extfieldsofstudy: Medicine Biology

text:
A v-SNARE implicated in intra-Golgi transport.

We report the identification of a putative v-SNARE (GOS-28), localized primarily to transport vesicles at the terminal rims of Golgi stacks. In vitro, GOS-28, A Golgi SNARE of 28 kD, is efficiently packaged into Golgi-derived vesicles, which are most likely COPI coated. Antibodies directed against GOS-28 block its ability to bind alpha-SNAP, partially inhibit transport from the cis to the medial cisternae, and do not inhibit budding of COP-coated vesicles, but do accumulate docked uncoated vesicles.

T RANSPORT vesicles shuttling cargo between intracellular organelles dock to their correct destination by the selective recognition of an address signal on vesicles by a receptor on the intended target membrane. According to the SNARE 1 hypothesis, this process is mediated at least in part by the unique pairing of SNAP receptors localized on vesicles (v-SNAREs) with their cognate t-SNARE on the target membrane (Srllner et al., 1993a). For example, in yeast, distinct compartment-specific SNARE complexes acting at the ER-Golgi (S,gaard et al., 1994) and at the Golgi-plasma membrane transport steps (Aalto et al., 1992(Aalto et al., , 1993Protopopov et al., 1993) have been demonstrated and a t-SNARE at the vacuole has been postulated (Bennett et al., 1993). In mammalian cells, SNARE complexes involved in regulated exocytosis have been isolated (Srllner et al., 1993a,b). The only known t-SNARE acting early in the secretory pathway is syntaxin 5 (Bennett et al., 1993) and its yeast homologue, Sed5 (Hardwick and Pelham, 1992), which have been localized to the cis-Golgi network (Banfield et al., 1994). In both eukaryotes, the SNARE pattern within the Golgi apparatus is largely unknown.
In contrast to SNARE proteins, which have to be compartment specific to fulfill their proposed function, the machinery acting in the consumption of docked vesicles, in the fusion process, is a general one used at multiple transport steps. Indeed, general fusion proteins such as the Please address all correspondence to Thomas Srllner, Cellular Biochemistry and Biophysics Program, Memorial Sloan-Kettering Cancer Center, 1275 York Avenue, Box 251, New York, NY 10021. Tel.: (212) 639-8598; Fax: (212) 717-3604.
1. Abbreviations used in this paper: ARF, ADP-ribosylation factor; GOS-28, Golgi SNARE of 28 kD; GST, glutathione S-transferase; NEM, N-ethylmaleimide; NSF, N-ethylmaleimide-sensitive fusion protein; SNAP, soluble NSF attachment protein; SNARE, SNAP receptor; VSV, vesicular stomatitis virus. N-ethylmaleimide-sensitive fusion protein (NSF)  and soluble NSF attachment proteins (SNAPs)  can be both found free in the cytoplasm and associated with the membranes of different organelles. The membrane interaction of NSF is mediated by the binding of SNAPs to SNAREs (Weidmann et al., 1989;Wilson et al., 1992). The assembled v-SNARE/t-SNARE complex provides high affinity sites for the binding of several SNAPs (Srllner et al., 1993b;Hayashi et al., 1995;Schiavo et al., 1995). A complex of SNAREs, SNAPs, and NSF can be isolated in vitro in the absence of hydrolyzable ATP, and migrates with 20 Svedberg (20S particle) in velocity centrifugation (Wilson et al., 1992). In the presence of ATP, the ATPase activity of NSF provides energy to dissociate the SNARE complex and to release SNAPs and NSF, thereby likely linking the process of vesicle docking to the initiation of vesicle fusion (Srllner et al., 1993b).
The secretory pathway is the major route for the delivery of newly synthesized molecules to their intracellular or extracellular destinations. This requires the maintenance of the compartments comprising the pathway (i.e., the constancy of their protein and lipid composition), despite a continuous flow of material passing through them. This appears to be accomplished by constant sorting events either excluding or enriching certain molecules in transport vesicles (Griffiths and Simons, 1986;Pelham and Munro, 1993). Transport vesicles enriched in certain cargo could either move in an anterograde or retrograde direction, either selecting molecules for their forward transport to a distinct organelle or retrieving escaped molecules backwards to their organelle of origin (Pelham, 1991). In addition to these sorting events, import into and export from a distinct compartment have to be balanced to maintain its size and volume. This implies that the transport containers, the vesicles, and the transport machinery have to be recycled, v-SNAREs that determine the address of a trans-port vesicle are themselves part of this machinery and may play a major role in these processes. For example, a v-SNARE that has guided a vesicle to its destination becomes part of the target membrane after fusion has occurred. Most likely, this v-SNARE has to be cycled back to its compartment of origin, and it would seem only natural that the number of v-SNAREs available for forward transport is dependent on their recycling from their target membrane. This would couple and balance the vesicle flow between two compartments. To test such predictions, it is necessary to identify v-SNAREs and their interacting t-SNAREs in the central turntable for vesicular trafficking, the Golgi apparatus, and its cisand trans-Golgi networks.
On one hand, the Golgi receives both anterograde transport vesicles from the ER and retrograde vesicles from downstream compartments; on the other hand, it produces retrograde vesicles moving back to the ER and anterograde vesicles moving toward the plasma membrane (Pelham, 1991;Miesenb6ck and Rothman, 1995). Therefore, the isolation and characterization of SNAREs from the Golgi will be essential for our understanding of how a cell maintains the integrity of its organelles by integrating vesicular flow patterns in a coordinated manner. Furthermore, the localization of SNAREs within the Golgi stack might provide us with information about the organization of the Golgi. For example, analysis of the distribution of SNAREs within a cisterna may help to define areas involved in vesicle budding and fusion.
In this paper, we describe the isolation of Golgi SNAREs and the identification, localization, and functional characterization of GOS-28, a v-SNARE, which can act in intra-Golgi transport.

Sequencing and Mass Analysis of Peptides
Proteins separated by SDS-PAGE were electroblotted onto nitrocellulose, and the visualized bands were processed for internal amino acid analysis, as described . Briefly, membrane-bound proteins were digested in situ with trypsin and the resulting peptides separated by narrowbore reverse-phase HPLC; the modular liquid chromatography system used in this study has been described elsewhere . Selected peptides were then subjected to chemical microsequencing and matrix-assisted laser desorption (MALDI) mass spectrometry, also described previously (S611ner et al., 1993a;Erdjument-Bromage et al., 1994;Tempst et al., 1994).

Isolation of GOS-28 cDNA
To isolate a cDNA encoding Golgi-SNARE of 28 kD (GOS-28), a twostage PCR was performed with degenerate primers based on peptide sequence data obtained as described (S611ner et al., 1993a). For the first stage of PCR, the following primers were used: ATGTT(TC)GA(AG)-AC(AGTC)ATGGC (codons Met66-Ala 71, sense, 0.5 p,M), TG(AG)AA-(TC)TC(AG)TG(AGTC)GT(AG)TA (codons TyrllS-His 123, antisense, 0.5 ~M) and a bovine liver cDNA (0.5 ng, Clontech) as template. The amplification reactions were performed for five cycles using an annealing temperature of 42°C (1 min at 94°C, 1 min at 42°C, and 2 min at 72°C) fol-lowed by an additional 30 cycles at an annealing temperature of 55°C (1 min at 94°C, 1 min at 55°C, and 2 min at 72°C). The last cycle was followed by an 8-min extension at 72°C. For the second stage, the PCR products from the first stage were diluted 1:1,000, and the same antisense primer as in stage 1 and the ATGGC(AGTC)AT(ATC)GA(AG)AT (ATC)GA (codons Met7°-Glu 75, sense) primer were added. The amplification conditions were the same as in stage 1. The products were fractionated by electrophoresis in 2% agarose gels. A major amplification product of ~180 bp was isolated, random labeled, and used as a probe for screening a CHO cDNA library constructed with the Uni-ZAP XR vector (Stratagene, La Jolla, CA). Hybridization was done at 50°C for 14 h in 5 x SSPE (1 × SSPE = 0.18 M NaC1, i mM EDTA, 10 mM NaH2PO4, pH 7.7), 5× Denhardt's solution, 0.5% SDS, in the presence of 20 Ixg/ml herring sperm DNA. The filters were washed twice in 2x SSPE, 0.1% SDS for 10 min at room temperature, followed by a 15-min wash at 50°C in 1× SSPE, 0.1% SDS. Positive clones were excised from the Uni-ZAP XR vector as inserts in pBluescript SK-by coinfection with the helper phage, ExAssist, as described by the manufacturer (Stratagene), and were sequenced for both sense and antisense strands.

Preparation of Antibodies
To generate a polyclonal antibody to GOS-28, a fusion protein of GOS-28 with an amino-terminal extension containing six histidine residues (His 6-GOS-28) was prepared by insertion of the GOS-28 cDNA into a pQE11 vector (Qiagen, Chatsworth, CA). After expression in Escherichia coli (XL1-Blue; Stratagene), the protein was purified from inclusion bodies under denaturing conditions by affinity chromatography using Ni-nitrilo-triacetic acid agarose as described by the manufacturer (Qiagen). His6-GOS-28 was eluted in 8 M urea, 0.1 M NaH2PO4, and 0.01 M Tris (pH 4.5), and then injected into rabbits for immunization. Antibodies directed against GOS-28 were purified as described (Harlow and Lane, 1988), by affinity chromatography using His6--CJOS-28 conjugated to cyanogen bromideactivated Sepharose 4B (Pharmacia Fine Chemicals, Piscataway, N J), and the elutate was dialyzed against PBS. Minor contaminations of antibodies cross-reacting with cellular antigens other than GOS-28 were affinity depleted. For this purpose, 500 Ixg of CHO protein was separated by SDS-PAGE, transferred onto nitrocellulose membrane, the region containing proteins with a 28-kD tool mass was cut out, and the remaining nitrocellulose membrane was incubated with the affinity-purified anti-GOS-28 antibodies in 0.1% BSA.
Fab fragments were prepared by incubating the affinity-purified anti-GOS-28 antibody with papain (Boehringer Mannheim Biochemicals, Indianapolis, IN), and any remaining intact antibodies and Fc fragments were removed by protein A-agarose chromatography (Mage, 1981). The Fab fragments were dialyzed against 10 mM MOPS-KOH (pH 7.5) and 150 mM KCI.

Expression of Epitope-tagged GOS-28 in CHO Cells
To obtain CHO cells that stably express GOS-28 tagged with a myc epitope (EQKLISEEDL), the coding sequence for the myc epitope was added by PCR to the 5' end of the coding region of the GOS-28 cDNA, and the resulting construct was subcloned into the mammalian cell expression vector pcDNA3 (InVitrogen, San Diego, CA). The FLAG epitope (DYKDDDDK) was inserted after the initiation codon into the GOS-28 cDNA by loop-in mutagenesis using the unique site elimination method (Deng and Nickoloff, 1992) and the Transformer site-directed mutagenesis kit (Clontech Laboratories, Palo Alto, CA). CHO cells grown in MEM-ct medium containing 10% FCS were transfected with the abovementioned constructs using the Lipofectamine reagent (GIBCO-BRL, Gaithersburg, MD), as described by the manufacturer. 48 h after the transfection, the cells were selected by the incubation with 1 mg/ml G418 sulfate (GIBCO-BRL). After 7 d, several resistant colonies were isolated by trypsinization in cloning cups, grown in a medium containing 0.5 mg/ml G418, and examined for myc-GOS-28 or FLAG--GOS-28 expression by Western blot analysis.

Immunolocalization
For immunofiuorescence, CHO cells expressing FLAG-and myc-tagged GOS-28 fusion proteins were fixed with 4% paraformaldehyde, and were permeabilized by dehydration and rehydration with ethanol. Cells were subsequently exposed for 2 h at room temperature to affinity-purified anti-GOS-28 antibodies (1:5) followed by a 1-h exposure to goat anti-rabbit IgG coupled to FITC. Anti-FLAG (M2) (Brizzard et al., 1994) or antimyc (9El0) (Evan et al., 1985) mAb revealed by FlTC-coupled antimouse IgG were used to localize FLAG-or myc-tagged GOS-28 in the transfected CHO cell lines. Cells were counterstained with 0.03% Evans blue and examined with a confocal fluorescence microscope. For electron microscope immunolocalization, cells were fixed with 2% glutaraldehyde in phosphate buffer (pH 7.4), infiltrated with 2.3 M sucrose, and processed for cryoultramicrotomy according to the method of Tokuyasu (1986). Anti-GOS-28 affinity-purified antibodies (diluted 1:3) were localized with the protein A-gold method (Roth et al., 1978) or by goat anti-rabbit IgGcoupled gold. Anti-myc (9E10) antibodies (diluted 1:100 or 1:300) were localized with rabbit anti-mouse IgG followed by protein A-gold. After immunolabeling, sections were absorption stained with uranyl acetate (Tokuyasu, 1986) and examined in the electron microscope.

Generation of Golgi-derived Coated Vesicles
Vesicle budding reactions contained CHO Golgi membranes (50 I~g/ml), bovine brain cytosol (2.4 mg/ml), 10 IxM palmitoyl Coenzyme A, 25 p.g/ml RNase A, and 20 ~M GTP~/S, in transport buffer. Control reactions were performed in the absence of GTP~/S. Reaction mixtures were assembled on ice and then incubated at 37°C for 15 rain. After the incubation, the Golgi membranes were isolated by centrifugation for 20 min at 15,800 g. The Golgi membrane pellet was resuspended in 0.6 ml low salt stripping buffer (25 mM Hepes [pH 7.2], 2.5 mM Mg(OAc)2, 0.2 M sucrose, and 50 mM KC1), reisolated by centrifugation, and subsequently washed with 0.6 ml high salt stripping buffer (25 mM Hepes [pH 7.2], 25 mM Mg(OAc)2, 0.2 M sucrose, 250 mM KC1). After centrifugation, the high salt supernatant was adjusted to 52% (wt/wt) sucrose. The high salt supernatant containing the coated vesicles was overlayered with 0.5 ml of 45% sucrose and 1 ml of (each) 40, 35, and 30% sucrose, and centrifuged for 15 h at 287,000 g. 350-1~1 fractions were collected from the gradient, diluted with 1 vol H20 , and centrifuged at 356,000 g for 30 min to pellet the vesicles. The isolated vesicles were resuspended in Laemmli buffer for Western blot analysis.

Assembly of 20S Particles on Immobilized GST-a-SNAP
A glutathione S-transferase (GST) fusion protein of a-SNAP was prepared by inserting the a-SNAP eDNA into the pGEX-2T vector (Pharmacia). The GST a-SNAP fusion protein was expressed in E. coli and purified from the bacterial lysate on glutathione-agarose (Sigma Chemical Co.). CHO Golgi membranes were stripped of peripherally associated proteins by incubating with 1 M KCI for 15 min on ice, and then recovered by centrifugation (15,800 g, 15 min, 4°C). The membrane pellet was resuspended in lysis buffer (20 mM Hepes-KOH [pH 7.2], 100 mM KCI, 1 mM DT1 ~, 0,5% Triton X-100, 1% glycerol) and solubilized for 45 min at 4°C with gentle mixing, followed by centrifugation for 45 rain at 100,000 g to remove detergent-insoluble material. The Golgi Triton X-100 extract (25 p~g of protein) was incubated with the indicated amounts of affinity-purified anti--GOS-28 antibody for 1 h at 4°C. Subsequently, the mixtures were incubated with GST-a-SNAP (1 p,g of protein) prebound to glutathione-agarose beads (15 p~l) without or with His6-NSF-myc (5 ~g of protein), in the presence of 0.5 mM ATP and either 2 mM EDTA or 2 mM MgC12, for 1 h at 4°C in a final volume of I ml. Beads were then washed four times with 1 ml of the appropriate buffer, and proteins were eluted from the beads by incubation with 15 p~l 2× Laemmli buffer and then subjected to Western blot analysis.

Electron Microscopy to Examine the Effect of the Anti-GOS-28 Antibody on Vesicular Transport
Reactions (500 Ixl) contained 10 p.M palmitoyl Coenzyme A, bovine brain cytosol (2.4 mg/ml), and CHO Golgi membranes (100 p,g/ml) in transport buffer. For the preparation of N-ethylmaleimide (NEM)-treated Golgi, CHO Golgi membranes were treated with 1 mM NEM for 15 min on ice, followed by an addition of 2 mM DTT and an incubation for 15 rain on ice. Reactions were assembled on ice, preincubated on ice for 15 rain with 5 p,g of affinity-purified anti-GOS-28 antibody and 20 ixM GTP~/S (see Table I), and then incubated at 37°C for 15 rain. The reactions were then treated with 1% glutaraldehyde, the membranes were isolated by centrifugation (15,800 g, 30 min, 4°C) and processed for electron microscopy, and the quantitative evaluation of vesicles and buds was performed as described previously .

Isolation of SNAREs from a Golgi-enriched Preparation
To gain insight into the SNARE composition of the Golgi, we made use of the mammalian Golgi apparatus, which is morphologically well defined by the existence of several stacked cisternae. Therefore, a preparation enriched in Golgi was isolated from bovine liver (Tabas and Kornfeld, 1979). The membranes were extracted at pH 11.5 (Fujiki et al., 1982) to remove peripheral associated proteins and lysed by addition of Triton X-100. From this extract, potential SNARE proteins were isolated by their ability to assemble together with SNAPs and NSF in a 20S particle in the absence of hydrolysable ATP. The whole particle was immobilized on beads containing covalently coupled anti-myc antibodies that bind the myc epitope-tagged NSF in the 20S particle. In a control reaction, the beads were treated with Mg-ATP~S to release proteins that were bound in an Mg 2+ sensitive, ATP hydrolysis-independent manner (Fig. 1,  lane 2). SNAPs and SNAREs were then eluted from the matrix by Mg-ATP treatment (S611ner et al., 1993a). In addition to several proteins in the molecular mass range of 70 kD, which are also visible in the control reaction, or-SNAP and six additional protein bands representing potential SNAREs were specifically eluted as shown in the Ponceau S-stained Western blot in Fig. 1, lane 1. These six proteins migrate with an apparent molecular mass of 45 (A), 31 (B), 26 (C), 24 (D), 21 (E), and 20 kD (F) in high-Tris urea SDS-PAGE. Microsequencing and mass spectrometric data obtained from several tryptic peptides derived from these six proteins were used to search protein sequence repositories and indicated that all bands corresponded to as yet unidentified proteins (data not shown). Further analy-  branes. SNARES were purified as described (S6Uner et al., 1993a). Briefly, I--Iis6-NSF-myc (500 ~g of protein) and t-Iis6--a-SNAP (500 0~g of protein) were assembled into 20S particles by incubating them together with Triton X-100 extracts of enriched Golgi membranes (250 mg of protein) from bovine liver in the presence of ATP~/S. The 20S particles were then isolated with anti-myc mAbs covalently coupled to protein G-Sepharose beads (2.5 ml of 2 mg 9El0 IgGs/mg protein G--Sepharose). The beads were washed, eluted with Mg-ATP3,S, free Mg 2÷ was complexed by addition of EDTA, the ATP~/S was exchanged against ATP, and then the bound 20S particles were disassembled by a specific elution step with Mg-ATP. Shown is a Ponceau S-stained Western blot of the specific (Mg-ATP) eluate and the control eluate (Mg-ATP3,S). The following tryptic peptide sequences (one-letter code) of the protein band D were obtained by microsequencing as described under Materials and Methods: (1)  sis, including the isolation of the encoding cDNA, the topological characterization, the intracellular localization, and functional studies, was performed with GOS-28 (represented by protein band D in Fig. 1).

GOS-28 Is a Type H Membrane Protein with the Bulk of Its Mass Exposed to the Cytoplasm
A full-length clone encoding GOS-28 was isolated from a C H O c D N A library, and the predicted molecular mass (28.5 kD) of the protein encoded by the open reading frame was in close agreement with the apparent molecular size of 24 kD (Fig. 1, protein band D). GOS-28 migrates in a standard Laemmli S D S -P A G E with an apparent molecular mass of 31 kD (data not shown). All six peptide sequences obtained by microsequencing of the protein hand D aligned perfectly with the translation product of the open reading frame (Fig. 2). GOS-28 has no obvious signal peptide, and contains a region (amino acids 63-95) with an inherently high probability to form coiled coil structures. The amino terminal region could also form coiled coil structures, but with lower probability. GOS-28 is further characterized by a carboxy-terminal hydrophobic region CHO cells stably transfected with the FLAG epitope-tagged GOS-28 were lysed in Laemmli buffer, and the proteins were separated by SDS-PAGE, transferred on nitrocellulose, and immunodecorated with affinitypurified anti-GOS-28 antibodies as described in Materials and Methods. The upper band represents the FLAG epitope-tagged GOS-28; the lower band represents the endogenous GOS-28. These stably transfected cells were also used for the immunolocalization studies. (B) GOS-28 is an integral membrane protein. CHO Golgi membranes were incubated either with 1 M NaC1 or 200 mM Na2CO3 (pH 11.3) for 30 min on ice. The membranes were isolated by centrifugation at 356,000 g for 45 rain. The supernatants were precipitated by TCA, and the pellets were analyzed by immunodecoration of Western blots with anti-GOS-28 antibodies, as described in Materials and Methods. (C) The bulk of GOS-28 is exposed to the cytoplasm. CHO Golgi membranes were treated with indicated concentrations of proteinase K (Boehringer Mannheim) at 16°C for 30 min in 20 mM Hepes-KOH (pH 7.2), 25 mM KC1, and 200 mM sucrose. Reactions were stopped by the addition of 10 mM PMSF. Samples were precipitated by TCA and analyzed for GOS-28 and p24 as described in B. presumably acting as a membrane anchor (see Fig. 2). Studies using a polyclonal antibody (Fig. 3 A) raised against recombinant GOS-28, which was expressed in and purified from E. coli, revealed that GOS-28 resists salt and sodium carbonate extraction of the Golgi (see Fig. 3 B). Both criteria are consistent with the properties expected for an in-tegral membrane protein. The bulk of the protein seems to be exposed to the cytoplasm since GOS-28 was degraded by addition of low amounts of protease to isolated Golgi (see Fig. 3 C). A control protein, p24, an integral membrane protein, the bulk of whose mass is localized in the Golgi lumen was protease resistant (Stamnes et al., 1995). Protease treatment of detergent-solubilized Golgi showed that p24 is not inherently protease resistant (data not shown). These data, together with the existence of a single membrane-spanning region at the carboxy terminus of GOS-28, argue for a type II membrane protein. These results also confirm the identity of the isolated clone and GOS-28 obtained by its association with SNAPs and NSF.

GOS-28 Is Localized to the Golgi Region
Since the starting material used for the SNARE isolation was enriched for Golgi membranes, one would predict that the majority of the recovered SNAREs should be localized within the Golgi or representing SNAREs that mediate the docking of anterograde or retrograde vesicles to the Golgi. Immunofluorescence localization experiments were performed to confirm this for GOS-28. Polyclonal antibodies raised against GOS-28 and mAbs directed against the FLAG and myc epitope of amino terminal FLAG-GOS-28 and myc-GOS-28 fusion proteins stably expressed in CHO cells were used for these studies (see Fig. 4). In all three cases, GOS-28 exhibits a clear juxtanuclear staining pattern that is characteristic of Golgi proteins.
Electron microscope immunocytochemistry revealed that GOS-28 is primarily localized to vesicles and buds at the ends of the cisternae, with little labeling of the cisternae themselves both in situ and in cell-free assays (Figs. 5 and 6 A-C). The very low levels of labeling of the central regions of stacked cisternae precluded an assignment of GOS-28 to c/s versus medial versus trans regions. Since this distribution of GOS-28 could be indicative of a functional specialization of the terminal rim of cisternae for vesicle budding and fusion, Golgi complexes were primed in vitro to accumulate coated buds and vesicles, and were further analyzed by electron microscopy (Fig. 6, B and C).
A few labeled vesicles were observed in relationship with the c/s-Golgi pole, and occasionally buds on the transitional elements of the ER were also found labeled (data not shown). A comparable distribution of immunolabeling was observed in insulin ceils of the rat pancreas (not shown).

GOS-28 Is Found in Coated Vesicles
The presence of GOS-28 in Golgi-associated buds and vesicles, as seen by electron microscopy, argues that GOS-28 is a v-SNARE. To confirm this, Golgi-derived transport vesicles were accumulated in vitro by incubating purified Golgi in the presence of cytosol, ATP, and a nonhydrolyzable GTP analogue, GTP~/S. GTP~/S binds to the ADP-ribosylation factor (ARF), thus keeping it in its active conformation, which recruits coatomer to membranes (Melancon et al., 1987;Serafini et al., 1991;Donaldson et al., 1992). The coat proteins deform the membrane, thereby shaping vesicles, which finally pinch off from their donor compartment (Ostermann et al., 1993). Since GTP~/S is, at best, only . Approximately 30% of the Golgi-associated vesicles were labeled. A comparable pattern of labeling of myc-tagged, overexpressed GOS-28 is obtained with the monoelonal anti-myc antibody on another set of cryosections (D and E) from stably transfected CHO cells. The predominant distribution of immunogold particles over the lateral bud/vesicular elements of the Golgi stack is particularly evident in E. Incubation with preimmune serum did not elicit labeling. The asterisk in A indicates the nucleus. In A, B, D, and E, antibodies were detected with protein A--gold (size of particles = 10 rim); in C, antibodies were detected with goat anti-rabbit IgG-gold (size = 15 nm). A, ×79,000; B, ×75,000;, C, ×56,000; D, ×67,000; E, ×62,000. Bar, 0.1 ~m. slowly hydrolyzed, ARF and coatomer are locked on the vesicles, thereby preventing fusion and accumulating COPIcoated vesicles, which differ in their density from the originating compartments. Density equilibrium centrifugation thus separates COPI-coated vesicles produced in this manner from parental Golgi membranes.
Indeed, GOS-28 is shifted into the COPI-coated vesicle peak (marked by C-COP) only when GTP~/S is added (Fig. 7), implying that GOS-28 is a v-SNARE because it is efficiently packed into vesicles. Interestingly, the distribution of GOS-28 only coincides with about half of the C-COP, suggesting that some COPI-coated vesicles contain GOS-28 while others do not. This is consistent with the finding that only 30% of vesicles label for GOS-28 (Fig. 5).
The presence of GOS-28 in COPI-coated vesicles was also shown by electron microscope immunocytochemical analysis of vesicles produced from isolated Golgi stacks by purified ARF and coatomer (Fig. 6 B), and in complete transport reactions inhibited by aluminum fluoride (Fig. 6  C). The cisternal areas labeled with anti-GOS-28 antibodies coincide with those labeled by anti-C-COP antibody (Fig. 6 D), indicating that the terminal rims of the Golgi cisternae are likely to be the active zones of vesicle budding. Although all these data strongly suggest the presence Figure 6. GOS-28 is concentrated in vesicles and buds at the terminal rims of Golgi cisternae. (A) Tangential view of a CHO cell Golgi complex (G) in situ showing the distribution of GOS-28 on peripherally located vesicular elements (arrows) in CHO cells stably transfected with overexpressed FLAG-tagged GOS-28. Immunolabeling was done with affinity-purified anti-GOS-28 antibodies. (B) Cell-free transport reactions. An isolated Golgi complex with cisternal and coated vesicular elements: only the latter shows a distinct GOS-28 immunolabeling on the outer aspect of the vesicle membrane. Golgi fractions were incubated with purified ARF and coatomer in presence of GTP, and were labeled with affinity-purified anti-GOS-28 antibodies. (C) Lateral part of an isolated Golgi complex. The GOS-28 immunolabeling appears restricted to the cisternal tips and associated vesicles. Golgi fractions were incubated in the presence of cytosol and aluminum fluoride, as described by Orci et al. (1989). Samples were labeled with affinity-purified anti-GOS-28 antibodies. (D) Section comparable to that in C, but immunolabeled with anti-I-COP antibodies. The labeling pattern is essentially similar to that with anti-GOS-28 antibodies. The Golgi fractions were incubated in presence of cytosol and GTP~/S, as described by Orci et al. (1989). In all experiments, the protein A-gold labeling technique described in Materials and Methods was used; size of gold particles = 10 nm. A, x90000; B, x65000; C and D, x64000. Bar, 0.1 Ixm.
of GOS-28 in COPI-coated vesicles, we can not exclude that GOS-28 might be also found in a different class of Golgi-derived vesicles.

Anti-GOS-28 Antibodies Inhibit Intra-Golgi Transport by Blocking the Interaction of GOS-28 and a-SNAP
Isolated Golgi membranes were incubated with increasing amounts of affinity-purified IgGs directed against GOS-28. The samples were then incubated under conditions reconstituting transport. Transport was measured by glycosylation of the cargo protein VSV G (incorporation of radiolabeled GIcNAc). Transport was blocked by the antibody in a dose-dependent manner. The maximum inhibition was 40-50%, and was reached by the addition of 0.5 l~g IgG to 5 Ixg Golgi protein in a 50-1xl reaction (see Fig. 8 A). Addition of 1 p~g purified cytoplasmic domain of GOS-28 to a reaction had no effect on transport, but completely prevented the inhibitory effect of 1 p.g anti-GOS-28 IgG (data not shown), thus excluding unspecific effects caused by the addition of IgG. In addition, control IgG prepared from preimmune serum had no significant effect on transport. To avoid potential aggregation of Golgi stacks or potential clustering of GOS-28 on the membranes caused by the bivalent IgGs, Fab fragments were prepared. The addition of Fab fragments also resulted in a partial inhibition of transport, similar to that obtained with the IgG, but the Fab had to be added at three-to fourfold higher concentrations to observe the maximal effect (see Fig. 8 B). This could be explained by an inactivation of the Fabs during their preparation procedure or by the lower avidity of Fabs.
To establish at which particular stage the antibody blocks transport, the reactions were analyzed by electron microscopy. Therefore, transport reactions were either incubated with GTP~/S, known to accumulate coated vesicles (Melancon et al., 1987), or with the anti-GOS-28 antibody to test for any morphological effects caused by the antibody. A combination of anti-GOS-28 antibody and GTP~/S was used to test if the antibody blocks the formation of coated vesicles or acts at a later step. In other reactions, the membranes were treated with NEM to block the NSF activity, thus accumulating docked uncoated vesicles (Malhotra et al., 1988). The control reaction consisted of standard transport conditions. As shown in Table I,  F/gure Z GOS-28 is present in Golgi-derived coated vesicles. Golgi membranes were incubated under transport conditions in the presence or absence of GTI~S, as described in Materials and Methods. Briefly, after the GTP~/S treatment, the membranes were isolated and incubated with buffer containing 250 mM KCI to remove associated vesicles. After centrifugation, the supematant coDtainin~ the vesicles was adjusted to 52% sucrose and overlayed with a 30-52% sucrose gradient. Gradient fractious were subjected to Western blot analysis with anti-X-COP polyclonal antibodies and anti--GOS-28 polydonal antibodies. cular profiles associated with the Golgi by 3.2-fold, similar to the effect of NSF inactivation. The antibody did not inhibit the accumulation of coated vesicles by GTI~S, demonstrating that the anti-GOS-28 antibody inhibits transport at some stage after coated vesicle budding. Therefore, while the anti--GOS-28 antibody does not block the formation of coated vesicles, it clearly blocks the comumption of docked uncoated vesicles before fusion can occur. A putative SNARE protein such as GOS-28 should bind SNAP, particularly when it is assembled in v-t-SNARE complexes. To test this, detergent extracts of Golgi membranes were incubated with an affinity matrix containing immobilized or-SNAP (GST--ct-SNAP fusion protein bound to glutathione-agarose). GOS-28 associated with a-SNAP (Fig. 9, lane 1), implying that GOS-28 can interact directly with a-SNAP or, more likely, that GOS-28 assembled in a v-t-SNARE complex provides high affinity sites for the binding of a-SNAP. This interaction is not a dead-end process because NSF releases the bound GOS-28 from the immobilized a-SNAP in presence of Mg-ATP, a hallmark property of SNAREs (S6Uner et al., 1993b) (Fig. 9, lane 9) Figure 8. Anti-GOS-28 antibodies inhibit c/s-to medial-Golgi transport in vitro. Enriched Golgi fractions were assayed for transport of VSV G protein between c/s-and medial-Golgi compartments, as described (Balch et al., 1984a,b; see also Materials and Methods). The reactions (50 ixl) were preincubated on ice for 1 h with the indicated amounts of preimmune IgG (A), affinity purilied anti--GOS-28 antibody (A), control Fab fragments prepared from preimmune IgG (B), or anti--GOS-28 Fab fragments (B) before incubation at 37"C for I h.
The accumulation of docked uncoated vesicles revealed by electron microscopy would imply that anti-GOS-28 antibody should block fusion, but not docking. Consistent with this, anti--GOS-28 antibody blocked the binding of GOS-28 to a-SNAP in a dose-dependent manner until a maximum inhibition of ~80% was reached by adding 2.5 ~g IgG to 25 ~g Golgi protein (see Fig. 9, lanes 2--4). Preimmune antibodies had no effect on the interaction of GOS-28 with a-SNAP (Fig. 9, lane 5). The anti--GOS-28 IgG/ Golgi ratio needed to obtain maximum inhibition of the GOS-28hx-SNAP interaction is the same as that is needed for maximum inhibition of transport, implying that the in- Untreated or NEM-txeated ClIO Golgi membranes were incubated under tlanstx~ conditions in the presence or absence of alTmity-purified anti-GOS-28 antibodies and GTP'yS, as described in Matedah and Metimds. For electron microscopy, nlemtmmes weze fixed, cullected bY centrifugatio~a, and ProCessed as described . For each of the samples the density of coaled vesicles/buds and uncoated circular profiles m a t e d with the Golgi wexe analyzed in 20-30 separate Golgi areas in each of the three independent expedmen~ Numbe~ are mean values ± SF.M. Figure 9. Anti-GOS-28 antibodies inhibit the assembly of the 20S fusion particle. Triton X-100 extract of CHO Golgi membranes (25 I~g protein) was preincubated with the indicated amounts of anti-GOS-28 antibody for 1 h on ice, as described in Materials and Methods. Then GST--a-SNAP (1 I~g) prebound to glutathione beads was added, and the incubation continued for another hour at 4°C in the absence or presence of 5 p.g His~-NSFmyc and either ATP/EDTA or ATP/Mg 2+ as indicated. Proteins bound to the beads were isolated, the samples were washed, and the proteins were further analyzed by SDS-PAGE, transferred to nitrocellulose membranes, and analyzed by labeling with anti-GOS-28 antibodies, as described in Materials and Methods.
teraction measured in the detergent extract faithfully represents the effects seen in the in vitro transport assay.

Discussion
Here, we report the isolation and characterization of a 28-kD Golgi membrane protein, GOS-28, fulfilling all of the criteria currently available to define a v-SNARE. GOS-28 has a single membrane-spanning region at its carboxy terminus, and the bulk of the protein (the amino terminal part) is exposed to the cytoplasm and has the inherent probability to form a coiled coil structure. Furthermore, GOS-28 binds directly or indirectly to a-SNAP, and ATP hydrolysis by NSF results in dissociation of this interaction. Morphological studies revealed that GOS-28 is localized to the Golgi apparatus with an enrichment on vesicular components at the terminal rims of the Golgi. Electron microscopy analysis and fractionation show the efficient inclusion of GOS-28 in COP-coated vesicles, budding in vitro. Antibodies directed against GOS-28 partially inhibit transport from c/s-to medial-Golgi, accumulating uncoated docked vesicles, and also inhibit the binding of GOS-28 to a-SNAP, implying that the binding of a-SNAP to complexes containing GOS-28is needed for fusion after docking. A possible discrepancy in our findings is that while anti-GOS-28 antibodies block transport by only 40--50%, they inhibit GOS-28 binding to a-SNAP by at least 80%. Among other possibilities, this difference could reflect an essential difference between the two assays. The membranes used in the transport assays contain sufficient endogenous a-SNAP to fulfill transport , implying that two populations of SNAREs might exist in them: one free of endogenous a-SNAP and another associated with endogenous a-SNAP. The former would be inhibited by GOS-28 antibody, but the latter would not, resulting in the partial inhibition of transport. On the other hand, in the binding assay, the GST--a-SNAP matrix would only bind the free GOS-28 pool, which will also be accessible to the anti-GOS-28 antibodies. The an-tibody would then efficiently block the GOS-28--~-SNAP association, but only partially inhibit transport.
What is the role of GOS-28 as a v-SNARE in Golgi transport processes? What routes within the stack are specified by this v-SNARE? Answering these important questions will require identifying the cisternae of origin of GOS-28, as well as the location of its partnering t-SNAREs in (or outside of) the stack. Unfortunately, because of the extremely efficient packing of GOS-28 into vesicles, it has not been possible to pinpoint the origin of vesicles containing GOS-28 in favor of c/s versus trans cisternae, and the parmering t-SNAREs are still unknown. Attempts to test for the interaction with one potential partner, syntaxin 5 (Sed5), have revealed binding (data not shown), but its specificity could not be conclusively demonstrated.
The inhibition of reconstituted c/s-to-medial transport by anti--GOS-28 antibody would suggest that GOS-28 is a v-SNARE for this step. The need for GOS-28, however, could be indirect; e.g., GOS-28 could be necessary for retrograde transport from medial-to c/s-Golgi, or for both anterograde and retrograde steps connecting these compartments. For example, Pelham and colleagues have reported that a v-SNARE, Sftlp, is localized to the medial-Golgi in yeast (Banfield et al., 1995) and partners the c/s-Golgi t-SNARE Sed5p (SCgaard et al., 1994). Yet, Sftlp is required for c/s-to-medial transport in vivo. As proposed by Pelham, Sftlp could be used for retrograde transport from later cisternae to the c/s compartment, and the block in anterograde transport could be a secondary consequence of the lack of recycling of critical anterograde machinery (perhaps Sftlp itself) back to the c/s-Golgi. In vivo (Banfield et al., 1995), the anterograde block is complete; in vitro, it would be partial if multiple rounds were required. This could be another explanation (see above) of why antibodies to GOS-28 partially inhibit c/s-to-medial transport in vitro, reflecting the fraction of anterograde transport that is dependent on recycling in vitro under the conditions we used.
Although the precise function of GOS-28 has not been completely established, GOS-28 is dearly a new member of the SNARE family, and the first mammalian v-SNARE localized to the Golgi and implicated in intra-Golgi transport. The predominant localization of GOS-28 to vesicular structures at the terminal rims of Golgi cisternae and its low abundance at the central cisternal region indicate that the terminal rims of Golgi cisternae are functionally specialized sites for the entry and exit of transport vesicles. Further understanding of the functional organiTation of the Golgi stack and its vesicular flow patterns will require the discovery of additional SNAREs and possibly other targeting molecules, as well as understanding their interactions.
Received for publication 17 November 1995 and in revised form 28 January 1996.


extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Chemistry

text:
The ATP-activated hexameric helicase of bacteriophage T4 (gp41) forms a stable primosome with a single subunit of T4-coded primase (gp61).

We have examined the formation of the primosome subassembly of the bacteriophage T4-coded DNA replication (elongation) complex from its helicase, primase, and DNA components. Previously, we had shown that the T4 helicase (gp41) exists in solution in a stable monomer ⇆ dimer equilibrium at physiological protein (and salt) concentrations and forms a hexamer upon activation by ATP (or GTP) binding (Dong, F., Gogol, E. P., and von Hippel, P. H. (1995) J. Biol. Chem. 270, 7462-7473). Here we report that the T4 primase (gp61) is a monomer in solution under the same conditions, and that the ATP-activated helicase binds to a single gp61 primase molecule on appropriate DNA templates to reconstitute a stable primosome. We show that: (i) the gp41 helicase alone does not form a stable complex with DNA templates, although this helicase by itself can carry out moderately processive ATP-driven translocation along single-stranded DNA (Young, M. C., Schultz, D. E., Ring, D., and von Hippel, P. H. (1994) J. Mol. Biol. 235, 1447-1458); (ii) the primase alone does form a stable complex with DNA; (iii) the helicase can bind to the primase-DNA complex in the presence of ATP or GTP to form a stable ternary complex; (iv) this complex consists of six helicase subunits and one primase subunit; and (v) the reconstituted primosome is stable for at least 10 to 20 min after NTP cleavage and dissociation of the hydrolysis products. These results strongly suggest that the functional T4 DNA replication primosome consists of an integrated 6:1 helicase-primase complex bound to DNA, and that the ATP-activated helicase hexamer remains intact throughout the processive DNA replication process.

The elongation phase of DNA replication in many organisms is characterized by similar patterns of molecular organization and mechanism (see Kornberg and Baker (1992)). The replication complex typically contains three functional subassemblies that operate at the replication fork. These are: (i) the central DNA polymerase that catalyzes template-directed 5Ј 3 3Ј leading and lagging strand synthesis of the nascent DNA and often carries a 3Ј 3 5Ј exonuclease (editing) activity as well; (ii) a set of polymerase accessory proteins that stabilize the polymerase on the DNA template by forming a "holoenzyme" complex to permit processive synthesis, especially of the leading strand within the DNA replication fork; and (iii) a primosome complex (composed of helicase and primase) that unwinds the doublestranded template DNA ahead of the replication fork and lays down small RNA primers for the multiple initiation events required for the discontinuous synthesis of lagging strand DNA. These subassemblies combine to form an integrated DNA replication complex that is capable of carrying out coordinated leading and lagging strand synthesis and drive the overall movement of the DNA replication fork.
The bacteriophage T4 utilizes seven discrete T4-coded proteins to carry out the elongation phase of DNA replication (Alberts, 1987;Cha and Alberts, 1988;Nossal, 1994). These proteins comprise a DNA-dependent DNA polymerase (gp43), 1 three accessory proteins (gp45, which forms the processivity "clamp" that binds the polymerase to the template DNA, and a gp44-gp62 complex, that uses ATP to drive the loading of the clamp onto the DNA and polymerase), and the T4-coded singlestranded DNA binding protein (gp32). In concert, these components form a polymerase holoenzyme that can carry out processive DNA replication on primed single-stranded DNA templates under physiological conditions. Efficient DNA replication on double-stranded DNA templates also requires the participation of a primosome subassembly, consisting of the helicase (gp41) and primase (gp61) components. The primosome catalyzes the ATP-(or GTP-) driven unwinding of the double-stranded DNA template and synthesizes the pentameric RNA primers required for coordinate lagging-strand DNA replication. In this paper we focus on the assembly and properties of the functional primosome.
Many studies have shown that while the helicase and primase alone can each carry out its specialized tasks to some extent, the full physiological properties and activities of these enzymes are only displayed when they function within an integrated primosome complex. Thus, the full primosome is required to permit the helicase to unwind double-stranded DNA at the high rates (up to 500 base pairs per s) characteristic of DNA replication in vivo and to permit efficient synthesis by the primase of the physiologically relevant pentaribonucleotide RNA primers (Cha and Alberts, 1986;Hinton and Nossal, 1987;Alberts, 1980, 1981). In addition, both proteins are needed to coordinate leading and lagging strand DNA synthesis at the moving DNA replication fork Cha and Alberts, 1990).
As an initial step in approaching the molecular properties of the T4 primosome, we examined the association states and the structures of the helicase itself (Dong et al., 1995) and showed that the subunits of the T4 DNA replication helicase assemble into a toroidal (ring-shaped) hexamer from its monomer and dimer solution constituents upon activation by binding of the nucleoside triphosphate substrate (ATP or GTP) of the helicase. This study also showed that the NTP-activated gp41 helicase exists as a hexagonal trimer of dimers and suggested that the hexamer may be characterized by pseudo-C3 or pseudo-D3 symmetry.
Here, we build on that study to present a physical biochemical description of the formation of the T4 primosome complex on DNA templates. We show that the ATP-activated gp41 helicase hexamer binds to a single subunit of gp61 primase on a DNA template to form the primosome subassembly of the T4 DNA replication complex. Our results also show that the primosome formed by the helicase hexamer and the primase is very stable and suggest that the primosome remains associated during the entire cycle of ATP hydrolysis, DNA template unwinding, and presumably lagging-strand primer formation.

EXPERIMENTAL PROCEDURES
Reagents and Buffers-The chemicals used for buffers, for gel electrophoresis (including acrylamide, bisacrylamide, TEMED, and ammonium persulfate), and for the silver staining of protein gels, were all Gel Electrophoresis or Analytical Reagent grade. Purine nucleoside triphosphates and their nonhydrolyzable analogs (ATP, GTP, ATP␥S, and GTP␥S) were obtained from Boehringer Mannheim. The homobifunctional protein-protein cross-linking reagent dithiobis(succinimidyl propionate), DSP, was purchased from Sigma.
Preparation of Proteins-The T4-coded DNA replication helicase (gp41) was overexpressed using Escherichia coli strain OR1265/ pDH518  and purified as described previously (Dong et al., 1995). The primase (gp61) was prepared as described . The E. coli strains used to overexpress both proteins were generously provided by Drs. Deborah Hinton and Nancy Nossal (NIH). Both purified proteins migrated as single bands in SDSpolyacrylamide gel electrophoresis when stained with Coomassie Blue G-250. Protein concentrations were determined by UV absorbance at 280 nm, using molar extinction coefficients (⑀ M, 280 ) of 7.6 ϫ 10 4 M Ϫ1 cm Ϫ1 (gp41) and of 6.9 ϫ 10 4 M Ϫ1 cm Ϫ1 (gp61) that were calculated as described (Young et al., 1994;Pace et al., 1995).
Preparation of Reconstituted Primosome Samples-Single-stranded M13mp18 DNA was prepared as described previously (Lechner and Richardson, 1983) and was used as the initial DNA template on which to reconstitute primosome complexes. At room temperature this material contains multiple DNA hairpins, each of which has the essential geometry of a small DNA replication fork. Reconstituted primosomes were usually prepared in the same buffer that we had used previously to form ATP-activated helicase hexamers (Dong et al., 1995). This buffer (TAMK) contained 33 mM Tris-OAc, 50 mM KOAc, 6 mM Mg(OAc) 2 (pH 7.8) and was made 1 mM (sometimes 2 mM) in ATP or GTP (or their nonhydrolyzable analogs, ATP␥S or GTP␥S) before the single-stranded M13 DNA template was added (usually to a final concentration of 5 or 10 nM in DNA molecules). The primase was then added, followed by the helicase. The final concentration of the primase was usually set at 0.5 or 1 M, and that of the helicase ranged from 0.5 to 6 M (in protein monomers). Due to carryover from the protein storage buffer, the experimental samples also generally contained 10 to 20% glycerol; control experiments (data not shown) indicated that glycerol concentrations in this range have little or no effect on the behavior of the reconstituted primosomes in the experiments described in this study. Samples were generally incubated at room temperature for 1 to 2 min after addition of all components (the helicase was added last) prior to being used for further characterization.
Agarose and Polyacrylamide Gel Electrophoresis-Native agarose gels were prepared using 0.5% agarose dissolved (with heating in a microwave oven) in a gel-running buffer containing 33 mM Tris-OAc, 50 mM KOAc, 0.1 mM Mg(OAc) 2 (pH 7.8). A minigel 10 cm in length and 6.5 cm wide was usually poured with a comb to form 12 sample wells. After the samples had been prepared and incubated at room temperature for 1 to 2 min, they were loaded onto the gel and were electrophoresed at ϳ2 V/cm (ϳ40 V for the minigel apparatus) at room temperature. One or two separate lanes were usually loaded with an indicator dye to monitor the progress of the electrophoresis. Gels were run until the bromphenol blue indicator dye had migrated for about 3 cm from the gel origin and were then stained with either ethidium bromide or with SYBR-Green I (Molecular Probes Inc.) to visualize the free DNA and the protein-DNA complex bands.
Discontinuous SDS-polyacrylamide gels for quantitative determination of the protein constituents of protein-DNA complexes were prepared using the method of Laemmli (Laemmli, 1970). A 12.5% (110:1 acrylamide:bisacrylamide) separating gel with a 6% (110:1 acryl:bisacrylamide) stacking gel system was usually used. This system provided good separation between denatured helicase and primase subunits. After electrophoresis, the gels were stained with silver and were analyzed by soft laser scanning densitometry.
Quantitation of Protein Constituents of the Primosome Complexes-Reconstituted primosomes were subjected to native gel electrophoresis to remove unbound protein, after which the protein-DNA complexes were excised from the native agarose gels and were analyzed by SDSpolyacrylamide gel electrophoresis to determine the amounts of helicase and primase present. A Zeineh SL-504-XL soft laser scanning densitometer was used for this quantitation, and data acquisition and analysis were performed using the Videophoresis II computer program (BioMed Instruments Inc.). Due to the shallow linear-response (dynamic) range of the silver-stained gels and the densitometry scanning technique, various amounts of each of the two proteins were also loaded onto gels to which sample proteins of known concentration had been added for calibration. The amount of each protein was then determined by measuring peak areas in densitometer scanning.
Protein-Protein Chemical Cross-linking-The bifunctional chemical cross-linker DSP, dithiobis(succinimidyl propionate), was used to analyze the association state(s) of the gp61 primase by cross-linking protein complexes, as described previously for the gp41 helicase (Dong et al., 1995). Cross-linking experiments were carried out in the same buffer used for the preparation of the primosome samples, either in the presence or in the absence of 1 mM purine nucleoside triphosphate cofactor. Samples were prepared by incubating the gp61 primase (1 M) in the buffer for 1 to 2 min at room temperature, after which cross-linking was initiated by adding small amounts (2% of the total reaction volume) of freshly prepared DSP stock solutions in dimethylformamide (final DSP concentrations in cross-linking reactions were 0.1% or 0.01%, w/v, depending on experiments). Cross-linking reactions were allowed to proceed at room temperature for 20 min 2 and were then quenched by adding a one-tenth volume of 1.4 M ethanolamine-HCl (pH 8.0). After further incubation at room temperature for 20 min, the samples were made 1% in SDS and 10% in glycerol (by volume), incubated at 37°C for 30 min, and analyzed by SDS-polyacrylamide gel electrophoresis using the phosphate-SDS gel system (Weber and Osborn, 1975). After electrophoresis, the protein bands were observed by silver staining.

RESULTS
Formation of the Primosome Complex-A native gel bandshift assay was used to measure the interactions between the components of the primosome. Previous functional and enzymatic studies had shown that the gp41 helicase can interact with single-stranded or forked DNA templates (Young et al., 1994;Dong et al., 1995). However, our native gel band-shift experiments, using single-stranded M13 DNA as a putative binding and translocation (and perhaps helicase) substrate, show that the gp41 helicase by itself does not form complexes with these single-stranded M13 DNA templates that are stable enough to be visualized by this technique (see Fig. 1A, lanes 2-5). Similar observations had been made previously with X174 single-stranded DNA Nossal, 1989a, 1989b).
The observation that the binding of gp41 with its DNA cofactors cannot be observed in standard gel-shift experiments is not necessarily at odds with earlier demonstrations that these DNA cofactors, in similar buffers and at comparable concentrations, can stimulate the ATPase activity of the helicase and perhaps serve as relatively inefficient helicase substrates. An important difference between these experiments is that while the samples for the band-shift experiments were prepared in the presence of ATP or GTP (or their analogs), they were typically run without nucleotide cofactors in the native agarose gel electrophoresis experiment itself. Thus, the interactions between gp41 and single-stranded or forked DNA that we have observed by ATPase stimulation (Young et al., 1994), by native polyacrylamide gel electrophoresis (with ATP present in the gels and the running buffer, Young et al. (1994)), and by direct cryoelectron microscopic imaging (Dong et al., 1995), were all dependent on the continuous presence of purine nucleoside triphosphates and were shown to be relatively weak or transient in the absence of these cofactors.
On the other hand, we now find that gp61 primase can be shown by standard gel-shift experiments to bind and remain stably associated with ss M13 DNA under all buffer conditions tested (Fig. 1A, lanes 6 -9). No nucleotide cofactors are required for this association (lane 6). However, the complexes formed by the primase and ss M13 DNA have a very strong tendency to aggregate, especially when the primase is present at high con-centrations such as those used in Fig. 1A. At lower primase concentrations (representing also a lower primase:DNA ratio), aggregation is less severe (Fig. 1B, lane 1). This aggregation of primase on ss M13 DNA is completely alleviated by the presence of proper amounts of the gp41 helicase, as shown in the experiments presented in Fig. 2A, which were performed at the same primase and ss M13 DNA concentrations (and under the same experimental conditions) as those in Fig. 1A.
Figs. 1B and 2A also demonstrate that gp41 binds stably to a primase-DNA complex in the presence of PuTP as indicated by the further retardation of the complex in the native agarose gels. (A similar increase in protein association on adding gp41 to primase-DNA complexes was observed previously in similar experiments using single-stranded X174 DNA as cofactor (Ri-FIG. 1. Association of primosome proteins with DNA. Native (0.5%) agarose gel-shift electrophoresis was used to analyze the association with single-stranded (hairpin-containing) M13 DNA of gp41 helicase or gp61 primase alone (A) and gp41 helicase and gp61 primase together (B). All reactions were carried out in TAMK buffer (see "Experimental Procedures"), and samples were loaded for gel electrophoresis after 2 min of incubation at room temperature. The concentration of ss M13 DNA in all reactions was 5 nM. A: lanes 2-5, 2 M helicase and ss M13 DNA in buffer only, plus 2 mM ATP, 2 mM GTP␥S, and 2 mM ADP, respectively, as indicated; lanes 6 -9, 1 M primase and ss M13 DNA in buffer only, plus 2 mM ATP, 2 mM GTP␥S, and 2 mM ADP, respectively, as indicated. Lanes 1 and 10 contained free singlestranded M13mp18 DNA only. B, titration of various concentrations of helicase to a constant concentration of primase and ss M13 DNA. The concentrations of the ss M13 DNA and of the primase were 5 nM and 0.5 M, respectively, in all reactions. The concentration of the helicase was 0, 0.5, 1, 1.5, 2, 3, 4, and 6 M in lanes 1 through 8. Lanes labeled D contained free SSM13 DNA only.
FIG. 2. Determination of the protein components of the primosome (helicase-primase-DNA) complex. A, electrophoresis in a native (0.5%) agarose gel electrophoresis to visualize protein-DNA complexes. DNA in the complexes were stained with ethidium bromide. Lanes 1 and 10, free DNA. Lanes 2-4, 1 M primase was preincubated with 5 nM ss M13 DNA in TAMK buffer containing 2 mM ATP for 1 to 2 min, after which 1, 2, or 6 M helicase was added. Lane 5, same binding reaction as in the previous lanes, but contained 0.5 M primase and 6 M helicase. Lane 6, binding reaction containing 0.25 M primase and 6 M helicase. Lane 7, similar to lane 5, but 2 mM GTP␥S was used instead of ATP. Lanes 8 and 9, control experiments using 2 M helicase or primase, without DNA, to show that these proteins alone do not migrate (by chance) to the region where the ss M13 DNA-protein complexes would run (see lanes 8 and 9 of B). B, electrophoresis in a 12.5% (110:1 acrylamide:bisacrylamide) SDS-polyacrylamide gel to analyze the protein contents of the ss M13-bound primosome complexes. Lanes 1 and 10, helicase and primase proteins as standards. Lanes 2-9, protein-DNA complexes cut out from lanes 2-9 of the native agarose gel (A) were recovered and loaded into the corresponding lanes (in the case of lanes 8 and 9, the agarose gel slices were cut to include all positions at which DNA and DNA-protein complexes were found in lanes 1 through 7). The gel was silver-stained after electrophoresis was complete. The total absence of gp41 and gp61 in control lanes 8 and 9 indicates that all proteins detected in lanes 2 through 7 exist in DNAprotein complexes. chardson and Nossal, 1989a).) This conclusion is confirmed by protein content analysis of these complexes using SDS-gel electrophoresis (Fig. 2B).
Many lines of evidence have suggested that the T4 helicase and primase proteins function together as a primosome complex within the DNA replication system. Our findings that the gp61 primase tends to aggregate when interacting with ssDNA by itself, but not when the gp41 helicase is also present, and also that the gp41 helicase alone interacts only weakly with the DNA template, but much more stably with a primase-DNA template complex, support the functional importance of the primosome assembly. The agarose gel shown in Fig. 1B also suggests that the size of the helicase-primase complex approaches a limit when the ratio of helicase to primase subunits exceeds approximately 6:1.
Composition and Stoichiometry of the Primosome-To determine directly the protein subunit stoichiometry of these reconstituted complexes, increasing amounts of gp41 were titrated into fixed concentrations of ss M13 DNA and gp61 until gp41 was clearly present in excess. The resultant DNA-bound complexes were then freed of excess components by native agarose gel electrophoresis, and the ratio of bound helicase to primase subunits was determined by densitometry of the component gp41 and gp61 bands on SDS-polyacrylamide gels (Fig. 2).
The results presented in Fig. 2, A and B, clearly show that in the presence of excess gp41, the amount of gp41 bound within the complex is limited by the amount of gp61 available. In support of this interpretation, we show (in Fig. 2A, lanes 4 -6) that reducing the amount of gp61 by 50% or 75% results in the loss of a corresponding amount of gp41 from the complexes. This shows that gp41 interacts stoichiometrically with gp61 and that the primase is not just acting as a "loading" factor to facilitate the interaction of DNA with gp41. Two-fold higher concentrations of proteins (relative to those used in Fig. 1B) were used in this experiment to permit more accurate measurement of protein concentrations in SDS-polyacrylamide gels by silver staining.
The ratio of gp41 to gp61 subunits in the complexes was determined by laser densitometric scanning of the silverstained SDS gels, using known amounts of each protein loaded onto the same gel for calibration, and was found to reach a limit ratio of 5:1 to 7:1 when excess of gp41 was present (see Table I). This subunit stoichiometry ratio is in accord with the titration experiment shown in Fig. 1B. A completely accurate stoichiometry ratio cannot be determined using this technique because of the very narrow dynamic quantitation range available with protein silver-stained SDS-polyacrylamide gels. However, since gp41 itself forms a hexamer upon activation by the binding of its PuTP substrate (Dong et al., 1995), these data strongly support a 6:1 stoichiometry of gp41:gp61 subunits within the primosome. This ratio is also in good accord with previous reports on the stimulation of gp41 ATPase and helicase activities by added gp61 Richardson and Nossal, 1989b).
To check for the possibility of additional subunit interactions within the primosome, we also used protein-protein cross-linking to see whether primase alone forms dimers or higher associated forms in solution. We used the protein-protein chemical cross-linking procedure that we had previously employed to demonstrate the various association forms of the gp41 helicase (Dong et al., 1995). Cross-linking experiments were performed with gp61 in both the presence and absence of purine nucleoside triphosphates. Our results (see Fig. 3) suggest that, unlike the helicase (lane S), the primase alone exists as a monomer under all solution conditions tested and does not associate to higher forms. This result, together with the fact that gp41 hexamers also do not associate further in solution (Dong et al., 1995), strongly suggests that the functional primosome subassembly contains six helicase subunits together with one monomer of primase. Effects of ADP on the Formation and Stability of the Primosome-ADP is produced by the ATPase activity of gp41; thus, it is important to ask whether ADP has any effect on the stability of the primosome. The experiments shown in Fig. 4 test the effects of this nucleotide on the stability of the primosome and show that ADP does seem to affect the formation or the stability of the primosome. When primosome assembly experiments were performed in the presence of ADP and analyzed by native agarose gel electrophoresis (using the same conditions as the previous native gels, i.e. no nucleotide cofactors in the running buffer), some complexes larger than the primase-DNA complexes seen in Fig. 1A (lanes 5-9) were detected. However, these complexes were much smaller than those formed in primosome assembly experiments carried out in the presence of ATP or GTP␥S. Based on these findings alone, we cannot know whether these intermediate-sized complexes correspond to "regular" (6:1) primosomes that have partially dissociated during gel electrophoresis or to complexes that were incompletely assembled prior to loading onto the gels. As we will show below, primosomes assembled by ATP binding are stable for long periods of time, even after the nucleoside triphosphate substrates have been completely removed. Such results strongly argue that the intermediate-sized complexes we observe here FIG. 3. Protein-protein chemical cross-linking suggests that primase alone exists as a monomer in solution. Lane S shows a cross-linking ladder of the T4 gp41 helicase in the presence of GTP␥S and is used here as a size standard and to show the levels of crosslinking expected in these experiments for oligomerized subunits. Lanes 1 and 2, gp41 and gp61 proteins (not cross-linked) were run as controls; lanes 3 and 4, gp61 was treated with cross-linkers in the absence (lane 3) and in the presence (lane 4) of 1 mM GTP␥S. Nearly all the gp61 primase protein was intramolecularly cross-linked (as shown by the increase in electrophoretic mobility of the bands) and no intersubunit cross-linking was seen. The protein content of the bands was analyzed by SDS-polyacrylamide gel electrophoresis using the phosphate buffergel system (see "Experimental Procedures") and silver-staining of the separated bands.  (Dong et al., 1995), we had shown that the gp41 helicase alone undergoes reversible hexamer formation that is dependent on the binding of its PuTP substrate. Therefore, it was important to determine whether the helicase within the primosome remains hexameric during the entire translocation/helicase/replication cycle or whether it dissociates and reassociates during each cycle of ATP binding and hydrolysis. We have approached this question by determining whether helicase hexamers within the reconstituted primosome complexes bound to ss M13 DNA remain hexameric after ATP removal. (We note that the persistence of such complexes in native gel-shift experiments, as in Figs. 1A and 2A, do not speak to this point since complexes that appear stable within gels may be held together artificially by gel "caging" effects.) Fig. 5 shows that primosomes formed as 6:1 helicase-primase complexes on ss M13 DNA and incubated for 10 min (Fig. 5) or 20 min (data not shown) at room temperature after ATP removal remain intact.
Stable Primosomes after ATP Depletion Do Not Contain ATP, ADP, or P i -We have shown above that primosomes formed by the interaction of helicase and primase are stable for prolonged periods, even after ATP depletion. We ask here whether these stable primosomes contain bound ATP or its hydrolysis products. To this end experiments were performed using radioactively labeled ATP carrying 32 P at either the ␥or the ␣-phosphate positions. We reasoned that if the primosomes remaining after removal of free ATP and hydrolysis products by gel filtration still bound intact or hydrolyzed ATP (i.e., both ADP and P i ), the primosome complexes should be labeled in both experiments; on the other hand, if the primosome binds only ADP or P i , only the primosome reconstituted in the presence of ␣or ␥-[ 32 P]ATP (but not both) will carry the radioactive label. Of course, the former outcome would not distinguish between intact bound ATP and its hydrolyzed products; however, previous studies (Young et al., 1994) have shown that the ATP hydrolysis rate by the helicase is very fast, and it is therefore unlikely that ATP bound to gp41 would remain unhydrolyzed after the long incubation periods used here.
The results of such experiments are shown in Fig. 6. Lanes 1 and 2 contained the primosome complexes reconstituted in the presence of ␥and ␣-32 P-labeled ATP, respectively. If all six helicase subunits in the primosome contain a bound ADP or P i (or both), the complexes will carry 32 P labels with equal amounts of radioactivity, as shown in control lanes 5 and 6 (which were loaded with the equivalent amounts of free radio-activity). The amounts of radioactivity in lanes 5 and 6 were chosen so that if all six subunits of the helicase bind 32 P, the radioactivity in the primosome complexes (loaded in lanes 1 and 2) would be equal to that seen in these lanes. We find, however, that no detectable radioactivity is found at the position at which the primosome complexes run on the native agarose gel. (The position is defined in the control experiment using SYBR-green I nucleic acid stains in Fig. 6B, lane 2). Therefore, the primosome complex remains intact as a 6:1 helicase:primase complex even after all nucleoside triphosphate and its hydrolyzed products have been released.
Taking these results together, it is clear that the PuTP substrate of the helicase is required to form the primosome. However, once formed, the primosome is very stable by itself. Thus, even though the ATP-activated hexamerization of the helicase is reversible (Dong et al., 1995), the helicase hexamer is stabilized against dissociation by the primase within the primosome. Hence, the primosome remains stable over a much longer period than the milliseconds required to complete the ATPase and helicase cycles, which strongly suggests that once the primosome is formed the gp41 helicase component remains hexameric during the full functional cycle and does not alternate between different oligomeric forms. FIG. 5. Primase stabilizes helicase hexamers in the primosome after ATP removal. A, samples were prepared by incubating the helicase, primase, and ss M13 DNA together in the presence of 1 mM ATP. After ATP was removed by gel filtration through a BioSpin column, the samples were incubated further at room temperature and were then run on the native (0.5%) agarose gel. Lane D, free ss M13 DNA; lane 1, complex without ATP removal; lane 2, sample applied to the agarose gel immediately after ATP removal; lane 3, sample incubated for an additional 10 min at room temperature after ATP removal prior to loading onto the gel. B, analysis of the protein composition of the bands in A on a 10% (30:0.8) SDS-polyacrylamide gel. Protein-DNA complexes isolated from the above agarose gel were mixed with SDS gel loading buffer, heated in boiling water for 5 min, and analyzed by SDS-polyacrylamide gel electrophoresis. The three lanes correspond, respectively, to the samples recovered from lanes 1-3 of the agarose gel of A.

Formation and Stabilization of the Primosome of the T4 DNA
Replication Complex-In this study we have shown that the gp61 primase binds stably by itself to appropriate DNA templates, while the gp41 helicase hexamer in the absence of the primase interacts with DNA cofactors relatively weakly and transiently. These observations support an assembly pathway like that outlined in Fig. 7. However, the order of addition of the components shown here may not be invariant, since we know that gp41 helicase alone can interact functionally with the DNA substrate as shown in previous studies (Young et al., 1994;Dong et al., 1995) and that primase and helicase in the absence of DNA can also form a weakly associated complex. 3 The definition of an unambiguous assembly pathway for primosome formation is not crucial to understanding the functional and mechanistic properties of this complex. The structure that the primosome adopts, and the interactions that stabilize it during its function, are more important concerns. Results from this study, together with our previous findings (Dong et al., 1995), clearly demonstrate that the subunits of the T4 DNA replication helicase form a hexamer when activated by the binding of PuTP substrate, and that this hexamer is capable of binding to a primase monomer and DNA to reconstitute a primosome. What we have learned here provides an observational basis that permit us to establish some qualitative "boundary conditions" for the formation and stability of complete and partial primosome complexes. These observations are summarized in Fig. 8. The quantitative magnitudes of the interactions that underlie this scheme will be presented elsewhere. 3 It is clear from this study that once properly assembled in the presence of purine nucleoside triphosphates, the primo-some remains stable for 20 min or more under the conditions tested even if the supply of unhydrolyzed PuTP is depleted. Physiological conditions, under which ample PuTP is available, certainly favor stable primosome formation, and we have shown that the stable primosome complex, containing 1 primase and 6 helicase subunits, is likely to remain assembled through the entire replication cycle on the T4 genome.
Implications of the Hexagonal Structure of the Active gp41 for the Structure of the Primosome-Determination of subunit stoichiometry and composition of the primosome is the first step to establishing the structure of the complex. The 6 to 1 stoichiometry established here, in combination with our previous demonstration that the helicase itself is a ring-shaped hexamer (Dong et al., 1995), seems to suggest that the helicase hexamer is likely to remain in this general conformation within the complex. However, this result does not define the position of the primase within the primosome. We know that the primase must interact with the DNA and, at least transiently, with all six helicase subunits. This raises the question of the location of gp61 inside the primosome complex. Considering that the helicase hexamer itself is a ring-shaped structure with pseudo-C3 or D3 symmetry, the primase is most likely to reside in a position close to the C3 rotational axis of the helicase hexamer. However, a primase subunit that shuttles from one helicase monomer (or dimer) binding site to another during primosome function is also possible. Such behavior has been seen for the subunits within the bovine heart mitochondrial F 1 -ATPase (Abrahams et al., 1994). In addition, we note that the bacteriophage T7 primosome complex is also hexameric, but that each subunit carries both a helicase and a primase domain (Mendelman et al., 1993;Patel and Hingorani, 1995). At present, there is no structural evidence to discriminate between a centrally located primase within the primosome and a primase that shuttles from one helicase subunit to another as the primosome functions.
Functional Integration of the Primosome within the T4 DNA Replication Complex-Formation of separate DNA polymerase holoenzyme and primosome subassemblies seems to be general in the DNA replication systems of many different organisms (Kornberg and Baker, 1992). In addition, many (most?) DNA helicases that are involved in genomic DNA replication seem to exist and function as hexamers (see Table II in Dong et al. (1995)) upon activation either by their PuTP substrates or by the DNA template. Furthermore, those hexameric helicases that have been investigated in detail all seem to adopt a toroi- FIG. 6. Primosome complexes remain stable after release of bound ATP and its hydrolyzed products. A, samples were prepared by the same procedures as for the sample in Fig. 5A, lane 4, in the presence of 1 mM ATP containing trace amounts of ␥-[ 32 P]ATP or ␣-[ 32 P]dATP. After removal of free ATP from the protein-DNA complexes using BioSpin gel filtration columns, the samples were incubated at room temperature for 10 min and were then analyzed on 0.5% native agarose gels to determine whether the complexes retained radioactivity (see text). Lanes 1 and 2 contained samples prepared in the presence of ␥and ␣-[ 32 P]ATP, respectively. Lanes 5 and 6 were loaded with amounts of radioactive ␥or ␣-[ 32 P]ATP equivalent to that expected in the primosome complexes in lanes 1 and 2 if each helicase subunit in the primosome complex were complexed with one 32 P-labeled ATP or its hydrolyzed products. Lanes 3 and 4 were loaded with sample buffer only to separate the control and experimental lanes. After electrophoresis, the gel was dried on a DEAE-filter paper (Whatman) with no heating and autoradiographed on Fuji x-ray film. B, control experiments on native agarose gels to monitor the position of primosome complex. Lane 1, free ss M13mp18 DNA. Lane 2, primosome complexes prepared in the presence of 1 mM ATP. The positions of the free or complexed DNA was monitored by SYBR-green staining.
FIG. 7. The association pathway proposed for the in vitro assembly of the T4 DNA replication primosome from the gp41 helicase, the gp61 primase, and the DNA template in the presence of nucleoside triphosphate substrates. Note that this is not the only pathway that is consistent with our data (see text). dal (ring-shaped) subunit arrangement. In spite of these large scale structural similarities, no atomic level structure has been established for any helicase or primosome. Nevertheless, it is likely that DNA replication primosomes from different organisms will be found to share a certain degree of structural similarity, and it is reasonable to speculate that the hexameric structure of these helicases (and the resulting common structural features that may characterize primosomes) will turn out to have general mechanistic significance.
Recent studies (Spacciapoli and Nossal, 1994;Barry and Alberts, 1994;Morrical et al., 1994) suggest that the T4-coded gp59 protein (the gp41 helicase assembly factor) is important under some conditions for the loading of the gp41 helicase onto DNA to form a functional complex with other replication proteins. Our finding here that the primosome can be formed on ss M13 DNA in the absence of the gp59 protein support the notion that this protein may only be required to assemble the helicase onto single-stranded DNA-binding protein (gp32)-coated DNA templates, in agreement with previous results (Barry and Alberts, 1994;Morrical et al., 1994;Spacciapoli and Nossal, 1994). Whether this assembly factor is required to load the activated helicase hexamer onto a primase that is already present on a gp32-coated DNA template is not yet known.


text:
A highly efficacious lymphocyte chemoattractant, stromal cell-derived factor 1 (SDF-1)

Chemotactic factors are postulated to direct emigration of lymphocytes from the blood stream into sites of inflammation. Members of a family of chemotactic cytokines, termed chemokines, have been shown to attract lymphocytes but efficacy, i.e., the maximal percentage of attracted cells, has been low. We have identified a highly efficacious lymphocyte chemotactic activity in the supernatants of the murine bone marrow stroma cell line MS-5 which attracts 10-fold more lymphocytes in vitro than currently described lymphocyte chemoattractants. Purification of this chemotactic activity revealed identity to stromal cell-derived factor 1 (SDF-1). SDF-1 acts on lymphocytes and monocytes but not neutrophils in vitro and is both a highly efficacious and highly potent mononuclear cell attractant in vivo. In addition, SDF-1 induces intracellular actin polymerization in lymphocytes, a process that is thought to be a prerequisite for cell motility. Since SDF-1 is expressed constitutively in a broad range of tissues it may have a role in immune surveillance and in basal extravasation of lymphocytes and monocytes rather than in inflammation.

L Ymphocytes function as the reservoir of immunological memory. In contrast to neutrophils and monocytes, lymphocytes may leave tissues through lymphatics, emigrate from the blood stream into lymphoid and non-lymphoid tissues, and recirculate multiple times during their life history (for review see reference 1). Chemoattractants are postulated to help direct lymphocyte traffic into sites of inflammation and into lymphoid and non-lymphoid tissues during recirculation (1,2). The best lymphocyte chemoattractants identified to date are members of a family of small, 60 to 80 amino acid chemotactic cytokines termed chemokines. The chemokines can be divided into two subfamilies, the CXC-and the CC-chemokines. The members of each subfamily are more homologous to one another than to members of the other subfamily, and differ in the spacing of the first two cysteines. The two subfamilies differ in their chromosomal localization and cluster on chromosome 4 (CXC-chemokines) and chromosome 17 (CC-chemokines), respectively. While CXC-chemokines, e.g., interleukin 8 (IL-8), act mostly on neutrophils, members of the CC-chemokines, e.g., monocyte chemoattractant protein 1 (MCP-1) 1, regulated on activation normal T cell expressed 1Abbreviations used in this paper: MCP-1, monocyte chemoattractant protein 1; MIP-10t, macrophage inflammatory protein lce; RANTES, regulated on activation normal T cell expressed and secreted; SDF, stromal cellderived factor. and secreted (RANTES), and macrophage inflammatory protein lcl (MIP-lo0, have been shown to exert function on monocytes, eosinophils and lymphocytes (2,3). However, the efficacy, i.e., the percent of attracted ceils, of lymphocyte chemoattractants described so far is low, whereas chemokines active on monocytes, neutrophils, and eosinophils are highly efficacious (2).
The bone marrow not only produces precursor and immature lymphocytes, but also harbors mature T lymphocytes. These give rise to graft versus host disease and are an important problem in bone marrow transplantation. To understand what drives recirculation of mature lymphocytes into the bone marrow we screened supernatants from bone marrow stromal cells for chemotactic activity. We subsequently identified a lymphocyte chemoattractant produced by bone marrow stromal cells and purified it to homogeneity. It is highly efficacious, attracting greater than 10-fold more lymphocytes than currently described chemoattractants. It stimulates actin polymerization, is active on monocytes and not on neutrophils in vitro, and in vivo is both a highly efficacious and highly potent mononuclear cell attractant. Its sequence shows identity with stromal cell-derived factor (SDF-1)(4-6), previously known to augment B cell proliferation. SDF-1 shows equidistant sequence homology to CC-and CXC-chemokines and strong conservation in evolution that suggest it as a primordial chemokine. SDF-1 by contrast to most chemokines is constitutively expressed in a broad range of tissues (4)(5)(6) and therefore may have a role in immune surveillance rather than in inflammation.

Materials and Methods
Three-Step Purification of a Lymphocyte Chemoattractant. Adherent MS-5 routine bone marrow stromal cells (7) (8). Molecular weight was determined by microcapillary HPLC/electrospray ionization/tandem mass spectrometry on a Finnigan TSQ7000 triple quadrupole mass spectrometer (San Jose, CA) and by matrix-assisted laser desorption mass spectrometry on a Finnigan Lasermat 2000 (Hemel, England) as described previously (9). Protein concentrations were determined by BCA protein assay (Pierce Chem. Co., Rockford, IL).

Cell Preparation and Chemotaxis Assay.
Human PBL were obtained from healthy donors by erythrocyte sedimentation with 6% Dextran T500 followed by Ficoll-Histopaque-1077 (Sigma Chem. Co., St. Louis, MO) separation. Monocytes were removed by two 30-min steps of plastic adherence (10). Human neutrophils were prepared as described (11). Human PBL (5 X 105) in 100 ILl RPMI-1640 medium, 0.25% human serum albumin (HSA) were added to the top chamber of a 6.5-mm diameter, 5-txm pore polycarbonate Transwell TM culture insert (Costar, Cambridge, MA) and incubated in duplicate with the indicated concentrations of purified material or dilutions of fractions from purification in the bottom chamber for 3 h. Fractions from the heparin, SP Sepharose, and HPLC columns were diluted 2-fold, 10-to 50-fold, and 40-to 1,000-fold, respectively, before assay. The chemotaxis assays across bare filters and endothelial monolayers cultured on filters were carried out as described previously (10,11), except cells that transmigrated into the lower chamber were vigorously suspended and counted with a FACScan | (Becton Dickinson, San Jose, CA) for 20 s at 60 Ixl/min, gating on the forward and side scatter of the cell type of interest. A 1:20 dilution of input PBL was similarly counted. For human monocytes, peripheral blood mononuclear cells (5 • 10s), i.e., the cells from Ficoll-Histopaque before monocyte depletion, were migrated for 2 h through 5-1xm pore polycarbonate filters and were scattergated on monocytes to exclude lymphocytes. Human neutrophils (5 • 105) were migrated for 1.5 h through 3-1xm pore polycarbonate filters. Murine spleen and bone marrow cells were obtained from 129 mice and erythrocytes were lysed in NH4C1buffer for 4 min on ice. Spleen cells and bone marrow cells were migrated for 3 h and 1.5 h and scatter-gated on lymphocytes and neutrophils, respectively. To effectively remove neutrophils and monocytes from plastic in the lower chamber a final concentration of 5 mM EDTA was added to each well and cells were resuspended vigorously. Actin Polymerization Assay. Actin polymerization was tested as described before (12). Briefly, human PBL (1.25 X 106/ml) in L15 medium at 37~ were incubated with the indicated chemokines at concentrations that were shown to be optimal in chemotaxis experiments or PMA at 100 ng/ml. At the indicated time points, cells (400 b~l) were added to 100 ILl of a solution at 37~ containing 4 X 10 -v M FITC-labeled phalloidin, 0.5 mg/ml 1-c~-lysophosphatidylcholine (both Sigma) and 18% formaldehyde in PBS. The fixed cells were subjected to fluorescence flow cytometry on a FACScan | (Becton-Dickinson, San Jose, CA) and the mean relative fluorescence of each sample determined. All data points are plotted relative to the mean relative fluorescence of the sample before addition of chemoattractant.
Biologic Activity of SDF-1 In Vivo. C57B1/6 mice were injected subcutaneously at a marked spot on the abdomen with 100 ng SDF-1 or murine MCP-1/JE in 20 ~1 sterile PBS or PBS alone. Injected protein solution was shown to contain <0.004 ng/ml LPS by the limulus assay (Sigma). Mice were sacrificed 18 h after injection, injection sites were excised and snap-frozen 5-txm cryosections were subjected to hematoxylin and eosin staining and for immunohistochemistry, sections were pretreated with avidin/ biotin blocking reagent (Vector Labs., Burlingame, CA), stained with mAbs to Thy

Results and Discussion
Purification of a Highly Efficacious Lymphocyte Chemoattractant. In peripheral blood lymphocyte chemotaxis assays, serum-free conditioned media of three murine bone marrow stromal cell lines, MS-5 (7), PA-6 (14), and ST-2 (15), consistently attracted 20-50% of the input lymphocytes, whereas supernatants from N I H 3T3 fibroblasts showed little activity. The chemotactic activity was purified 1,310fold from MS-5 conditioned medium by sequential hep-arin, cation exchange, and reversed-phase H P L C chromatography ( Table 1). The chemoattractant activity coeluted with a single protein peak in reversed-phase H P L C and migrated as a single band in SDS-PAGE of 8 kD (Fig. 1). W e consistently obtained 80-90 Ixg of purified protein from 1,000 ml of conditioned medium. The NH2-terminal sequence of the material was KPVSLSYRXPXRFFE, with smaller amounts of a sequence lacking the first five residues of the above sequence, S Y R X P X R F F E S H I A . This sequence, and the amino acid composition, showed identity with stromal cell-derived factor-1 (SDF-1), a chemokine identified by signal sequence trap cloning (4) and as a factor that augments IL-7-stimulated pre-B cell growth (5). Activity of SDF-1 in chemotaxis assays has not previously   (A-C) or murine (D-E) leukocyte subtypes were subjected to chemotaxis through 5-1*m pore (A, B, and D) or 3-pmi pore (C and E) Transwell TM filters to the indicated chemoattractant in the lower chamber and the % of input cells that accumulated in the lower chamber was measured. The bars and symbols show duplicates and mean for one representative experiment of three. (F) Actin polymerization in human PBL. SDF-1 (1,000 ng/ml), huMCP-1 (100 ng/ml), huMIP-la (100 ng/ml), and PMA (100 ng/ml) at the indicated optimal concentrations were added to PBL at time 0 and intracellular F-actin content in PBL at the indicated times was measured with FITC-phalloidin in permeabilized cells.
been reported. Two alternatively spliced forms of SDF-1 differ in the COOH-terminal four amino acid residues (4). Electrospray and matrix-assisted laser desorption mass spectrometry gave a mass of 7847.1 +_ 0.4 daltons consistent with that of 7849.2 daltons predicted for SDF-hx beginning with the KPVSL sequence and with the COOH-terminal lysine residue removed by processing. SDF-1 is not glycosylated, as shown by its mass and lack of effect of N-glycanase or O-glycanase (Fig. 1), and lacks predicted glycosylation sites.

SDF-I E~ciently Attracts Lymphocytes and Monocytes
In Vitro. Purified SDF-1 was a highly efficacious lymphocyte chemoattractant, attracting 70% of human peripheral blood lymphocytes in filter chemotaxis assays at optimal concentrations of about 1 I*g/ml (Fig. 2 A). Consistent with the high percentage of enfigrating cells, the propor-tions of CD4 +, CD8 +, CD26 +, CD45RA +, and CD45R, O + T lymphocyte subsets were similar in the starting and emigrated populations by immunofluorescent flow cytometry. The proportions in the starting and emigrating populations were for CD4 47 and 48%, for CD8 24 and 28%, for CD26 64 and 66%, for CD45RA 42 and 44%, and for CD45RO 40 and 43%, respectively. SDF-1 was also an efficacious chemoattractant and showed a similar doseresponse for murine lymphocytes (Fig. 2 D) and human monocytes (Fig. 2 B), but was not active on human or murine neutrophils (Fig. 2, C and E). Chemotaxis to SDF-1 was dependent on a G~i-coupled receptor. Pretreatment of PBL with 50 ng/ml of pertussis toxin inhibited migration to 1 Ixg/ml of SDF-1 of 85 + 2.9% of input lymphocytes to background levels of 3.2 -+ 0.7%.
We compared SDF-1 to other lymphocyte chemoattrac- tants in a range of concentrations previously reported to be most active; the greatest migratory response to each and the concentration giving the greatest response are shown in Table 2. MCP-1, RANTES, and MIP-lc~ consistently induced migration over background, and were more potent than SDF-1, i.e., active at lower concentrations, but were substantially less effective, attracting only 2-4% of the input cells. Furthermore, the chemotactic index, the ratio of specific to background migration is only 1.3-to 2.3-fold for MCP-I, R_ANTES, and MIP-I& in filter chemotaxis assays. Fig. 2 A visualizes this difference in efficacy between SDF-1 and proinflammatory chemokines over a broad range of chemokine concentrations. Two different sources of lymphotactin showed little activity in our assay, as did different interleukins (Table 2), basic fibroblast growth factor, leukemia inhibitory factor, transforming growth factorq3, platelet-derived growth factor AA, and granulocyte monocyte-colony stimulating factor (data not shown).

SDF-I Induces Actin Polymerization in Lymphocytes.
To monitor intracellular actin reorganization, a process that is thought to be a prerequisite for cell movement, we assayed actin polymerization (12). SDF-1 (1,000 ng/ml) induced a transient twofold increase in intracellular filamentous-actin within 15 s in human peripheral lymphocytes (Fig. 2 F). Actin polymerization in neutrophils is stimulated by IL-8, fMLP, C5a, and LTB4 with the same kinetics (16,17) but to our knowledge an increase in F-actin has not been previously observed with chemokines in lymphocytes. Consistent with our findings in the chemotaxis assay, the other lymphocyte chemoattractants stimulated little actin polymerization.  *Chemokines and interleukins were assayed over a 1,000-fold concentration range and results at the optimal concentration are shown. Results are mean and SD of three independent experiments. Chemotactic index was defined as the ratio between the percentage of cells migrated to sample and to medium alone.

SDF-I Is an Efficacious and Potent Chemoattractant for Mononudear Cells In Vivo. Subcutaneous injection of 100
ng of purified SDF-1 into the skin of C57B1/6 mice elicited a dense infiltrate of mononuclear cells in the subcutaneous tissue throughout a substantial surrounding area (Fig. 4, B and C). Multiple sections failed to reveal any infiltrate at PBS injection sites (Fig. 4 A). Injection of 100 ng of mufine MCP-1 elicited an infiltrate of far fewer cells that were localized to the point of injection, and contained both monocytes and neutrophils (Fig. 4 F), in agreement with a Table 3   CC-and C-Chemokines CXC-chemokines SDF-1 Figure 5. A phylogenetic tree of chemokines. The mature amino acid sequences of known chemokines in Swiss-Prot were aligned using the clustal method with a PAM250 residue weight table and default settings of gap penalties for pairwise and multiple alignments of 3 and 10, respectively (Megalign; DNASTAR., Madison, WI). Distances to branch points are proportional to amino acid sequence divergence from predicted ancestral sequences. The clustal alignment method grouped the CC-chemokines and the C-chemokine lymphotactin together in the same subfamily. Comparing mature human protein sequences, SDF-la is 21 + 4% and 27 -+ 3% (mean + SD) identical to CC-and CXC-chemokines, respectively; for comparison, MCP-1 is 49 + 19% and 17 + 2% identical and IL-8 is 19 -+ 3% and 35 + 7% identical to other human CC-and CXC-chemokine subfamily members, respectively.
previous report (18). Less than 1% of the cells infiltrating to SDF-1 were neutrophils. Immunohistochemistry with mAb to Thy-1 (19) revealed the presence of up to 10% lymphocytes ( Fig. 4 D) among a majority of monocytes/macrophages as demonstrated by staining with mAb MOMA-2 (20; Fig. 4 E).
We have shown that SDF-1 exhibits unparalleled efficacy as a lymphocyte chemoattractant in vitro and effectively attracts mononuclear cells in vivo. It attracts ~-,10-fold more lymphocytes than previously described chemoattractants, and achieves an efficacy comparable only to that seen previously with attractants ofneutrophils, eosinophils, and monocytes (2). In vitro SDF-1 is less potent, i.e., requires higher concentrations for maximal activity, than MCP-1, RANTES, and MIP-lc~. However, in vivo SDF-1 is both a highly potent and highly efficacious chemoattractant for attraction of mononuclear cells. The number of cells infiltrating to 100 ng SDF-1 is far greater than to 100 ng MCP-1 in parallel experiments, and greater than seen in previous in vivo studies to all doses of MCP-1, MCP-2, MCP-3, and RANTES (18,(21)(22)(23). It is intriguing to observe the high percentage of monocytes in the infiltrate although SDF-1 attracts a similar percentage of lymphocytes and monocytes with similar dose-response in vitro (Fig. 2, A and B). This finding may be due to the higher migratory capability of monocytes compared to lymphocytes that has been observed in different in vivo models (24). Issekutz and Issekutz show that skin injections of zymosan-activated serum (ZAS), TNF-0q IFN-~/, or LPS in rats induce consistently more than fivefold greater monocyte than lymphocyte accumulation at all time points tested (24). This is comparable to the ratio of monocyte to lymphocyte emigration seen here with SDF-1 in vivo. Greater expression of adhesion molecules, greater motility, or a lower frequency of exit from the site after entry, may favor greater accumulation of monocytes than lymphocytes in vivo, even when the strength of the chemoattractive stimulus is comparable.
The in vivo potency of SDF-1 is remarkable. SDF-1 is highly basic with a predicted pI of 10.9 compared to 9.8 and 9.9 for IL-8 and MCP-1, respectively. SDF-1 binds heparin with higher affinity than IL-8 or MCP-1 in that elution from heparin Sepharose requires 800-900 mM NaC1, whereas IL-8 and MCP-1 elute at ~500 mM NaC1 (3,25). Strong binding to glycosaminoglycans may present SDF-1 in an effective manner or help retain it at sites of production, and may in part explain its observed potency in vivo.
The evolutionary and sequence homology properties of SDF-1 are unique among chemokines. First, the human homologue of SDF-1 (26)has a single conservative Ile to Val substitution compared to the murine SDF-1 studied here. This strong conservation (99% identity) is unparalleled among chemokines; for comparison MCP-1 and RANTES are 60 and 83% identical between mouse and human. Second, based on amino acid sequence homologies, SDF-1 is almost equally related/distant from both CC-and CXC-chemokines ( Fig. 5 and legend). This coincides with the finding that the human SDF-1 gene is localized on chromosome 10 while all other known CXC-and CC-chemokines cluster on chromosomes 4 and 17, respectively (26). Alignment of the mature amino acid sequences of known chemokines generates a phylogenetic tree that confirms that the CC-and CXC-chemokines belong to separate subfamilies, and suggests that SDF-1 belongs to neither and forms an outgroup, SDF-1 is predicted among currently known chemokines to have diverged the least from a primordial chemokine ancestor; this is consistent with the high conservation of SDF-1 between species. Because of both its functional and predicted phylogenetic distinctions from CC-and CXC-chemokines, SDF-1 is thus far a unique chemokine; whether it represents the first member of a new chemokine subfamily remains to be determined, SDF-1 is constitutively produced by bone marrow stro-real cells, and therefore could be involved in trafficking of lymphocytes and monocytes to this site. Bone marrow stromal cells produce it in large quantities, with 0.8 b~g/ml recovered from cultured cells. Whether SDF-1 plays a special role in bone marrow, or whether production is higher to compensate for rapid loss through the fenestrated endothelium of bone marrow, remains to be determined. IfSDF-1 is important in attracting mature lymphocytes to bone marrow, antagonism of this function before harvest of bone marrow for transplantation could be clinically beneficial. SDF-1 may be of wider importance. SDF-1 mRNA is constitutively expressed in every tissue thus far examined, including heart, liver, lung, brain, muscle, spleen, and kidney (4,6,27), although protein expression has not yet been characterized. Most other chemokines are only expressed in response to proinflammatory stimuli, and are thought to regulate the emigration and activity of leukocytes at inflammatory sites. By contrast, expression of SDF-1 is unaltered by inflammatory stimuli (27). Normal trafficking of lymphocytes through tissues during immune surveillance requires G protein-coupled receptors (1) but the identity of the receptors on lymphocytes and the ligand(s) remain uncharacterized. In the absence of inflammation blood monocytes constantly replace mononuclear phagocytes in solid tissues maintaining a steady state by emigrating from the blood stream and differentiating into macrophages. SDF-1 has a tissue distribution that suggests it as a candidate for a function in lymphocyte recirculation, and in basal recruitment ofmonocytes in normal replenishment and turnover of tissue mononuclear phagocytes, rather than in inflammation.