©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
The Myristoyl Moiety of Myristoylated Alanine-rich C Kinase Substrate (MARCKS) and MARCKS-related Protein Is Embedded in the Membrane (*)

(Received for publication, May 22, 1995)

Guy Vergères (§) Stéphane Manenti (¶) Thomas Weber (1)(**) Christoph Stürzinger

From the Department of Biophysical Chemistry, Biocenter of the University of Basel, 4056 Basel, Switzerland and theDepartment of Biochemistry, Swiss Federal Institute of Technology, 8092 Zürich, Switzerland

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Members of the myristoylated alanine-rich protein kinase C substrate (MARCKS) family are involved in several cellular processes such as secretion, motility, mitosis, and transformation. In addition to their ability to bind calmodulin and to cross-link actin filaments, reversible binding to the plasma membrane is most certainly an important component of the so far unknown functions of these proteins. We have therefore investigated the binding of murine MARCKS-related protein (MRP) to lipid vesicles. The partition coefficient, K(p), describing the affinity of myristoylated MRP for acidic lipid vesicles (20% phosphatidylserine, 80% phosphatidylcholine) is 5-8 10^3M, which is only 2-4 times larger than the partition coefficient for the unmyristoylated protein. Interestingly, the affinity of MRP for acidic lipid membranes is 20-30-fold smaller than reported for murine MARCKS (Kim, J., Shishido, T., Jiang, X., Aderem, A. A., and McLaughlin, S.(1994) J. Biol. Chem. 269, 28214-28219). Since only a marginal binding could be observed with neutral phosphatidylcholine vesicles, we propose that electrostatic interactions are the major determinant of the binding of MRP to pure lipid membranes. Although the myristoyl moiety does not contribute drastically to the binding of MRP to vesicles, photolabeling experiments with a photoreactive phospholipid probe show that the fatty acid is embedded in the bilayer. The same membrane topology was found for bovine brain MARCKS. Since the relatively low affinity of MRP for vesicles is insufficient to account for a stable anchoring of the protein to cellular membranes, insertion of the myristoyl moiety into the bilayer might favor the interaction of MRP with additional factors required for the binding of the protein to intracellular membranes.


INTRODUCTION

During the last decade, a growing number of myristoylated proteins have been identified in eukaryotic cells. Myristoylation refers to the attachment of a C14 saturated fatty acid (myristate) to the N-terminal glycine residue of substrate proteins. The soluble enzyme N-myristoyl transferase (NMT) (^1)cotranslationally transfers a myristoyl moiety from the cofactor myristoyl-coenzyme A to the N terminus of nascent proteins containing the consensus sequence (Gly-X-X-X-Ser/Thr) (Johnson et al., 1994). Although myristoylation is believed to be irreversible (James and Olson, 1989), some evidence has been presented for a demyristoylation activity in cytoplasmic fractions of brain synaptosomes (da Silva and Klein, 1990; Manenti et al., 1994). The importance of myristoylation has been highlighted by several studies in which removal of the fatty acid leads to dramatic alterations of cellular functions; myristoylation is necessary for the transforming activity of v-Src (Kamps et al., 1985), for the transcriptional suppressor activity of the HIV nef gene product (Huang and Jolicoeur, 1994), and for signaling and transforming functions of G protein alpha subunits (Gallego et al., 1992). How myristoylation mediates the function of these proteins is, however, unclear. One function of the fatty acid moiety is to promote protein-protein interactions. In this respect myristoylation is required for high affinity interaction of G protein alpha subunits with the beta subunits (Linder et al., 1991) and enhances the binding of the HIV nef gene product to CD4 (Harris and Neil, 1994). Myristoylation also influences the conformation of proteins as seen for the catalytic subunit of cAMP-dependent protein kinase (Yonemoto et al., 1993) and for p56lck, a member of the Src family, which are stabilized by the acylation (Nadler et al., 1993). Myristoylation also promotes protein-ligand interactions as indicated by the increased ability of myristoylated ADP-ribosylation factor to exchange nucleotides (Franco et al., 1995). The most obvious function of the myristoyl moieties is to mediate membrane binding. Evidences for this role have been presented for several proteins by analyzing the subcellular localization of unmyristoylated mutants. This approach has successfully demonstrated that the myristoyl moieties of nitric oxide synthase (Busconi and Michel, 1994), the myristoylated alanine-rich protein kinase C substrate (MARCKS) (Graff et al., 1989), the product of the HIV nef gene (Yu and Felsted, 1992), Src (David-Pfeuty et al., 1993), and the alpha subunit of Gz (Hallak et al., 1994) are necessary for membrane binding. Myristoylation, however, is not sufficient to bind proteins to membranes for several reasons. First, myristoylated proteins are found in almost every subcellular compartment including the cytosol (Magee and Courtneidge, 1985; Blenis and Resh, 1992). Second, the apparent partition coefficient, K(p), describing the binding of acylated peptides to vesicles is about 10^4M and corresponds to a Gibbs free energy of binding of 8 kcal, which is not sufficient to firmly anchor proteins to membranes (Peitzsch and McLaughlin, 1993). These observations indicate that proteins need some information, in addition to myristoylation, in order to be targeted to their final subcellular localization and to bind to membranes. A search for receptors for proteins such as Src (Sigal and Resh, 1993), MARCKS (George and Blackshear, 1992), and nitric oxide synthase (Busconi and Michel, 1994) has, however, been unsuccessful so far. Recently, the emphasis has shifted to dual binding motifs. Some myristoylated proteins, such as several members of the G protein alpha subunit family (Casey, 1994) as well as several members of the Src family (Resh, 1994), are palmitoylated at a cysteine residue adjacent to the myristoylated N-terminal glycine residue. This double acylation is sufficient to anchor proteins firmly to the membrane (Blenis and Resh, 1992). Other proteins such as MARCKS (Taniguchi and Manenti, 1993; Kim et al., 1994a, 1994b) and Src (Sigal et al., 1994) contain a basic domain that interacts electrostatically with membranes containing negatively charged lipids. The affinity of the basic domains for acidic phospholipids could provide the cell with a targeting mechanism to the inner leaflet of the plasma membrane, which contains up to 30% negatively charged lipids (Op den Kamp, 1981). In this respect, McLaughlin and co-workers have proposed that the myristoyl moiety and the basic domain of MARCKS act synergistically to firmly anchor the protein to acidic lipid membranes (Kim et al., 1994b) explaining the association of this protein with intracellular membranes (Albert et al., 1986; Thelen et al., 1991) (for a review see Vergères et al.(1995)).

The MARCKS family comprises two groups of proteins. The members of the first group, referred as to MARCKS, are 30-35-kDa proteins encoded by the macs genes. The second group, referred as to MARCKS-related protein (MRP), is composed of 20-kDa proteins encoded by the mrp genes and also called MacMARCKS and F52 in the literature (Aderem, 1992; Blackshear, 1993). Although macs and mrp are ubiquitously expressed, higher levels of mRNA are found in brain (MARCKS and MRP) and in reproductive tissues (MRP) (Lobach et al., 1993). The N terminus, which contains the consensus sequence for myristoylation, and the basic domain with the site of phosphorylation by protein kinase C, are conserved in MRP and MARCKS (Fig. 1). Receptor-mediated activation of protein kinase C alters the subcellular localization of MARCKS. Phosphorylation of the basic domain results in the translocation of MARCKS from the plasma membrane to the cytosol. This phenomenon is reversible, since dephosphorylation of MARCKS is concomitant with the relocation of the protein at the plasma membrane (Thelen et al., 1991). Recently, it was proposed that phosphorylation induces the cycling of MARCKS between the plasma membrane and lysosomes (Allen and Aderem, 1995). Although the molecular mechanisms underlying the association of MARCKS with membranes have been relatively well characterized (Taniguchi and Manenti, 1993; Kim et al., 1994a, 1994b; Allen and Aderem, 1995), nothing has been known about MRP so far. In analogy to MARCKS, one would expect that MRP binds to membranes. The slight differences in the sequences of the N terminus and of the basic domain might, however, modulate the binding and consequently give us some insights on how myristoylated proteins interact with membranes (Fig. 1). For these reasons we have purified the unmyristoylated form (unmyr MRP) as well as the myristoylated form (myr MRP) of recombinant murine MRP expressed in Escherichia coli and investigated the ability of the myristoyl moiety and of the basic domain to promote membrane binding. Although the myristoyl moiety is believed to contribute to the binding of proteins to membranes by inserting into the bilayer, no direct proof of this putative hydrophobic interaction has been presented until now. We have therefore also investigated the membrane topology of murine MRP and bovine MARCKS using a novel photoreactive phospholipid probe.


Figure 1: Structure of proteins of the MARCKS family. The amino acid sequence of the myristoylated N terminus, which contains the consensus sequence GXXXS recognized by NMT, as well as of the basic domain, which contains the serines phosphorylated by protein kinase C, are shown for murine MRP (uppersequence). Nonconserved residues (underlined in MRP sequence) are shown for murine and bovine MARCKS (lowersequences). The amino acid residues are numbered (top) according to the MRP sequence.




EXPERIMENTAL PROCEDURES

Materials

Calmodulin-agarose, protease inhibitors (leupeptin, chymostatin, aprotinin, phenylmethysulfonyl fluoride (PMSF)), dansyl-calmodulin, and ATP were from Sigma. Egg L-alpha-phosphatidylcholine (PC) and brain L-alpha-phosphatidylserine (PS) were from Aventi Polar Lipids (Alabaster). The DE52 anion exchanger was obtained from Whatman, phenyl-Sepharose CL-4B from Pharmacia Biotech Inc., and hydroxylapatite from Bio-Rad Laboratories. Tryptone and yeast extract were from Oxoid Ltd. (Basingstoke, United Kingdom). Ampicillin, kanamycin, and isopropyl-1-thio-beta-D-galactopyranoside (IPTG) were from Gerbu Trading GmbH (Gaiberg, Germany). [-P]ATP (3000 Ci/mmol) and [^3H]myristate (50-80 mCi/mmol) were obtained from Amersham Corp. 1-O-Hexadecanoyl-2-O-[9-[[[2-(I)iodo-4-((trifluoro-methyl)-^3H-diazirin-3-ylbenzyl]oxy]carbonyl]nonanoyl]-sn-glycero-3-phosphocholine ((I)-TID-PC/16) (>2000 Ci/mmol) was synthesized as described (Weber et al., 1994) from the corresponding tin precursor (Weber and Brunner, 1995). The E. coli strain JM109(DE3) was obtained from Novagen. The plasmids pET3dF52M1 and pBB131NMT were gifts from Perry Blackshear (Duke University Medical Center, Durham) and Jeffrey Gordon (Washington University, School of Medicine, St. Louis), respectively.

Expression of MRP

Expression of unmyr MRP

The E. coli strain JM109(DE3) was transformed by electroporation with the plasmid pET3dF52M1, which contains the gene coding for mouse MRP (Umekage and Kato, 1991; Blackshear et al., 1992). The cells containing the plasmid were selected with 50 µg/ml ampicillin. A frozen stock of transformed colonies was used to inoculate 30 ml of LB media (10 g/liter tryptone, 5 g/liter yeast extract, 10 g/liter NaCl, pH 6.5, with HCl) containing 50 µg/ml ampicillin and grown overnight at 37 °C. 2 liters of LB media containing 50 µg/ml ampicillin was inoculated with 20 ml of the overnight culture and grown at 37 °C to log phase (A = 0.5-0.8). At this point 0.4 mM IPTG was added to induce protein expression, and the culture was grown for 5 additional hours. The cells were harvested and the paste (about 7-8 g, wet weight) was kept frozen at -20 °C.

Expression of myr MRP

The E. coli strain JM109(DE3) was transformed with the plasmid pBB131NMT, which contains the gene coding for yeast NMT (Duronio et al., 1990) and selected with 50 µg/ml kanamycin. The strain JM109(DE3)-pBB131NMT was then transformed with the plasmid pET3dF52M1 and selected in the presence of 50 µg/ml ampicillin and kanamycin. The bacterial culture was performed exactly as described above except that 50 µg/ml kanamycin was added to the media. Coexpression of MRP and NMT was induced by addition of 0.4 mM IPTG to the log phase culture.

Radioactive Labeling of MRP with

myristate

Two ml of LB containing the appropriate antibiotics were inoculated with 20 µl of an overnight culture of either the JM109(DE3)-pET3dF52M1 or the JM109(DE3)-pBB131NMT-pET3dF52M1 strain and grown for 2 h at 37 °C. The culture was transferred to a tube, in which 100 µCi of [^3H]myristate in ethanol had previously been dried, and grown for an additional hour. The expression of MRP and NMT was induced by adding 0.4 mM IPTG, and the cells were grown for 5 additional hours before harvesting.

Analysis of MRP Expression

The cells from 1 ml of culture (3-4 mg, wet weight) were washed with 1 ml of buffer A (10 mM Tris, 1 mM EDTA, pH 7.4) and resuspended in 50 µl of lysis buffer (buffer A + 1% Triton X-100). The solution was heated at 95 °C for 10 min and cooled on ice for 5 min, and the precipitate was removed by centrifugation in an Eppendorf centrifuge at 14,000 rpm for 5 min. A volume of supernatant corresponding to 150 µg, wet weight, of cell paste in sample buffer (50 mM Tris, pH 6.8, 10% glycerol, 0.2% SDS, 0.02% bromphenol blue) was loaded on a 10% polyacrylamide gel. Proteins were visualized by Coomassie Blue staining of the gels. Myristoylation of MRP was detected by fluorography of the cell extracts grown in the presence of [^3H]myristate (see above).

Purification of Proteins under Nondenaturing Conditions

Purification of unmyr MRP

5 g, wet weight, of cell paste (about 1.5 liters of culture) was resuspended by pipetting in 60 ml of lysis buffer. The cells were lysed by sonicating twice for 4 min with a 4-min rest. Since sonication was sufficient to reduce the viscosity of the solution, DNase was not used. Following addition of 1 mM fresh PMSF, the suspension was centrifuged at 20,000 rpm in a SS34 Sorval rotor for 30 min at 4 °C. The supernatant was collected, and the volume was adjusted to 100 ml with lysis buffer following addition of solid NaCl to a final concentration of 3 M. The solution was loaded with a flow rate of 120 ml/h on a 60-ml phenyl-Sepharose column (2.4-cm diameter) equilibrated in buffer A containing 3 M NaCl. The fractions of the pass-through, which contains unmyr MRP, were pooled, 0.1% Triton X-100 was added, and the solution was dialyzed overnight in 3 liters of buffer A with a change after 3 h. The protein solution was loaded on a 20-ml hydroxylapatite column (1.0-cm diameter, 120 ml/h) equilibrated in buffer B (10 mM Tris, 1 mM EDTA, 0.1% Triton X-100, pH 7.4). After washing with 50 ml of buffer B, unmyr MRP was eluted with a phosphate gradient in buffer B (2 120 ml; 0-0.3 M phosphate). The fractions containing unmyr MRP (about 60-120 mM phosphate) were pooled and dialyzed overnight in 1 liter of buffer A. Triton X-100 was removed by loading the protein solution on a 5-ml DE52 column (1.0-cm diameter, 120 ml/h) equilibrated in buffer A and washing the column with 50 ml of buffer A. unmyr MRP was finally eluted from the DE52 column at 30 ml/h with buffer A containing 0.3 M NaCl. The fractions containing unmyr MRP were pooled, dialyzed in 1 liter of buffer C (10 mM Tris, 0.1 mM EDTA, pH 7.4), and stored frozen at -70 °C. In order to prevent proteolysis, protease inhibitor mixtures (PIC) were added to the solutions; the lysis buffer contained a concentrated mixture (cPIC: 1 mM PMSF and 1 µg/ml each of aprotinin, chymostatin, and leupeptin), and all subsequent solutions contained a ``medium'' mixture (mPIC: 0.1 mM PMSF and 0.1 µg/ml each of aprotinin, chymostatin, and leupeptin) with the exception of the final dialysis solution, which contained a diluted mixture (dPIC: 10 µM PMSF and 0.1 µg/ml each of aprotinin, chymostatin, and leupeptin). Protein concentration was determined by the method of Lowry using bovine serum albumin as a reference (Lowry et al., 1951).

Purification of myr MRP

5 g, wet weight, of cells expressing myr MRP (about 1.5 liters of culture) were lysed and loaded on a 60-ml phenyl-Sepharose column as described for unmyr MRP. Following loading of the cell extract, the column was washed with 50 ml of buffer A containing 3 M NaCl. Myr MRP was eluted with 200 ml of buffer A containing 2 M NaCl and with a NaCl gradient in buffer A (2 150 ml; 2.0-0.5 M NaCl). Under these conditions, myr MRP elutes with the 2 M NaCl wash as well as at the beginning of the gradient. 0.1% Triton X-100 was added to the fractions containing myr MRP, and the protein was further purified on hydroxylapatite and DE52 as described for unmyr MRP. Since this procedure does not allow the purification of myr MRP to homogeneity, an additional step was introduced. The fractions containing unmyr MRP were pooled and dialyzed overnight in 1 liter of buffer A. After the addition of 7 mM CaCl(2) the solution was loaded on a 5-ml calmodulin-agarose affinity column (1.0-cm diameter, flow rate of 120 ml/h) equilibrated with buffer D (10 mM Tris, 7 mM CaCl(2), pH 7.4). The column was washed with 50 ml of buffer D, and myr MRP was eluted at 30 ml/h with 25 ml of buffer E (10 mM Tris, 2 mM EGTA, 0.3 M NaCl, pH 7.4). The fractions containing myr MRP were pooled, dialyzed in 1 liter of buffer C, and stored frozen at -70 °C. PIC was added to all solutions as described for unmyr MRP.

Purification of MARCKS

unmyr MARCKS (Manenti et al., 1993) as well as myr MARCKS (Manenti et al., 1992) were purified from calf brain under nondenaturing conditions as described previously.

Binding of MRP to Calmodulin

The binding of MRP to dansyl-calmodulin was measured fluorimetrically in a Jasco FP-777 spectrophotometer at 22 °C (Johnson and Wittenauer, 1983; Blackshear et al., 1992). The solutions contained 10 mM Tris, pH 7.4, 0.1 mM EDTA, 90 mM KCl, 1 mM CaCl(2), 50 nM dansyl-calmodulin, and 250 nM MRP. The excitation wavelength was 340 nm (2-nm bandwidth), and the emission spectra were measured between 390 and 600 nm (2-nm bandwidth).

Phosphorylation of MRP

MRP was phosphorylated by the catalytic domain of protein kinase C (PKM) (Uchida and Filburn, 1984). The reaction mixture contained 1 µM MRP, 2.5 nM PKM, 200 µM [-P]ATP (0.25 µCi), 10 mM MgCl(2), 100 mM NaCl, 0.1 mM EDTA, 10 mM Tris, pH 7.4, and mPIC. Phosphorylation of MRP was allowed to proceed for 2 h at 37 °C. The reaction was stopped by the addition of sample buffer, heated for 3 min at 95 °C, and loaded on a 10% polyacrylamide SDS gel. Phosphorylated MRP was detected by autoradiography of the gels.

Preparation of Sucrose-loaded Vesicles

A suspension containing 125 mg of egg PC or 125 mg of a mixture of egg PC and brain PS (4:1, w/w) was dried, resuspended in 1.5 ml of buffer F (10 mM Tris, 1 mM EDTA, 100 mM NaCl, pH 7.4) containing 170 mM sucrose (Hope et al., 1985). The suspensions were freeze-thawed 10 times in liquid nitrogen and extruded 10 times through a 100-nm carbonate filter. The vesicles were diluted with 5 volumes of buffer F and centrifuged at 50,000 rpm in a TL-100.3 rotor (Beckman) for 30 min at 22 °C. The vesicles were resuspended in 200 µl of buffer F, and the lipid concentration was measured by determining the phosphate content (Böttscher et al., 1961). The suspension was adjusted to 20 mM phosphate with buffer F.

Binding of MRP to Vesicles

The binding of MRP to sucrose-loaded vesicles was performed as described (Hope et al., 1985). 100 µl of a solution containing 1 µM MRP and 0-10 mM lipid vesicles in buffer F containing dPIC and 0.001% Triton X-100 was incubated at 30 °C for 30 min. MRP bound to vesicles was separated from unbound MRP by centrifugation at 70,000 rpm in a TL-100.3 rotor for 50 min at 22 °C. To estimate the partitioning of MRP between membrane and solution, 25 µl of the supernatant in sample buffer was loaded on a 10% SDS-polyacrylamide gel. The gel was silver-stained, and the unbound MRP concentration was determined by volume integration with local background subtraction at 100-µm resolution on a personal densitometer from Molecular Dynamics. The intensity of the MRP band in which lipids have been omitted was used as a reference (100% or 1 µM unbound MRP). The signal was linear in the range used for these studies (0-500 ng of MRP).

Determination of the Partition Coefficient for the Binding of MRP to Membranes

The partition coefficient K(p) (M) describing the binding of MRP to membranes was determined as described by McLaughlin and co-workers (Kim et al., 1994b) by fitting the equation,

where f(b) is the fraction of MRP bound to the membrane. f(b) was calculated by estimating the concentration of MRP in the supernatant following centrifugation of the vesicles as described in the preceding paragraph. [L] (M) is the accessible lipid concentration and can be estimated to 50% of the total lipids for 100-nm vesicles. K(p) (M) relates the mole ratio of MRP bound to the vesicles to the unbound MRP concentration and can be regarded as an apparent association constant. K(p) was obtained by a least square fitting of the above equation. The reciprocal of K(p), 1/K(p) (M), can be regarded as an apparent dissociation constant that is equal to the lipid concentration at which 50% of MRP is bound to the vesicles (Peitzsch and McLaughlin, 1993).

Photolabeling of Membrane-bound MRP and MARCKS

1.5 µM MARCKS or 5 µM MRP were incubated with 60 µM 100-nm lipid vesicles containing 5 µCi of (I)-TID-PC/16 (Weber et al., 1994) in buffer A containing 50 mM NaCl for 30 min at 30 °C in a final volume of 110 µl. For photoactivation, the lipid-vesicles solution in an Eppendorf tube was irradiated for 1 min in a Pyrex vessel mounted approximately 10 cm from a Suss LH 1000 lamphouse (Karl Suss, Waterbury Center, VT) equipped with a 350-watt high pressure mercury lamp. Following photolysis, the protein solution was delipidated in the presence of 1% Triton X-100 as follows: 15 µl of buffer A containing 50 mM NaCl and 10% Triton X-100 was added to the protein solution and incubated for 15 min at room temperature. The solution was transferred to an Eppendorf tube containing 50 µl of DE52, equilibrated in buffer A containing 50 mM NaCl, and incubated for 15 min at room temperature to allow binding of MARCKS or MRP to the gel matrix. Under these conditions the fraction of (I)-TID-PC/16 that has not reacted with MRP or MARCKS does not bind to the gel. The excess reagent was removed following centrifugation of the suspension for 30 s at 14,000 rpm in an Eppendorf centrifuge. After six washes with 500 µl of buffer A containing 50 mM NaCl almost no additional radioactivity could be removed. MRP and MARCKS were eluted by incubating the gel material with 50 µl of buffer A containing 400 mM NaCl. The label associated with the proteins was detected by loading 10 µl of the samples on a 10% SDS-polyacrylamide gel. The proteins were visualized by Coomassie Blue staining, and (I)-TID-PC/16 associated with the proteins was detected by autoradiography of the gels (4 days exposure). After delipidation, the protein solution still contained a large fraction of (I)-TID-PC/16-lipid-cross-links, which were associated noncovalently with the proteins and which migrated with the front during electrophoresis. Some radioactivity, however, still bound nonspecifically to the protein, thus increasing the background signal. This problem arises most likely from the strong association of (I)-TID-PC/16-PS-cross-link products to MRP and MARCKS and can be eliminated by adding 2 µg of phosphatidylserine to the protein solution before loading the sample on the gel. The radioactivity associated with the proteins was quantitated by volume scanning densitometry of the autoradiograms.

Enzymatic Demyristoylation of MRP and of MARCKS

Myr MRP and MARCKS were demyristoylated enzymatically as described previously (Manenti et al., 1994). The incubation medium contained 18 µl of delipidated MRP or MARCKS that had previously been labeled with (I)-TID-PC/16 and 12.5 µl of a cytosolic fraction of brain synaptosomes. In addition, the solution contained 1 mM EGTA, 0.5 mM ATP, 10 mM Tris, pH 7.0, 20 mM Hepes, pH 7.0, and cPIC in a final volume of 50 µl. After 24 h at 37 °C a fresh synaptosomal fraction (9 µl) as well as ATP and cPIC were added, and the solution was incubated for 20 additional hours. In control experiments, the synaptosomal fraction was omitted and replaced by water. The reactions were stopped by heating at 95 °C for 10 min. The precipitated proteins were removed by centrifugation, and MRP and MARCKS were analyzed by loading 30 µl of the supernatant on a 10% SDS-polyacrylamide gel.


RESULTS

Expression of MRP

In order to bypass the inability of bacteria to N-myristoylate proteins, Gordon and co-workers have developed a system capable of producing myristoylated proteins by coexpressing yeast NMT (Duronio et al., 1990). This system was used to express myr MRP in E. coli (Blackshear et al., 1992). Since the capacity of this coexpression system to produce myr MRP in quantities sufficient for its purification has not been investigated (Blackshear et al., 1992) our first goal was to characterize the expression of MRP.

Addition of IPTG to a log phase culture of JM109(DE3) cells containing the plasmid pET3dF52M1 encoding MRP induces the expression of a heat-stable protein, which appears as a double band on a 10% SDS-polyacrylamide gel and whose apparent molecular weight is 37 kDa (Fig. 2, lane4). Since the double band is not present in the strain containing the pBB131NMT plasmid only (lane2), the protein results from transcription of the mrp gene. Coexpression of NMT with MRP results in a slight decrease in the apparent molecular weight of the double band (lane6). That this shift results from myristoylation of MRP is shown in Fig. 3. When E. coli cells are grown in the presence of [^3H]myristate a double band, whose apparent molecular weight corresponds to MRP, is labeled radioactively in the strain coexpressing MRP and NMT (lane1) but not in the strain expressing MRP only (lane2). Inspection of the Coomassie Blue-stained gel (Fig. 2) shows that the myr MRP double band is apparently not significantly contaminated with unmyr MRP (lane6). NMT can therefore efficiently myristoylate MRP (>90%) in the E. coli strain JM109(DE3). A comparison of the intensities of the Coomassie Blue-stained MRP bands of the heated cell extracts with known amount of the purified protein (see next paragraph) shows that approximately 20 mg of MRP is produced per 5 g of cell paste (about 1.5 liters of culture). In the absence of IPTG, unmyr (lane3) and myr (lane5) MRP are expressed to a lower but measurable extent, showing that the DE3lacUV5 promoter is partially leaky. Finally, we have observed that the starting pH of the media has an effect on the expression of MRP. Since the expression gradually decreases above pH 7.5 (data not shown) we have chosen a starting pH of 6.5 for the LB media.


Figure 2: Expression of MRP in E. coli. The E. coli strains JM109(DE3) containing the plasmids with the genes coding for NMT (lanes1 and 2), MRP (lanes3 and 4), and MRP + NMT (lanes5 and 6) were grown in the absence (lanes1, 3, and 5) or in the presence (lanes2, 4, and 6) of 0.4 mM IPTG. The cells were lysed for 10 min at 95 °C in the presence of 1% Triton X-100, and the equivalent of 150 µg, wet weight, of cell paste was loaded on a 10% polyacrylamide SDS gel. Proteins were visualized by Coomassie Blue staining. Molecular weight standards are shown to the right. The MRP double bands are indicated with arrowheads.




Figure 3: Myristoylation of MRP in E. coli. The equivalent of 150 µg, wet weight, of cells grown in the presence of [^3H]myristate was lysed as described in Fig. 2and analyzed on a 10% SDS-polyacrylamide gel. The radioactive label was detected by fluorography. Lane1, heated extract of cells containing the plasmids with the genes coding for MRP and NMT. Lane2, cells containing the plasmid coding for MRP only.



Purification of MRP

Proteins of the MARCKS family remain soluble at high temperature and low pH. These properties have allowed the purification of myr MRP from mouse alveolar macrophages (Li and Aderem, 1992) as well as of recombinant murine unmyr MRP from E. coli (Verghese et al., 1994). A recent report shows that heating increases the binding of a monoclonal antibody to MARCKS, indicating that extreme conditions might alter the structure of MARCKS irreversibly (Rose et al., 1994). In order to avoid potential denaturation problems we have developed a mild purification method for MRP using a combination of absorption (phenyl-Sepharose, hydroxylapatite) and anion exchange (DE52) chromatography. This procedure allows the reliable purification of about 2-3 mg of unmyr MRP/1.5 liters of culture (Fig. 4, lane2). In contrast to the unmyristoylated protein, myr MRP is retained on phenyl-Sepharose in the presence of 3.0 M NaCl and slowly elutes at lower salt concentrations (about 2.0 M NaCl) together with other proteins. An affinity chromatography on calmodulin-agarose, is required to obtain a pure protein. Although the yield is lower than for unmyr MRP, we could purify the myristoylated protein in quantities sufficient for biochemical analysis (typically 1 mg/1.5 liters of culture) (lane3). We should point out that the behavior of myr MRP on the phenyl-Sepharose column strongly depends on the chromatographic conditions used. Changing the amount of cell paste, the size of the column, or the volume of buffers dramatically decreases the yield of the purified protein.


Figure 4: Purification of MRP. Coomassie Blue-stained SDS-polyacrylamide gel (10%) of 0.4 µg each of purified unmyr MRP (lane2) and myr MRP (lane3). Molecular mass standards (67, 43, and 30 kDa) are shown in lane1.



Characterization of Purified MRP

Before investigating the interactions of MRP with membranes we have characterized the purified proteins. The amino acid composition of unmyr and myr MRP are identical and consistent with the published sequence (Umekage and Kato, 1991). In addition, sequencing of both proteins indicates that the N terminus of unmyr MRP is free, whereas myr MRP is blocked. Interestingly, Blackshear etal.(1992) have reported an apparent molecular weight of 50 kDa, which significantly differs from the value we have observed (37 kDa). This discrepancy probably results from differences in the conditions used for electrophoresis (e.g. SDS, acrylamide, and bisacrylamide concentrations). These differences prompted us to determine the molecular weight of the unmyristoylated protein by sedimentation equilibrium. In agreement with the calculated value of 20,165, a molecular mass of 20 ± 1 kDa was measured for the monomeric form of the protein. (^2)

Since protein kinase C-dependent phosphorylation and binding to calmodulin are two hallmarks of proteins of the MARCKS family, we have characterized the interactions of purified MRP with these proteins. The binding of MRP to calmodulin was assessed using dansyl-calmodulin as a probe (Fig. 5). Incubation of 50 nM dansyl-calmodulin with 250 nM unmyr and myr MRP induces a shift in the fluorescence emission spectrum of dansyl from 500 to 490 nm as well as a 2.3-fold increase in the fluorescence intensity at 490 nm. These data show that both unmyr and myr MRP bind to calmodulin, in agreement with the observed retention of the proteins on a calmodulin affinity column. Finally, both unmyr and myr MRP are substrates for PKM, the catalytic fragment of protein kinase C, as judged by the incorporation of radioactive phosphate in the MRP double band (Fig. 6, lanes1 and 2, respectively).


Figure 5: Binding of MRP to calmodulin. 250 nM unmyr (3) and myr (4) MRP were incubated with 50 nM dansyl-calmodulin in the presence of 1 mM CaCl(2). Fluorescence spectra were recorded between 390 and 600 nm with excitation at 340 nm. 1, buffer; 2, calmodulin alone.




Figure 6: Phosphorylation of MRP by PKM. 1 µM unmyr (lane1) and myr (lane2) MRP were incubated with 2.5 nM PKM in the presence of 200 µM [P]ATP (0.25 µCi) and 10 mM MgCl(2) for 2 h at 37 °C. The samples were analyzed on a 10% SDS-polyacrylamide gel. A, Coomassie Blue staining; B, autoradiography.



The appearance of MRP as a double band raises the possibility that E. coli might produce heterogeneous proteins. Several lines of evidence suggest, however, that this is not the case. First, the double band could never be separated on the various columns used for the purification. In addition, myristoylation as well as phosphorylation do not alter the double band pattern, indicating that the structure of the N terminus and the phosphorylation state of the basic domain are identical for each band. Finally, MRP is also recognized as a double band on Western blots of macrophage extracts, (^3)suggesting that this pattern might result from a conformational equilibrium between two forms of the protein in electrophoresis buffer as a consequence of the weak affinity of the negatively charged SDS for the acidic MRP.

Binding of MRP to Lipid Vesicles

In order to estimate the contribution of the myristoyl moiety and of the basic domain to the binding of MRP to membranes, we have incubated unmyr and myr MRP with sucrose-loaded vesicles containing either neutral or acidic lipids. Unbound MRP was separated from the membrane-bound population by sedimentation of the vesicles at 100,000 g, and the supernatant was analyzed on a 10% SDS-polyacrylamide gel (Fig. 7). This analysis shows that 1 µM unmyr MRP binds to 2.5 mM PC/PS (4:1) vesicles (lane2) but not to PC vesicles (lane3). The behavior of myr MRP is qualitatively similar since the protein binds to acidic lipid vesicles (lane2) much stronger than to neutral vesicles (lane3). There is, however, a notable quantitative difference, since 1 µM myr MRP completely binds to PC/PS (4:1) vesicles whereas a fraction of unmyr MRP still remains unbound. In order to further investigate the role of the myristoyl moiety in the increased affinity of myr MRP for PC/PS (4:1) vesicles, we have compared the binding of myr and unmyr MRP as a function of the lipid concentration (Fig. 8, A and B, respectively). In order to minimize self aggregation and/or nonspecific binding of MRP to the polypropylene tubes, and in analogy to studies with MARCKS (Kim et al., 1994b), we have added 0.001% Triton X-100 to the solutions. Apart from allowing more reproducible measurements, the detergent has no measurable effect on the interaction of MRP with membranes. A 10-fold increase in Triton X-100, e.g. 0.01%, does not change the apparent partition coefficient for the binding of MRP to PC/PS vesicles (4:1) (not shown). Since unmyr and myr MRP remain in solution following centrifugation in the absence of lipids, nonspecific binding of the proteins to the microcentrifuge tubes or aggregation can be ruled out. We can therefore calculate the fraction of membrane-bound protein following quantification of the amount of protein remaining in the supernatant after centrifugation of the vesicles. By fitting the data with (see ``Experimental Procedures'') we have estimated that the apparent partition coefficient for unmyr MRP is 2 10^3M. The data for myr MRP could not be fitted as well as for unmyr MRP. A K(p) of 5 10^3M perfectly fits the data up to 70% binding but underestimates the affinity of the myristoylated protein at higher lipid concentrations. On the other hand, a K(p) of 8 10^3M slightly overestimates the affinity at low lipid concentrations but better fits the upper part of the curve. We therefore estimate that the myristoylated protein binds to PC/PS (4:1) vesicles with an apparent partition coefficient ranging between 5 10^3 and 8 10^3M. Since the data describing the binding of myr MRP to PC/PS (4:1) vesicles cannot be perfectly fitted with a single partition coefficient, the mechanism of binding of the myristoylated protein might be more complex than for unmyr MRP and awaits further investigation. Finally, a weak but significant binding of myr MRP to 10 mM PC vesicles could be observed (not shown). However, a partition coefficient could not be obtained since the affinity of the protein is too low to allow reproducible measurements. Since we have consistently observed that <50% myr MRP binds to 10 mM PC vesicles, we can, however, set an upper limit for the partition coefficient of myr MRP to neutral vesicles, namely 10^2M.


Figure 7: Binding of MRP to phospholipid vesicles. 1 µM MRP was incubated with 2.5 mM PC/PS (4:1) (lane2) or 2.5 mM PC (lane3) sucrose-loaded vesicles. The vesicles were pelleted by centrifugation, and 25 µl of the supernatant, containing unbound MRP, was loaded on a 10% SDS-polyacrylamide gel. The gel was stained with silver. In a control experiment, the vesicles were omitted (lane1). Upperpanel, unmyr MRP; lowerpanel, myr MRP.




Figure 8: Determination of apparent partition coefficients for the binding of MRP to PC/PS (4:1) vesicles. 1 µM myr (A) and unmyr (B) MRP were incubated with increasing amounts of PC/PS (4:1) vesicles and centrifuged to separate free from membrane-bound protein. The percentage of MRP bound to vesicles was calculated following scanning densitometry of silver-stained SDS-polyacrylamide gels as described under ``Experimental Procedures'' and is expressed as a function of the accessible lipid concentration. The datapoints from five experiments are shown. The data is fitted according to (see ``Experimental Procedures'') using K(p) values of 2 10^3M for unmyr MRP and 5 10^3M (brokencurve) and 8 10^3M (plaincurve) for myr MRP.



Hydrophobic Photolabeling of MRP and MARCKS

Although the myristoyl moiety of proteins is believed to insert in the bilayer, no direct proof of such an interaction has been presented so far. To address this question, we have labeled two proteins of the MARCKS family, namely mouse MRP and bovine brain MARCKS with the photoreagent (I)-TID-PC/16 (Weber et al., 1994, Weber and Brunner, 1995) (Fig. 9). This reagent is a phospholipid analogue and can thus be incorporated into bilayers. Upon photolysis, the diazirine group is activated and generates an extremely reactive carbene. The carbene reacts with its immediate surroundings, which include mostly lipids but also the membrane-bound apolar core of proteins. The radioactive iodine allows detection of the protein-bound label and consequently a determination of the membrane topology of the protein of interest.


Figure 9: Structure of (I)-TID-PC/16.



Fig. 10shows the results of the labeling experiments with MARCKS and MRP. In the presence of PC/PS (4:1) vesicles, myr MARCKS and MRP are heavily labeled. In the absence of acidic lipids, i.e. with PC vesicles, the extent of labeling of myr MARCKS and MRP is decreased more than 10-fold. When unmyr MARCKS and MRP are photolyzed in the presence of PC/PS (4:1) vesicles (arrowheads), only a weak labeling is observed. This labeling is, however significantly over control values; we have estimated that unmyr MRP incorporates as much as 10-15% of the radioactivity compared with the myristoylated protein. In order to exclude the possibility that the radioactivity associated with the proteins results from noncovalent binding of unreacted reagent, vesicles containing (I)-TID-PC/16 were photolyzed prior to incubation with unmyr MARCKS or MRP. These controls, which are shown for the myristoylated proteins on the rightpanel, demonstrate that noncovalently bound reagent is almost completely removed and consequently does not contribute to the labeling observed when the solutions are photolized subsequent to incubation of the proteins with the vesicles.


Figure 10: Hydrophobic photolabeling of MRP and MARCKS. unmyr as well as myr MRP and MARCKS were incubated with 100-nm vesicles containing (I)-TID-PC/16. The vesicles consisted of either PC or PC/PS (4:1). Following photolysis of the protein-vesicle solutions, lipids were removed on DE52 gel, and the protein-bound label was detected on a 10% SDS-polyacrylamide gel. The arrowheads indicate the position of the unmyristoylated proteins. A, Coomassie Blue staining; B, autoradiography.



The weak labeling of the unmyristoylated proteins strongly suggests that the major site of labeling with (I)-TID-PC/16 is the myristoyl moiety. To confirm this hypothesis we have used the ability of cell extracts to enzymatically demyristoylate MARCKS (Manenti et al., 1994). myr MARCKS and MRP were labeled with (I)-TID-PC/16, delipidated on DE52 gel, and finally demyristoylated with a cytosolic fraction of brain synaptosomes (Fig. 11). Demyristoylation can easily be checked with MARCKS since removal of the fatty acid shifts the apparent molecular mass of the protein from 80 to 70 kDa (Fig. 11A) (Manenti et al., 1994). Since the 70-kDa band (arrowhead) contains <5% of the radioactivity originally associated with the intact protein, we conclude that (I)-TID-PC/16 is mostly bound to the myristoyl moiety of MARCKS (Fig. 11B). The demyristoylation experiments with MRP follow essentially the same pattern. Upon incubation with the synaptosomal extract, MRP is shifted to an apparent higher molecular weight on SDS-polyacrylamide gel (panelA), suggesting that the protein is demyristoylated (see Fig. 2and Fig. 4). Most of the radioactivity originally associated with the protein is removed following demyristoylation (panelB). The fraction of label still associated with the protein after demyristoylation (about 30%) is, however, significantly higher than for MARCKS. Because of the relative minor difference in apparent molecular weights between myr and unmyr MRP (see Fig. 2and Fig. 4) it is unclear whether the label remaining associated with the protein reflects an incomplete removal of the myristoyl moiety or results from the labeling of other parts of the protein. Finally, upon demyristoylation of MARCKS and MRP photolyzed in the presence of PC vesicles, the low amount of label associated with the proteins is removed (Fig. 11).


Figure 11: Demyristoylation of photolabeled MRP and MARCKS. Following photolysis of vesicles incubated with myr MRP and MARCKS, the solutions were delipidated. Myr MARCKS and MRP were demyristoylated enzymatically with cytosolic extracts of brain synaptosomes and subsequently loaded on a 10% SDS-polyacrylamide gel. Demyristoylation was assessed by analyzing the apparent molecular weight of the proteins on a Coomassie Blue-stained 10% SDS-polyacrylamide gel (A). (I)-TID-PC/16 remaining associated with MARCKS and MRP was detected by autoradiography (B). The arrowheads indicate the position of the unmyristoylated proteins.



Interestingly, the labeling of MARCKS with (I)-TID-PC/16 does not change the apparent molecular mass of the protein, whereas addition of the myristoyl moiety increases the apparent molecular mass from 70 to 80 kDa (Fig. 11). These observations indicate that, in contrast to the phospholipid analogue (I)-TID-PC/16, the myristoyl moiety might specifically alter the conformation of MARCKS and consequently its affinity for SDS. Another intriguing observation is that the myristoyl moiety of MARCKS can still be removed following labeling with (I)-TID-PC/16. The putative ``demyristoylase'' therefore most likely does not specifically recognize the myristoyl moiety of MARCKS. The information contained at the N terminus of MARCKS is thus likely to be important for enzymatic demyristoylation.


DISCUSSION

In order to investigate how MRP interacts with membranes we have compared the binding of unmyr and myr MRP to vesicles of different lipid compositions. As a source of proteins we have used an E. coli system in which myr MRP can be produced by coexpressing yeast NMT (Blackshear et al., 1992). Both unmyr and myr MRP were expressed, purified to homogeneity under mild conditions, and characterized. We have shown that both unmyr and myr MRP bind dansyl-calmodulin and are phosphorylated by the catalytic fragment of protein kinase C. Whether myristoylation modulates the interactions of MRP with calmodulin and protein kinase C is currently under investigation and will be reported elsewhere.

The role of the basic domain was analyzed by changing the lipid composition of the vesicles and introducing acidic lipids to mimic the composition of the plasma membrane (Kim et al., 1994a, 1994b). Both unmyr and myr MRP require acidic lipids for their binding, in agreement with the model proposed for MARCKS that predicts that the basic domain interacts electrostatically with acidic lipid membranes (Taniguchi and Manenti, 1993; Kim et al., 1994a, 1994b). The partition coefficient of unmyr MRP in the presence of PC/PS (4:1) vesicles is, however, 3 orders of magnitude smaller than for the peptide corresponding to the basic domain of MARCKS (K(p) = 4 10^6M) (Kim et al., 1994a). This large difference might result from a steric hindering of the basic domain in the protein. Alternatively, negative charges, close to the basic domain, could inhibit the binding of MRP to acidic lipid membranes via electrostatic repulsion, as suggested for MARCKS (Kim et al., 1994b). A third explanation might also be found in the structure of the basic domain; MRP contains a proline at position 96 instead of a serine in other MARCKS proteins (see Fig. 1). Since the basic domain has been hypothesized to be an amphipathic alpha-helix (Graff et al., 1989), this proline might alter the structure of the helix by introducing either a 30-degree kink or a turn (Richardson and Richardson, 1989) and potentially decrease the partition coefficient of MRP. This hypothesis is supported by the observation that unmyr MARCKS has a 50-fold higher affinity for PC/PS (4:1) vesicles (K(p) = 1.3 10^5M) (Kim et al., 1994b).

Expression of myristoylation-deficient mutant proteins shows that the myristoyl moiety is necessary to promote the binding of otherwise acylated proteins to membranes (see Introduction). Our results show that myristoylation is clearly not sufficient for membrane binding since we have estimated that the apparent partition coefficient of myr MRP in the presence of PC vesicles is smaller than 10^2M. The affinity of myr MRP for neutral membranes is therefore at least 2 orders of magnitude smaller than for myristoylated model peptides, which have partition coefficients of 10^4M (Peitzsch and McLaughlin, 1993). This observation suggests that the myristoyl moiety of MRP is in a conformation that inhibits its insertion into neutral membranes. Furthermore, in the presence of acidic lipid membranes, the myristoyl moiety only increases the binding of MRP to PC/PS (4:1) vesicles by a factor of 2-4. In contrast to MARCKS (Kim et al., 1994b), a strong cooperative effect between the basic domain and the myristoyl moiety is not observed, and we therefore propose that the major determinant of the binding of MRP to pure lipid membranes is the basic domain rather than the myristoyl moiety. Interestingly, the conclusion that the myristoyl moiety of a protein is not required for its binding to phospholipid vesicles has also been proposed for the ADP-ribosylation factor ARF1 (Franco et al., 1993) and is likely to be extended to other proteins.

Fluorescent fatty acid labels incorporated into the palmitoylation sites of rhodopsin have demonstrated that the palmitoyl moiety of this protein is situated in the membrane (Moench et al., 1994). In spite of the considerable recent interest in myristoylated proteins, a direct demonstration of such a topology for myristoyl moieties has not been presented so far. In order to investigate this aspect, we have photolabeled MRP as well as MARCKS with (I)-TID-PC/16. The observations that the myristoyl moiety is required to obtain a strong labeling and that demyristoylation results in a loss of the label, demonstrate that the myristoyl moiety of MRP and MARCKS is incorporated in the bilayer. This represents to our knowledge the first direct evidence that the fatty acid moiety of myristoylated proteins can insert into phospholipid bilayers. In the presence of PC vesicles, the labeling of myr MRP and MARCKS is drastically reduced, demonstrating that electrostatic interactions are required to promote the insertion of the myristoyl moiety into the membrane.

A low but nonetheless significant labeling can be observed when unmyr MRP and MARCKS are photolized in the presence of PC/PS (4:1) vesicles. This observation indicates that secondary hydrophobic interactions might also be involved in membrane binding. The basic domain, which can be modeled as an amphipathic alpha-helix, contains 6 hydrophobic residues, which are clustered in two domains of the putative alpha-helix (Graff et al., 1989). These residues could potentially insert in the hydrophobic bilayer and participate in the binding. In this respect, replacement of Phe and Phe (Phe and Phe in the MRP sequence) by alanines results in a twofold decrease in the binding of the basic domain of MARCKS to membranes (Kim et al., 1994a). In analogy to MARCKS proteins, the N-terminal domain of the ADP-ribosylation factor ARF1 also contains a repeat of hydrophobic residues, IFXXLFXXF, whose structure is compatible with that of an amphipathic helix and which might be critical to the membrane attachment of that protein (Kahn et al., 1992).

myr MRP binds to PC/PS (4:1) vesicles with a partition coefficient of 5-8 10^3M which is 20-30-fold smaller than for MARCKS (Kim et al., 1994b). Whereas the affinity of MARCKS for PC/PS (4:1) vesicles is high enough to account for a stable binding of the protein at the plasma membrane (Kim et al., 1994b), the partition coefficient of MRP is clearly insufficient. Since myristoylation does not drastically increase the binding of MRP to membranes, the cooperative model proposed for MARCKS (Kim et al., 1994b) cannot be applied to MRP. Based on our finding, as well as on the conclusions obtained for MARCKS by McLaughlin and co-workers (Peitzsch and McLaughlin, 1993; Kim et al., 1994a, 1994b), we propose a model for the binding of MRP to membranes (Fig. 12). In solution, the myristoyl moiety of MRP is in a close conformation. The basic domain targets MRP to a negatively charged membrane, such as the plasma membrane. This interaction is too weak to allow a stable anchoring of the protein but allows an efficient insertion of the myristoyl moiety into the bilayer. This particular conformation is then recognized by a membrane protein, which in turn provides the additional binding energy to firmly anchor MRP at the membrane.


Figure 12: Putative model for the binding of MRP to acidic lipid membranes.




FOOTNOTES

*
This research was supported by Swiss National Foundation for Promotion of Science Grants 31-32188.91 (to G. Schwarz, Biocenter, Basel) and 31-36193.92 (to J. Brunner, Swiss Federal Institute of Technology, Zürich). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Dept. of Biophysical Chemistry, Biocenter of the University of Basel, Klingelbergstrasse 70, 4056 Basel, Switzerland. Tel: 41-612672179; Fax: 41-612672189; vergeres{at}urz.unibas.ch.

Present address: Program of Cellular Biochemistry and Biophysics, Rockefeller Research Laboratories, Memorial Sloan-Kettering Cancer Center, New York, NY 10021.

**
Present address: Hopital PURPAN, Unité INSERM 326, 31059 Toulouse, France.

(^1)
The abbreviations used are: NMT, N-myristoyl transferase; IPTG, isopropyl-1-thio-beta-D-galactopyranoside; (I)-TID-PC/16, 1-O-hexadecanoyl-2-O-[9-[[[2-(I)iodo-4-((trifluoromethyl)-^3H-diazirin-3-yl)benzyl]oxy]carbonyl]nonanoyl]-sn-glycero-3-phosphocholine; MARCKS, myristoylated alanine-rich protein kinase C substrate; MRP, MARCKS-related protein; myr, myristoylated; PC, phosphatidylcholine; PIC, protease inhibitor mixture; PMSF, phenylmethysulfonyl fluoride; PKM, catalytic domain of protein kinase C; PS, phosphatidylserine; unmyr, unmyristoylated.

(^2)
A. Lustig and G. Vergères, unpublished results.

(^3)
G. Vergères, unpublished results.


ACKNOWLEDGEMENTS

We thank Josef Brunner for stimulating discussions. We are also grateful to Jeffrey Gordon and Perry Blackshear for giving us the plasmids pBB131NMT and pET3dF52M1, respectively. Finally we acknowledge Hisaaki Taniguchi for suggesting the use of the phenyl-Sepharose column to purify MRP.


REFERENCES

  1. Aderem, A. A. (1992) Cell 71,713-716 [Medline] [Order article via Infotrieve]
  2. Albert, K. A., Walaas, S. I., Wang, J. K. T., and Greengard, P. (1986) Proc. Natl. Acad. Sci. U. S. A. 83,2822-2826 [Abstract]
  3. Allen, L. A. H., and Aderem, A. A. (1995) EMBO J. 14,1109-1121 [Abstract]
  4. Blackshear, P. J. (1993) J. Biol. Chem. 268,1501-1504 [Free Full Text]
  5. Blackshear, P. J., Verghese, G. M., Johnson, J. D., Haupt, D. M., and Stumpo, D. J. (1992) J. Biol. Chem. 267,13540-13546 [Abstract/Free Full Text]
  6. Blenis, J., and Resh, M. D. (1992) Cur. Opin. Cell Biol. 119,984-989
  7. Böttscher, C. J. F., van Gent, C. M., and Fries, C. (1961) Anal. Chim. Acta 24,203-204 [CrossRef]
  8. Busconi, L., and Michel, T. (1994) J. Biol. Chem. 269,25016-25020 [Abstract/Free Full Text]
  9. Casey, P. J. (1994) Cur. Opin. Cell Biol. 6,219-225 [Medline] [Order article via Infotrieve]
  10. da Silva, A. M., and Klein, C. (1990) J. Cell Biol. 111,401-407 [Abstract]
  11. David-Pfeuty, T., Bagrodia, S., and Shalloway, D. (1993) J. Cell Sci. 105,613-628 [Abstract/Free Full Text]
  12. Duronio, R. J., Jackson-Machelski, E., Heuckeroth, R. O., Olins, P. O., Devine, C. S., Yonemoto, W., Slice, L. W., Taylor, S. S., and Gordon, J. I. (1990) Proc. Natl. Acad. Sci. U. S. A. 87,1506-1510 [Abstract]
  13. Franco, M., Chardin, P., Chabre, M., and Paris, S. (1993) J. Biol. Chem. 268,24531-24534 [Abstract/Free Full Text]
  14. Franco, M., Chardin, P., Chabre, M., and Paris, S. (1995) J. Biol. Chem. 270,1337-1341 [Abstract/Free Full Text]
  15. Gallego, C., Gupta, S. K., Winitz, S., Eisfelder, B. J., and Johnson, G. L. (1992) Proc. Natl. Acad. Sci. U. S. A. 89,9695-9699 [Abstract]
  16. George, D. J., and Blackshear, P. J. (1992) J. Biol. Chem. 267,24879-24885 [Abstract/Free Full Text]
  17. Graff, J. M., Gordon, J. I., and Blackshear, P. J. (1989) Science 246,503-506 [Medline] [Order article via Infotrieve]
  18. Hallak, H., Brass, L. F., and Manning, D. R. (1994) J. Biol. Chem. 269,4571-4576 [Abstract/Free Full Text]
  19. Harris, M. P. G., and Neil, J. C. (1994) J. Mol. Biol. 241,136-142 [CrossRef][Medline] [Order article via Infotrieve]
  20. Hope, M. J., Bally, M. B., Webb, G., and Cullis, P. R. (1985) Biochim. Biophys. Acta 812,55-65
  21. Huang, M., and Jolicoeur, P. (1994) J. Virol. 68,5648-5655 [Abstract]
  22. James, G., and Olson, E. N. (1989) J. Biol. Chem. 264,20928-20933 [Abstract/Free Full Text]
  23. Johnson, J. D., and Wittenauer, L. A. (1983) Biochem. J. 211,473-479 [Medline] [Order article via Infotrieve]
  24. Johnson, D. R., Bhatnagar, R. S., Knoll, L. J., and Gordon, J. I. (1994) Annu. Rev. Biochem. 63,869-914 [CrossRef][Medline] [Order article via Infotrieve]
  25. Kahn, R. A., Randazzo, P., Serafini, T., Weiss, O., Rulka, C., Clark, J., Amherdt, M., Roller, P., Orci, L., and Rothman, J. E. (1992) J. Biol. Chem. 267,13039-13046 [Abstract/Free Full Text]
  26. Kamps, M. P., Buss, J. E., and Sefton, B. M. (1985) Proc. Natl. Acad. Sci. U. S. A. 82,4625-4628 [Abstract]
  27. Kim, J., Blackshear, P. J., Johnson, J. D., and McLaughlin, S. (1994a) Biophys. J. 67,227-237 [Abstract]
  28. Kim, J., Shishido, T., Jiang, X., Aderem, A. A., and McLaughlin, S. (1994b) J. Biol. Chem. 269,28214-28219 [Abstract/Free Full Text]
  29. Li, J., and Aderem, A. A. (1992) Cell 70,791-801 [Medline] [Order article via Infotrieve]
  30. Linder, M. E., Pang, I. H., Duronio, R. J., Gordon, J. I., Sternweis, P. C., and Gilman, A. G. (1991) J. Biol. Chem. 266,4654-4659 [Abstract/Free Full Text]
  31. Lobach, D. F., Rochelle, J. M., Watson, M. L., Seldin, M. F., and Blackshear, P. J. (1993) Genomics 17,194-204 [CrossRef][Medline] [Order article via Infotrieve]
  32. Lowry, O., Rosebrough, N., Farr, A., and Randall, R. (1951) J. Biol. Chem. 193,265-275 [Free Full Text]
  33. Magee, A. I., and Courtneidge, S. A. (1985) EMBO J. 4,1137-1144 [Abstract]
  34. Manenti, S., Sorokine, O., Van Dorsselaer, A., and Taniguchi, H. (1992) J. Biol. Chem. 267,22310-22315 [Abstract/Free Full Text]
  35. Manenti, S., Sorokine, O., Van Dorsselaer, A., and Taniguchi, H. (1993) J. Biol. Chem. 268,6878-6881 [Abstract/Free Full Text]
  36. Manenti, S., Sorokine, O., Van Dorsselaer, A., and Taniguchi, H. (1994) J. Biol. Chem. 269,8309-8313 [Abstract/Free Full Text]
  37. Moench, S. J., Moreland, J., Steward, D. H., and Dewey, T. G. (1994) Biochemistry 17,5791- 5796
  38. Nadler, M. J. S., Harrisson, M. L., Ashendel, C. L., Cassady, J. M., and Geahlen, R. L. (1993) Biochemistry 32,9250-9255 [Medline] [Order article via Infotrieve]
  39. Op den Kamp, J. A. F. (1981) Membrane Structure (Finean, J. B., and Michell, R. H., eds) pp. 83-126, Elesevier, Amsterdam
  40. Peitzsch, R. M., and McLaughlin, S. (1993) Biochemistry 32,10436-10443 [Medline] [Order article via Infotrieve]
  41. Resh, M. D. (1994) Cell 76,411-413 [Medline] [Order article via Infotrieve]
  42. Richardson, J. S., and Richardson, D. C. (1989) Prediction of Protein Structure and the Principles of Protein Conformation (Fasman, G. D., ed) pp. 1-98, Plenum Press, New York
  43. Rose, S. D., Cook, H. W., Palmer, F. B., Ridgway, N. O., and Byers, D. M. (1994) J. Neurochem. 63,2314-2323 [Medline] [Order article via Infotrieve]
  44. Sigal, C. T., and Resh, M. D. (1993) Mol. Cell. Biol. 13,3084-3092 [Abstract]
  45. Sigal, C. T., Zhou, W., Buser, C. A., McLaughlin, S., and Resh, M. D. (1994) Proc. Natl. Acad. Sci. U. S. A. 91,12253-12257 [Abstract/Free Full Text]
  46. Taniguchi, H., and Manenti, S. (1993) J. Biol. Chem. 268,9960-9963 [Abstract/Free Full Text]
  47. Thelen, M., Rosen, A., Nairn, A. C., and Aderem, A. A. (1991) Nature 351,320-322 [CrossRef][Medline] [Order article via Infotrieve]
  48. Uchida, T., and Filburn, C. R. (1984) J. Biol. Chem. 259,12311-12314 [Abstract/Free Full Text]
  49. Umekage, T., and Kato, K. (1991) FEBS 286,147-151 [CrossRef][Medline] [Order article via Infotrieve]
  50. Verg è res, G., Manenti, S., and Weber, T. (1995) Signaling Mechanisms: From Transcription Factors to Oxidative Stress (Packer, L., and Wirtz, K. W. A., eds) Vol. H92, pp. 125-137, Springer Verlag, Berlin, Heidelberg
  51. Verghese, G. M., Johnson, J. D., Vasulka, C., Haupt, D. M., Stumpo, D. J., and Blackshear, P. J. (1994) J. Biol. Chem. 269,9361-9367 [Abstract/Free Full Text]
  52. Weber, T., and Brunner, J. (1995) J. Am. Chem. Soc. 117,3084-3095
  53. Weber, T., Paesold, G., Galli, C., Mischler, R., Semenza, G., and Brunner, J. (1994) J. Biol. Chem. 269,18353-18358 [Abstract/Free Full Text]
  54. Yonemoto, W., McGlone, M. L., and Taylor, S. S. (1993) J. Biol. Chem. 268,2348-2352 [Abstract/Free Full Text]
  55. Yu, G., and Felsted, R. L. (1992) Virology 187,46-55 [Medline] [Order article via Infotrieve]

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