Protein acyl thioesterases (Review)
Running title: Protein acyl thioesterases
Ruth Zeidman, Caroline S. Jackson and Anthony I. Magee
Address:
Molecular Medicine
National Heart & Lung Institute
Sir Alexander Fleming Building
South Kensington
Imperial College London
London SW7 2AZ
UK
Corresponding author: Anthony I. Magee
E-mail: t.magee@imperial.ac.uk
Telephone: +44 20 7594 3135
Keywords
APT1
Palmitoylation
Protein acyl thioesterase
Protein acyl transferase
S-acylation
1
Abstract
Many proteins are S-acylated, affecting their localization and function. Dynamic Sacylation in response to various stimuli has been seen for several proteins in vivo. The
regulation of S-acylation is beginning to be elucidated. Proteins can autoacylate or be Sacylated by protein acyl transferases (PATs). Deacylation, on the other hand, is an
enzymatic process catalyzed by protein thioesterases (APT1 and PPT1) but only APT1
appears to be involved in the regulation of the reversible S-acylation of cytoplasmic
proteins seen in vivo. PPT1, on the other hand, is involved in the lysosomal degradation
of S-acylated proteins and PPT1 deficiency causes the disease infant neuronal ceroid
lipofuscinosis.
S-acylation
Proteins are modified in many ways as a means to control their localisation and
function. Many proteins that exert their function at or near a cellular membrane are
modified by the covalent attachment of lipids. S-acylation is the post-translational
attachment of fatty acids to proteins through a thioester bond, usually on a cysteine
residue. Palmitic acid, a 16-carbon saturated fatty acid, is the most common fatty acid to
be linked to proteins in this manner, explaining why S-acylation is often referred to as
palmitoylation, although proteins can also be modified in this manner by many other
fatty acids, both saturated and unsaturated, including myristic, oleic, arachidonic and
stearic acids.
A wide variety of proteins are acylated, including transmembrane proteins like Gprotein coupled receptors, T cell co-receptors CD4 and CD8 and ion channels, cytosolic
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and membrane-associated proteins like members of the Ras and Src families,
heterotrimeric G-proteins, endothelial nitric oxide synthase (eNOS), SNARE vesicle
fusion proteins, as well as secreted and viral proteins. Addition of fatty acids to a
protein has various effects on signalling, trafficking, protein stability, protein-protein
interaction and membrane and membrane subdomain association [1-3].
In the cases where the acylation takes place on an N-terminal cysteine, the fatty acid is
often found linked to the protein through an amide bond to the N-terminus. This type of
acylation is referred to as N-acylation and often occurs in secreted proteins, for example
Hedgehog/Sonic hedgehog [4], but also in Gαs [5]. This may well occur in two steps; Sacylation via a thioester bond between the fatty acid and the sulphur in the cysteine,
followed by an internal S to N acyl shift to the α-amino group.
S-acylation is a reversible modification
One interesting difference between S-acylation and N-acylation is the reversibility of
the S-acylation. In fact, S-acylation is the only lipid modification of proteins that is
readily reversible, seen as the half time of the attachment of the fatty acid side chain
being significantly shorter than the life time of the protein. Many proteins have been
shown to be dynamically S-acylated in vivo with different turnover rates. For instance,
S-acylation of Fyn, a member of the Src kinase family, is required for its plasma
membrane localisation and has been shown to be reversible, with a half life of 1.5-2 h
[6]. The unpalmitoylated pool of Fyn was found in intracellular membranes. Another
Src family kinase, Lck, has also been demonstrated to be reversibly palmitoylated in
pervanadate-stimulated human T blasts, with a half life of 15-30 minutes (Fig. 1B;
unpublished work, C.S. Jackson and A.I. Magee). Ras proteins were one of the first
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proteins to be reported to have dynamic S-acylation [7] and different H-Ras variants are
also deacylated at variable rates. Oncogenic H-Ras mutants, which are GTP-bound to a
higher proportion than wild-type H-Ras, were determined to have shorter palmitate half
lives than the wild type H-Ras, even though the steady state levels of palmitoylation did
not differ [8].
(Insert Fig 1 near here)
For some proteins, the increased acyl turnover seen on many proteins in vivo is in
response to various stimuli. One such example is the acylation turnover of the neuronal
protein PSD-95, which is increased upon neuronal activity and causes a clustering of
PSD-95 at the synapse [9]. eNOS deacylation is more efficient in the presence of Ca2+calmodulin, an eNOS activator [10]. Many G-protein coupled receptors - including the
β2-adrenergic, α2A-adrenergic, m2 muscarinic acetylcholine, 5-HT4a serotonin, V1a
vasopressin and δ opioid receptors - are also dynamically acylated after agonist binding
[11-16]. The effects of increased fatty acyl turnover on the receptors are not clear. The
steady state levels of acylation will depend on the relative reacylation and deacylation
rates. Even if the steady state levels are unchanged, under conditions with of fatty acyl
turnover there will be a proportion of transiently deacylated receptors. The relative size
of this pool and the time the receptors stay deacylated might be a mechanism for
controlling signalling through the receptors. In fact, receptor desensitization by
phosphorylation is increased in depalmitoylated receptors compared to their
palmitoylated counterparts [17-19].
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The turnover of palmitate on Gα subunits is also regulated by stimulation of their
associated receptors [20-24] as activation of the receptor leads to the dissociation of the
α subunit from the βγ subunits, making the cysteine on the α subunit available for
depalmitoylation by the thioesterase APT1 [25].
Autoacylation
Even though proteins have been known to be modified by fatty acids and the effects of
acylation on localization and signalling have been known for many years, the
mechanism behind the actual process of attaching the acyl chains has not been very
clear. Many proteins and peptides can autoacylate when incubated with the appropriate
acyl-CoA in vitro. Myristoylated Giα1 is autopalmitoylated in the presence of palmitoylCoA at the same cysteine resdidue that is acylated in vivo [26]. Similarly, the
autoacylation of other proteins have been reported, including rhodopsin [27], SNAP-25
[28], carbamoyl-phosphate synthetase 1 [29], Bet3 [30], β2-adrenergic receptor [31] and
c-Yes [32]. Autoacylation does not seem to be a universal mechanism as, for example,
GAP-43 and Fyn do not autoacylate under the same conditions as Giα1 does [26]. In
addition, not all of these in vitro reactions take place at physiological pH and/or acylCoA concentrations. In some cases the reaction times are also too long to accommodate
the fast palmitate cycling seen on many proteins in vivo.
DHHC PATs
It seems more likely that, at least for most proteins, S-acylation is an enzymatic process.
In recent years there has been much progress in identifying protein-acyl transferases
(PATs). Two families of these enzymes are the subjects of other reviews in this issue.
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Briefly, one such group of PATs is the DHHC family, consisting of multi-spanning
transmembrane proteins that have a cysteine-rich domain containing a conserved
asparatate-histidine-histidine-cysteine (DHHC) motif, with is essential for the PAT
activity [33, 34]. The PAT activity of the DHHC proteins was originally discovered in
yeast, where the DHHC protein Erf2, together with Erf4, mediates the acylation of yeast
Ras [33]; another DHHC protein, Akr1, is a PAT for yeast casein kinase 2 [34]. The
palmitoylation of specific proteins could not be detected in yeast strains deficient in
different combinations of the yeast DHHC proteins, confirming the in vivo acyltransferase activity of the DHHC proteins [35]. In a screen on potential substrates, the
23 DHHC proteins predicted in the mammalian genome had variable substrate
specificities, with some overlap [36]. The DHHC PAT substrates are intracellular
proteins [1, 37, 38] including: H-and N-Ras which are S-acylated by DHHC9 together
with a Golgi-associated protein GCP16, echoing the yeast Erf2/Erf4 complex [39]; the
DHHC21 substrate eNOS [40]; huntingtin which is acylated by HIP14/DHHC17 [41,
42]; the γ2 subunit of the GABAA receptor which is acylated by GODZ/DHHC3 [43]
and the neuronal PSD-95 protein, which in slightly contradictory reports has been
suggested to be the substrate of DHHC2, 3, 7 and 15 (but not DHHC17) [36] or of
HIP14/DHHC17, in addition to other neuronal substrates [41].
MBOAT PATs
A different group of PATs appear to have secreted proteins and peptides as their
substrates. Hedgehog (Hh) proteins e.g. Sonic hedgehog (Shh), Spitz, Wnts and ghrelin
require acylation for their function and are acylated by members of the MBOAT
(membrane-bound-O-acyltransferase) family [44]. A common feature of this group of
proteins is that they have several membrane-spanning domains and that they transfer
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organic acids, typically fatty acids, onto hydroxyl groups of membrane-embedded
targets [45]. Only a small subset of members is known to transfer fatty acids and other
lipids to proteins. Rasp/Skinny hedgehog/Hedgehog acyl transferase (Hhat) is the PAT
for Hh/Shh [46] and Spitz [47], Porcupine (Porc) is the PAT for Wnts [48] and GOAT
is the PAT for ghrelin [49]. The acylation of secreted proteins is reviewed elsewhere in
this issue.
Ykt6-type PATs
A third mechanism for fatty acid attachment to proteins incorporates elements of both
autoacylation and enzymatic acyl transfer. The yeast SNARE protein Ykt6 binds
palmitoyl-CoA at an N-terminal longin domain and autoacylates cysteines at its Cterminus [50] in addition to mediating the palmitoylation of Vac8, presumably in a nonenzymatic way as equimolar amounts of Ykt6 and Vac8 are needed [51]. The exact
mechanism behind Vac8 palmitoylation is not clear, and further complicated by the fact
that the DHHC protein Pfa3 can palmitoylate both Vac8 [52] and Ykt6 [53]. One
possibility is that Ykt6 could act as a co-factor for Pfa3 in the way that Erf4 does for
Erf2, although this remains to be elucidated.
Regulation of deacylation
It is thus not completely clear if all fatty acylations are enzymatic and if they are
regulated. This raises the possibility that the level of regulation of acylation lies on the
deacylation step, especially for those proteins where autoacylation has been suggested.
Deacylation of proteins appears to be an enzymatic process. Despite the fast progress
that has been made, and is continously being made, in identifying and characterizing
PATs, much less is known about the thioesterases that deacylate proteins. So far, only
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two thioesterases have definitively been shown to catalyze the removal of fatty acids
from proteins, and out of those two, only one appears to be involved in regulation of the
dynamic acylation cycles seen for many intracellular proteins.
Acylprotein thioesterases
Acyl protein thioesterase 1 (APT1) was originally isolated from rat liver as a
lysophospholipase [54], but was later shown to have a preferred substrate specificity for
palmitoylated proteins over lysophosphatidylcholine and palmitoyl-CoA [25]. It is a
cytosolic protein [25] with a widespread tissue distribution [55]. Several proteins have
been identified as APT1 substrates in in vitro assays, like Ras [25], various
heterotrimeric G protein α subunits [25, 56], eNOS [10], RGS4 [25] and SNAP-23 [57].
Viral proteins, like the spike glycoprotein E2 from the Semliki Forest virus, HEF and
HA from influenza viruses and the G protein from vesicular stomatitis virus (VSV) are
also deacylated by APT1 in in vitro assays [58], although whether this has any
biological significance remains to be elucidated.
Cell-based experiments have validated APT1 as a thioesterase. In yeast cells deficient in
Apt1p, the yeast homologue of APT1, no Gαi1 deacylation takes place [56], confirming
its status as an APT1 substrate. In permeabilized platelets, infusion of APT1 protein
causes an almost 50% reduction of the steady state levels of acylation and, more
specifically, a reduction of the acylation of Gαq and concomitant reduction of membrane
association [59]. Co-expression of APT1 and eNOS increases the level of eNOS
depalmitoylation compared with expressing eNOS alone [10]. Furthermore, the
dislocation of the APT1 substrates SNAP-23 and a mutant Gsα from the plasma
membrane has also been seen on elevating APT1 levels in cells [59, 60].
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There is no specific consensus sequence for S-acylation, although some patterns of Sacylation exist, for instance N-terminal dual S-acylation and myristoylation, C-terminal
dual S-acylation and prenylation, and multiple S-acylation at cysteine string motifs [1].
Following on from the lack of distinct S-acylation motifs, there is also no clear
consensus for the sequences surrounding the thioacyl group recognized by APT1. In
fact, APT1 appears very promiscuous in its substrate specificity. In vitro, APT1 can
depalmitoylate structurally different proteins, both soluble intracellular proteins, like
eNOS and Gαi1, which are S-acylated on cysteines in proximity to an N-terminal
myristoylation site [61, 62] and Ras, which is modified at its C-terminus by both
acylation and prenylation [63], as well as integral membrane proteins. SNAP- 23 is Sacylated at multiple cysteines located in the transmembrane domain [64, 65] and the
viral proteins E2, HEF, HA and VSV G protein are S-acylated at the interface between
the transmembrane domain and the intracellular part of the proteins [66, 67].
APT1 does not, however, deacylate proteins without any discrimination. Caveolin, for
instance, an integral membrane protein which is acylated on cysteine residues in the
intracellular C-terminal domain [68] is not deacylated by recombinant APT1 under
conditions where eNOS deacylation is readily seen [10]. Moreover, not all substrates
are deacylated with the same efficiency. Rat APT1 is 10-fold more efficient in
deacylating Gαi1 compared to Ras. This preference for Gα subunits over Ras is even
more pronounced in yeast APT1, where the difference is 70-fold [56]. Also, the
activational status of the substrates may influence the catalytic efficiency displayed by
APT1 towards it; free Gαi1 is a more readily deacylated by APT1 than heterotrimeric
Gαi1 [56].
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Another explanation for the variable substrate preference could be the possibility that
APT1 could have selectivity for different fatty acids linked to its substrates. This is,
however, contradicted by the fact that two good viral APT1 substrates, the Semliki
Forest virus (SFV) protein E2 and the influenza virus protein HEF, are acylated almost
exclusively with palmitic and stearic acid, respectively, whereas the SFV E1 protein,
which is mainly palmitoylated [69], is a poor APT1 substrate [58].
The crystal structure of human APT1 has been solved and shows that the enzyme is a
member of the large enzyme group α/β hydrolases, which also includes lipases,
esterases and dehalogenases, and has a classical catalytic triad made up of Ser-114, His203, and Asp-169 in APT1 [70]. A Blast search of the APT1 sequence reveals
homologues in a wide variety of species, including humans and other mammals as well
as lower organisms such as Drosophila melanogaster, Saccharomyces cerevisiae,
Caenorhabditis elegans, Arabidopsis thaliana and Mycobacterium leprae, implying an
essential role for ATP1 as it is so well conserved throughout evolution.
APT1 inhibitors have been developed based on the structure of the H-Ras C-terminus
and in vitro assays show promise for some of the compounds, with IC50 values in the
low nanomolar ranges [71]. Their effects in vivo have not yet been determined but they
could provide a useful tool for studying the role of APT1 in cells. In general, the field
of protein acylation has suffered from a lack of specific potent inhibitors so these
compounds could be an important addition to the toolbox.
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A homologue of APT1, called lysophospholipase II or APT2 with 64% identity to
APT1, has been cloned [55]. It was reported to have activity against several lipid
substrates, with varying efficiencies. APT2 mRNA transcripts can be found in a wide
variety of tissues, suggesting it is ubiquitously expressed [55]. An interesting question
that has not yet been answered is whether APT2, just like APT1, is a cytosolic protein
thioesterase involved in protein acylation regulation. In fact, in extracts from yeast
strains where the APT1 gene is disrupted, the thioesterase activity against acylated HRas is similar to that of extracts from wild-type yeast, despite deacylation of Gαi1 being
almost completely abolished [56]. It is tempting to speculate that APT2 could be
responsible for this residual thioesterase activity. However, Blast searches using either
yeast APT1 or human APT2 sequences do not suggest that there is an APT2 protein in
yeast (R. Zeidman and A.I. Magee). Instead, the Ras deacylation could be caused by
another still unidentified thioesterase activity.
A Blast search of the human APT1 sequence reveals another homologue of APT1, with
31% identity to APT1 and with the catalytic triad conserved (R. Zeidman and A.I.
Magee). This protein was named lysophospholipase-like 1 when its gene was
discovered during the sequencing of chromosome 1 [72]. As of yet there are no reports
of this protein’s biological activity and substrates, but if any thioesterase activity of this
protein will be identified, we suggest that it should be called acyl protein thioesterase
like 1 (APTL1) in accordance with the naming of APT1.
APT1 was purified from the soluble S100 fraction of a rat liver homogenate [25] and it
does not contain any predicted transmembrane domains or obvious sites for lipid
modification. The substrates of APT1 are, however, membrane-associated, which raises
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the question of how APT1 can access its substrates efficiently. One answer could be
that APT1 might interact with membrane proteins, either its direct substrates or other
proteins that bring APT1 in close proximity to its substrates. Our own data suggest that
a significant proportion of APT1 and APT2 sediment with cellular membranes from
cultured cells (R. Zeidman and A.I. Magee, unpublished work), supporting this
speculation.
Protein-palmitoyl thioesterases
Palmitoyl-protein thioesterase 1 (PPT1) was first isolated from bovine brain extracts
based on its ability to depalmitoylate [3H]-palmitate-labeled H-Ras [73] and was also
found to depalmitoylate Gα subunits and acyl-CoAs in vitro, with a preference for a
chain length between 14 and 18 carbons [74]. Further expression studies revealed that
PPT1 is a lysosomal enzyme [75] and it is therefore unlikely to play a role in regulated
deacylation of cytoplasmic proteins. The gene encoding PPT1 was found to be located
on chromosome 1p32, the region linked to the neurodegenerative disease infantile
neuronal ceroid lipofuscinosis (INCL), and as mutations in the PPT gene were found in
INCL patients, mutated PPT1 was identified as the underlying cause of INCL [76]. One
of the features of INCL is the accumulation of granular deposits inside the cells.
[35S]cysteine-labeled lipid thioesters accumulate in immortalized lymphoblasts from
patients with INCL, and this accumulation can be reversed by adding recombinant
PPT1 to the cells [77], which is taken up and trafficked to lysosomes [78]. The normal
function of PPT1 is thus to remove the acyl chains from proteins being degraded in the
lysosome.
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A homologue of PPT1, called PPT2, has been cloned. It is also a lysosomal
thioesterase, but has a substrate specificity for palmitoyl-CoA and not palmitoylated
proteins [79] and therefore likely does not act as a thioesterase for acylated proteins
during the degradation process.
Deregulation of S-acylation implicated in diseases
Aberrant regulation of S-acylation and deacylation has been implicated in a number of
human diseases. The dysregulation of S-acylation can be caused by mutations in the
acylated proteins, or by abnormal expression of either the PAT or thioesterase involved.
Huntington’s disease, a neurodegenerative disease, is caused by a dominant mutation in
the protein huntingtin that expands the number of glutamines within an existing
polyglutamine stretch [80]. Within this region, there is a cysteine that is transiently
acylated, both in the wild-type and mutated huntingtin, but the level of acylation is
lower in the glutamine-expanded mutant than in the wild-type protein. Less acylated
forms of huntingtin display increased aggregation and formation of insoluble inclusions
in neuronal cells. Overexpression of the PAT responsible for acylation of huntingtin,
DHHC17/HIP14, reduces the number of inclusions in cells expressing mutant
huntingtin [42]. Finding a way to decrease the activity of the thioesterase responsible
for deacylation of huntingtin, which still needs to be identified, or increasing the
activity of DHHC17/HIP14 in the affected neuronal cells in patients would be an
attractive model for Hungtington’s disease treatment or prevention.
Targeting the S-acylation regulation machinery could also be a potential taget for anticancer therapies. Ras proteins, which can be both farnesylated and S-acylated, play an
13
important role in tumour progression and Ras mutations are common in human cancers
[81]. Oncogenic H-Ras mutants undergo increased cycles of acylation and deacylation
compared to wild-type H-Ras and are GTP-bound to a higher extent [8]. This opens up
the possibility to modulate Ras activity by preventing dynamic acylation of Ras and also
suggests that either inhibiting the acylation or stimulating the deacylation could have
similar effects on reducing the oncogenicity of Ras. Ras lipidation is already an antioncogenic drug target; farnesyl transferase inhibitors (FTIs) have been tested for their
efficacy as cancer drugs. Despite initial promising results, the FTIs have not been
efficient anti-tumour drugs in clinical trials, presumably because Ras is instead
geranylgeranylated following FTI treatment [82]. It is possible that S-acylation
inhibitors could prove a more efficient route in the development of treatments for Rasmediated tumours.
In fact, the deacylation of another protein, tubulin, might be a mechanism already in use
in anti-cancer therapy. Treatment of leukemic lymphoblasts with clinically relevant
concentrations of vinblastine was reported to lower the levels of [3H]-palmitate
incorporated into tubulin, as well as resulting in the known effects of vinblastine,
microtubule disassembly and apoptosis [83]. As tubulin can autopalmitoylate [84], the
regulating step in the tubulin acylation/deacylation control could be at the level of
thioesterases. Modulating thioesterase activity in cancer cells sensitive to vinblastine
might be a useful therapeutic strategy.
Future perspectives
In summary, only a single thioesterase, APT1, is known to act on cytoplasmic Sacylated proteins, although the protein acylthioesterase activity of other candidates has
14
not been thoroughly tested. Whether the substrate specificity and/or activity of this
enzyme are regulated by post-translational modification or subcellular localisation is an
open question. In this context, the association of a substantial fraction of APT1 with
cellular membranes is intriguing. However, comparison to the large family of DHHCcontaining PATs for cytoplasmic proteins (23 in man) might suggest that APT1 cannot
confer enough specificity in substrate deacylation to be a likely regulatory point for all
of them. Nevertheless, in order fully to understand the role of S-acylation in
contributing to the regulation of the function of cytoplasmic proteins it is essential to
study thioesterases as well as PATs and to elucidate the control of their opposing
effects in vivo.
Current evidence suggests that PPT1 is mainly involved in the lysosomal degradation of
acylated proteins. However, with respect to extracellular acylated proteins it will be
interesting to see whether PPT1, which can be released from cells before being taken up
into lysosomes, could deacylate some signalling molecules (e.g. Wnts) extracellularly
and thus modulate their spread within tissues and function.
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Figure legend
Figure 1: Dynamic S-acylation of Lck.
A: The S-acylation (represented by thick grey lines) of Lck at two cysteines near an Nterminal myristoylation (represented by thin black line). A protein acyl transferase
(PAT) mediates the addition of palmitate from palmitoyl coenzyme A (PalCoA)
whereas a thioesterase (TE) catalyzes the hydrolysis of palmitic acid (PalCOOH) from
Lck. B: Accumulation of proteins with phosphorylated tyrosines in lysates from human
blast T cells exposed to 1 mM pervanadate (PV) for the times indicated, analyzed by
phosphotyrosine (PY) western blotting. C: Lck immunoprecipitated from human T
lymphoblast cells pulsed with 3H-palmitate, and chased for the indicated times.
Pervanadate (PV) treatment causes increased activation of Lck, seen as increasing
26
amounts of active, serine-phosphorylated Lck (p60 SP) and also increases the palmitate
turnover on Lck compared to control cells not treated with pervanadate (ctrl).
Method: T lymphoblasts were a kind gift of Dr. Julian Ng and Dr. Doreen Cantrell at
CRUK-LRI. They were prepared from outdated blood bags and maintained for up to 2
weeks in RPMI 1640 medium containing 10% foetal calf serum and Interleukin 2
(20ng/ml) at 50-200 x 104 cells/ml [85]. Cells were quiesced by removal of IL2 for 2-3
days, spun down and resuspended in labelling medium containing 5mM pyruvate, then
labelled with 3H palmitic acid (30-60Ci/mmol, approx. 200 µCi/ml) and incubated at
37C for 45 min, then spun down and resuspended in chase medium (labelling medium
containing 80µM palmitic acid, approx. 20 fold excess over tritiated palmitic acid, with
or without 1 mM PV). This was followed by rapid washing with ice-cold PBS, lysis,
immunoprecipitation , separation by SDS-PAGE and fluorography to detect 3Hpalmitate labelled bands.
Acknowledgements
We thank Julian Ng and Doreen Cantrell at Cancer Research UK-London Research
Institute for the kind gift of human T lymphoblasts. Work in the Magee laboratory is
supported by the UK Medical Research Council and Biotechnology and Biological
Sciences Research Council.
27