating on the lipid droplets (Fig.
6). However, the magnitude of uptake and
storage of [3H]oleic acid in the TAG pool was dependent
on which protein coated the droplets. Cells expressing perilipin A
accumulated the greatest amount, 148.9 ± 19.4 nmol oleic acid,
greater than the perilipin B-expressing cells, 125.6 ± 3 nmol
oleic acid, which, in turn, was greater than control cells with
ADRP-coated droplets, 103.0 ± 10.3 nmol oleic acid (mean ± S.E., n = 6). These accumulations represent the
steady state balance between lipogenesis and lipolysis during the lipid
loading phase; accordingly, these levels reflect the differences in
unstimulated lipolysis since the cells were loaded under the
non-stimulated condition, i.e. no IBMX or forskolin. After
the 3-h efflux incubation, the non-TAG pools lost no measurable [3H]oleic acid (Fig. 6), whereas the TAG pools exhibited
substantial losses, which again was dependent on the lipid droplet coat
protein and the PKA activation state (Fig.
7). Moreover, TLC analysis revealed that
all of the radiolabel that appeared in the medium was in the form of
[3H]oleic acid, which could be accounted for entirely by
the loss of radiolabeled fatty acid from the intracellular TAG
pool.
|
|
The percentages of TAG hydrolyzed over 3 h in control, Peri
A, and Peri A 3XN cells are summarized in Fig. 7. It is clear that the
TAG pool, presumably within the Peri A-coated pool, is the source of
the accelerated oleic acid release under PKA-stimulation.
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In the present study, we have demonstrated that expression of perilipin in CHO fibroblasts results in perilipin-coated lipid droplets from which the ADRP has disappeared. Perilipin A, but not perilipin B, suppresses lipolysis of droplet TAG, and PKA phosphorylation of perilipin A engenders a robust lipolytic reaction despite the absence of HSL or any other PKA-mediated lipase. Unlike PKA-mediated lipolysis in adipocytes, the perilipin A-mediated lipolysis occurs only after a lag of ~30 min. Lipolysis is not stimulated when the serine residues within the three PKA sites at the N-terminal region of perilipin A are mutated to alanines.
The finding that the proteinaceous coating of the lipid droplets may confer PKA-mediated regulation of TAG hydrolysis prompts a reconsideration of the conventional view of PKA-mediated lipolysis in adipocytes. Heretofore, such activation was attributed primarily to the phosphorylation of HSL, following which the lipase was activated, but only minimally, and translocated from the cytosol to the lipid droplet surface. Clearly, given the above findings, one must account for the contribution of PKA phosphorylation of perilipin to the regulation of lipolysis. Interestingly, the HSL null mouse retains a modicum of isoproterenol-stimulated lipolytic activity, which led to the suggestion that adipocytes may contain a second hormone-sensitive lipase (24). Given the present results, plus the likelihood that the HSL null mouse expresses perilipins in its adipocytes, the presence of these lipid droplet-coating proteins may explain the residual PKA-activated lipolysis in the HSL null mouse (25).
Despite the earlier report suggesting that HSL may be expressed in CHO cells (26), we detected no HSL by immunoblotting the these cells nor could we detect any PKA-stimulated lipolytic activity in CHO cell homogenates (data not shown). Thus, the neutral lipid lipase responsible for the TAG hydrolysis in the CHO cells appears unresponsive to PKA, and it is concluded that PKA regulation of lipolysis in CHO cells expressing perilipin A results solely from the PKA phosphorylation of the perilipin. Clearly, the ability of the endogenous CHO cell lipase(s) to hydrolyze stored TAG under the PKA-stimulated condition is dependent on the phosphorylation of perilipin A. This conclusion is supported by the finding that mutation of selected PKA sites eliminated the regulated response. In a separate study, we show that perilipin A acts cooperatively with HSL in lipolysis when both proteins are introduced into CHO cells.2 Thus, the ability of perilipin to regulate lipolysis of the TAG within lipid droplets appears to be manifested with any lipase that has access to the lipid droplet surface. The enhancement of lipolytic activity by perilipin A and the endogenous lipase of CHO cells is clearly a time-dependent process that requires at least 30 min after stimulation to become manifest. In contrast, when both HSL and perilipin are introduced into CHO cells, the lipolytic activation is nearly immediate with no detectable lag.2 We assume that the phosphorylation of HSL fosters this immediate reaction in contrast to the endogenous lipase, which is apparently not a PKA substrate. Oddly, perilipin B neither protected against lipolysis nor conferred significant PKA regulation of lipolysis, which was not anticipated given the results of Souza et al. (10). However, in the present study, we expressed perilipin B without perilipin A, whereas in the studies of Souza et al. (10), perilipin B was introduced into cells that contained abundant perilipin A.
The fatty acids that appeared in the CHO cell medium in the present study derived from the triolein stored in the intracellular neutral lipid droplets, which were coated with either ADRP or the perilipins introduced into the cells. No other lipid pool contributed to the fatty acid that was released. Moreover, stimulation of cells expressing perilipin A led to a loss solely from the triolein from within the perilipin-coated droplets. Thus, one may assume that the PKA-mediated increase in fatty acid release stems from changes in the lipid droplet surface upon phosphorylation of perilipin A, a conclusion supported by the finding that the 3XN mutant species failed to respond to PKA activation.
The present data permit speculation on the structural determinants for perilipin function at the lipid droplet surface. If perilipins A and B assume similar extended conformations at the droplet surface, the greater protection against hydrolysis of the longer splice variant, perilipin A, may be attributed to its greater length and thus greater coverage of droplet surface, i.e. it is the extended C-terminal tail (~12 kDa) of perilipin A that is important in protecting against lipolysis in the unstimulated state. It follows that phosphorylation of PKA sites in the N-terminal region induces a conformational change in the C-terminal region that exposes portions of the lipid droplet surface to lipase action under conditions of PKA activation.
In addition to the present studies on CHO cells, it is reasonable to
assume that phosphorylation of perilipin contributes to the lipolytic
activation in adipocytes. This conclusion is reinforced by the finding
that lipolytic stimulation by -adrenergic agonists of adipocytes
from perilipin null mice is strongly blunted despite the presence of
normal amounts of HSL (12).
After completion of the present studies, a similar study appeared that largely agrees with a number of the findings reported herein (13), but that study did not include studies with perilipin B, and mutational analysis was limited to the construct we term Peri A 3XN (13). Although the present studies and those of Souza et al. (13) agree qualitatively, there are large quantitative differences between the two studies. For example, we find that introduction of perilipin A suppresses lipolysis by ~90% when compared with control cells, whereas Souza et al. (13) observed a suppression of about 50%. Also, we find that activation of Peri A cells leads to 162% greater lipolysis than in control cells, as compared with a 50% increment in the other study. Further, PKA activation of perilipin A cells produced a 700% increase in lipolysis, as compared with a 160% increase in the published study. Our findings suggest that PKA phosphorylation of perilipin A does far more than merely abrogate the protective effect of the perilipin, as suggested by Souza et al. (13), but additionally, exposes considerably more TAG to lipase activity than is exposed in the control cells containing droplets coated with ADRP. Reasons for these differences may include the use by Souza et al. (13) of NIH 3T3 fibroblasts engineered to be more lipogenic by using cells that also expressed an ectopic fatty acid transporter and an ectopic acyl synthase, whereas in the present studies, we used an acyl synthase inhibitor to block re-esterification of fatty acids. Also, we used CHO cells that were stably transfected with the various perilipin constructs, whereas Souza et al. (13) infected the cells with perilipin adenovirus. In separate studies, we, too, used adenovirus to introduce perilipin into cells and find modest effects with CHO cells similar to those reported by Souza et al. (13) for NIH 3T3 cells.2
In parallel studies, we demonstrate both that HSL translocation to
lipid droplets requires that such droplets be coated with PKA-phosphorylatable perilipin A2 and that HSL be
phosphorylated at one of its C-terminal PKA sites, Ser-659
or 660.3 Thus, the
physiological activation lipolysis in adipocytes is a concerted
reaction between phospho-HSL and phospho-perilipin.
![]() |
FOOTNOTES |
---|
* 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.
Present address: Dept. of Chemistry, Otterbein College,
Westerville, OH 43081-2006.
§ Present address: Dept. of Nutritional Sciences, Rutgers, The State University of New Jersey, New Brunswick, NJ 08901.
¶ To whom correspondence should be addressed: Bldg. 50, Rm. 3140, National Institutes of Health, Bethesda, MD 20892-8028. Tel.: 301-496-6991; Fax: 301-496-5239; E-mail: DeanL@intra.niddk.nih.gov.
Published, JBC Papers in Press, December 10, 2002, DOI 10.1074/jbc.M211005200
2 C. Sztalryd, G. D. H. Xu, A. R. Kimmel, and C. Londos, submitted for publication.
3 C.-L. Su, C. Sztalryd, J. A. Contreras, C. Holm, A. R. Kimmel, and C. Londos, submitted for publication.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: PKA, cAMP-dependent protein kinase; Peri, perilipin; Peri A, perilipin A; Peri B, perilipin B; ADRP, adipose differentiation-related protein; IBMX, isobutylmethylxanthine; TAG, triacylglycerol; HSL, hormone-sensitive lipase; E-MLV, ecotropic murine leukemia virus.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. |
Honnor, R. C.,
Dhillon, G. S.,
and Londos, C.
(1985)
J. Biol. Chem.
260,
15130-15138 |
2. |
Londos, C.,
Brasaemle, D. L.,
Schultz, C. J.,
Adler-Wailes, D. C.,
Levin, D. M.,
Kimmel, A. R.,
and Rondinone, C. M.
(1999)
Ann. N. Y. Acad. Sci.
892,
155-168 |
3. | Holm, C., Osterlund, T., Laurell, H., and Contreras, J. A. (2000) Annu. Rev. Nutr. 20, 365-393[CrossRef][Medline] [Order article via Infotrieve] |
4. | Brasaemle, D. L., Levin, D. M., Adler-Wailes, D. C., and Londos, C. (1999) Biochim. Biophys. Acta 1483, 251-262 |
5. |
Greenberg, A. S.,
Egan, J. J.,
Wek, S. A.,
Garty, N. B.,
Blanchette-Mackie, E. J.,
and Londos, C.
(1991)
J. Biol. Chem.
266,
11341-11346 |
6. |
Servetnick, D. A.,
Brasaemle, D. L.,
Gruia-Gray, J.,
Kimmel, A. R.,
Wolff, J.,
and Londos, C.
(1995)
J. Biol. Chem.
270,
16970-16973 |
7. | Londos, C., Brasaemle, D. L., Schultz, C. J., Segrest, J. P., and Kimmel, A. R. (1999) Semin. Cell Dev. Biol. 10, 51-58[CrossRef][Medline] [Order article via Infotrieve] |
8. | Greenberg, A. S., Egan, J. J., Wek, S. A., Moos, M. C., Jr., Londos, C., and Kimmel, A. R. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 12035-12039[Abstract] |
9. |
Brasaemle, D. L.,
Rubin, B.,
Harten, I. A.,
Gruia-Gray, J.,
Kimmel, A. R.,
and Londos, C.
(2000)
J. Biol. Chem.
275,
38486-38493 |
10. |
Souza, S. C.,
de Vargas, L. M.,
Yamamoto, M. T.,
Lien, P.,
Franciosa, M. D.,
Moss, L. G.,
and Greenberg, A. S.
(1998)
J. Biol. Chem.
273,
24665-24669 |
11. | Martinez-Botas, J., Anderson, J. B., Tessier, D., Lapillone, A., Chang, B. H. J., Quast, M. J., Forenstein, D., Chen, K. H., and Chan, L. (2000) Nat. Genet. 26, 474-479[CrossRef][Medline] [Order article via Infotrieve] |
12. |
Tansey, J. T.,
Sztalryd, C.,
Gruia-Gray, J.,
Roush, D. L.,
Zee, J. V.,
Gavrilova, O.,
Reitman, M. L.,
Deng, C. X.,
Li, C.,
Kimmel, A. R.,
and Londos, C.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
6494-6499 |
13. |
Souza, S. C.,
Muliro, K. V.,
Liscum, L.,
Lien, P.,
Yamamoto, M. T.,
Schaffer, J. E.,
Dallal, G. E.,
Wang, X.,
Kraemer, F. B.,
Obin, M.,
and Greenberg, A. S.
(2002)
J. Biol. Chem.
277,
8267-8272 |
14. | Saiki, R. K., Gelfand, D. H., Stoffel, S., Scharf, S. J., Higuchi, R., Horn, G. T., Mullis, K. B., and Erlich, H. A. (1988) Science 239, 487-491[Medline] [Order article via Infotrieve] |
15. | Muller, A. J., Young, J. C., Pendergast, A. M., Pondel, M., Landau, N. R., Littman, D. R., and Witte, O. N. (1991) Mol. Cell. Biol. 11, 1785-1792[Medline] [Order article via Infotrieve] |
16. | Wilson, C. A., and Eiden, M. V. (1991) J. Virol. 65, 5975-5982[Medline] [Order article via Infotrieve] |
17. | Miller, D. G., and Miller, A. D. (1992) J. Virol. 66, 78-84[Abstract] |
18. | Bligh, E. G., and Dyer, W. J. (1959) Can. J. Biochem. Physiol. 37, 911-917 |
19. | Levacher, C., Sztalryd, C., and Picon, L. (1985) Reprod. Nutr. Dev. 25, 169-173[Medline] [Order article via Infotrieve] |
20. | Brasaemle, D. L., Barber, T., Wolins, N. E., Serrero, G., Blanchette-Mackie, E. J., and Londos, C. (1997) J. Lipid Res. 38, 2249-2263[Abstract] |
21. | Igal, R. A., Wang, P., and Coleman, R. A. (1997) Biochem. J. 324, 529-534[Medline] [Order article via Infotrieve] |
22. |
Tomoda, H.,
Igarashi, K.,
Cyong, J. C.,
and Omura, S.
(1991)
J. Biol. Chem.
266,
4214-4219 |
23. | Tomoda, H., Igarashi, K., and Omura, S. (1987) Biochim. Biophys. Acta 921, 595-598[Medline] [Order article via Infotrieve] |
24. |
Slatiel, A. R.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
97,
535-537 |
25. |
Osuga, J.,
Ishibashi, S.,
Oka, T.,
Yagyu, H.,
Tozawa, R.,
Fujimoto, A.,
Shionoiri, F.,
Yahagi, N.,
Kraemer, F. B.,
Tsutsumi, O.,
and Yamada, N.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
787-792 |
26. | Osuga, J., Ishibashi, S., Shimano, H., Inaba, T., Kawamura, M., Yazaki, Y., and Yamada, N. (1997) Biochem. Biophys. Res. Commun. 233, 655-657[CrossRef][Medline] [Order article via Infotrieve] |
27. | Student, R. (1908) Biometrika 6, 1-25 |