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Renografin Classification Essay

Abstract

The plasma membrane ATPase, encoded by PMA1, is delivered to the cell surface via the secretory pathway. Previously, we characterized a temperature-sensitive pma1 mutant in which newly synthesized Pma1-7 is not delivered to the plasma membrane but is mislocalized instead to the vacuole at 37°C. Severalvps mutants, which are defective in vacuolar protein sorting, suppress targeting-defective pma1 by allowing mutant Pma1 to move once again to the plasma membrane. In this study, we have analyzed trafficking in the endosomal system by monitoring the movement of Pma1-7 in vps36, vps1, andvps8 mutants. Upon induction of expression, mutant Pma1 accumulates in the prevacuolar compartment in vps36cells. After chase, a fraction of newly synthesized Pma1-7 is delivered to the plasma membrane. In both vps1 andvps8 cells, newly synthesized mutant Pma1 appears in small punctate structures before arrival at the cell surface. Nevertheless, biosynthetic membrane traffic appears to follow different routes in vps8 and vps1: the vacuolar protein-sorting receptor Vps10p is stable in vps8 but not in vps1. Furthermore, a defect in endocytic delivery to the vacuole was revealed in vps8 (andvps36) but not vps1 by endocytosis of the bulk membrane marker FM 4-64. Moreover, in vps8 cells, there is defective down-regulation from the cell surface of the mating receptor Ste3, consistent with persistent receptor recycling from an endosomal compartment to the plasma membrane. These data support a model in which mutant Pma1 is diverted from the Golgi to the surface invps1 cells. We hypothesize that in vps8and vps36, in contrast to vps1, mutant Pma1 moves to the surface via endosomal intermediates, implicating an endosome-to-surface traffic pathway.

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INTRODUCTION

In mammalian cells, entry of newly synthesized hydrolases into the lysosomal pathway is mediated by a sorting receptor at thetrans-Golgi complex (Kornfeld and Mellman, 1989). Analogously, in Saccharomyces cerevisiae, Vps10 is a sorting receptor that recycles between the trans-Golgi and the endosome, recognizing a signal within soluble hydrolases and thereby effecting their delivery to the vacuole (Marcusson et al., 1994; Cooper and Stevens, 1996). Genetic screens to identify yeast mutants defective in delivery of the vacuolar marker carboxypeptidase Y (CPY) have resulted in >40 VPS genes required for proper vacuolar protein sorting, revealing the complexity of the vacuole biosynthetic pathway (Rothman and Stevens, 1986; Robinson et al., 1988). Because biosynthetic traffic to the vacuole or lysosome intersects with endocytic traffic at the endosome, a subset of the vps mutants also displays defects in endocytosis (Daviset al., 1993; Munn and Riezman, 1994; Singer-Kruger et al., 1994).

The general organization of the endocytic pathway has been especially well characterized in mammalian cells. Specifically, extracellular molecules and plasma membrane proteins travel through the endocytic pathway to the lysosome via two sequential intermediates, early and late endosomes. Similarly, in yeast, two populations of endosomes have been defined morphologically as well as biochemically (Singer-Kruger et al., 1993; Hicke et al., 1997;Mulholland et al., 1999). Nevertheless, a molecular understanding of endosome function remains far from complete (Gruenberg and Maxfield, 1995; Mellman, 1996; Riezman et al., 1997). In mammalian cells, a recycling pathway is well established by which certain internalized proteins are delivered back to the plasma membrane (Gruenberg and Maxfield, 1995; Mellman, 1996). In contrast, in yeast, the existence of a direct traffic pathway from the endosome to the cell surface has not been established to date, although a number of recent studies suggest such a pathway (Harsay and Bretscher, 1995; Yuanet al., 1997; Ziman et al., 1998). Indeed, two populations of Golgi-derived secretory vesicles have been isolated from yeast; the finding that preventing endocytosis results in the accumulation of only one of these populations is consistent with the idea that one of these classes of vesicles may derive from endocytic recycling (Harsay and Bretscher, 1995).

We were prompted to examine whether an endosome-to-surface traffic pathway exists in yeast by studies of PMA1, which encodes the plasma membrane ATPase. Normally, Pma1 reaches the cell surface via the secretory pathway (Brada and Schekman, 1988; Chang and Slayman, 1991). However, in the temperature-sensitive pma1-7mutant, newly synthesized Pma1 is defective for targeting to the plasma membrane at 37°C and instead is delivered to the vacuole via the endosome (Chang and Fink, 1995; Luo and Chang, 1997). Although the molecular basis for vacuolar delivery of Pma1-7 is unknown, we have considered the possibility that there is a post-endoplasmic reticulum quality control mechanism that recognizes and targets mutant Pma1 into the endosomal/vacuolar system (Chang and Fink, 1995; Hong et al., 1996; Li et al., 1999). Several vpsmutants, which are defective in vacuolar protein sorting, have been identified that cause rerouting of mutant Pma1 to the plasma membrane (Luo and Chang, 1997). By disrupting the recycling of a Golgi-based quality control receptor, these vps mutants might allow Pma1-7 to travel directly from the Golgi to the cell surface. With this in mind, we have compared trafficking pathways of mutant Pma1 invps1, vps8, and vps36. Remarkably, the data suggest that in vps8 and vps36 cells, Pma1-7 moves to the plasma membrane only after it has entered the endosomal system.

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MATERIALS AND METHODS

Media and Strains

Standard yeast media and genetic manipulations were as described (Sherman et al., 1986). Yeast transformations were performed by the lithium acetate method (Gietz et al., 1992). Strains used in this study are listed in Table 1. All strains except those marked with asterisks are isogenic with L3852.vps8-Δ1::LEU2 and vps36-Δ1 were isolated as suppressors of pma1-7 after insertional mutagenesis (Luo and Chang, 1997). ACY76 was generated in a one-step gene replacement by transformation of L3852 with pPS83, aHIS3-marked VPS8 disruption construct (Horazdovsky et al., 1996) provided by B. Horazdovsky (Texas Southwestern Medical Center, Dallas, TX). ACY33 was generated in a one-step gene replacement by transformation of L3852 with pKJH2, aLEU2-marked VPS27 disruption construct (Piperet al., 1995) provided by T. Stevens (University of Oregon, Eugene). WLY65 was generated in a one-step gene replacement by transformation of L3852 with pBS-YPT51-LYS2 (Singer-Kruger et al., 1994) provided by B. Singer-Kruger (University of Stuttgart, Germany). WLX20-5D is an ascospore from a cross between WLX16-1A and WLX4-2A (MATa lys2Δ201 leu2-3,112 ura3-52 ade2 vps27-Δ1::LEU2). Integration of MET-HA-PMA1and MET-HA-pma1-7 at ura3-52 was accomplished by transforming yeast with pWL10 and pWL9 linearized with NcoI. ACY72 was constructed by pop-in, pop-out gene replacement ofSTE3 by transformation of ACX66-2D (MATα his3Δ200 leu2-3,112 ura3-52 ade2 trp1Δ63 GAL+) with pSL1904 (provided by N. Davis, Wayne State University, Detroit, MI), resulting in replacement of the STE3 promoter with aGAL1,10 promoter. ACY81 was constructed by transformation of ACY72 with pPS83, a vps8::HIS3disruption construct. ACY84 and ACY85 were constructed by transformation of ACY72 and ACY81 with pAS173, apep4::hisG-URA3-hisG disruption construct (Chang and Fink, 1995), to disrupt PEP4.

Table 1.

Yeast strains used in this study

Molecular Biology

Plasmids with HA-tagged pma1-7 and PMA1under the control of the MET25 promoter were constructed as follows. With the use of XhoI (polylinker) andBstEII sites, a 1.7-kilobase (kb) fragment from the 5′ region was removed from pAC7 and pAC4 bearing 4.5-kbHindIII–HindIII pma1-7 andPMA1 inserts, respectively (Chang and Fink, 1995). The fragment was replaced with a 750-base pair (bp) fragment from pFT4 (provided by C. Slayman, Yale University, New Haven, CT), which has aHindIII site introduced at −27 bp from the start codon, generating pWL1 and pWL2. A 4.2-kbHindIII–HindIII fragment bearing thePMA1 coding sequence was excised from pWL1 and pWL2 and placed after the MET25 promoter of FB1521 (Mumberg et al., 1994). The 4.6-kb fragments containing MET-pma1-7and MET-PMA1 were excised with the use ofSacI–XhoI polylinker sites and placed into pRS306, a URA3-marked YIp, generating pWL5 and pWL6, respectively. To introduce an HA epitope, the plasmid pXZ28 (containingPMA1 with an HA epitope introduced after the second amino acid; provided by J. Haber, Brandeis University, Waltham, MA) was used as a template for PCR. A fragment of 0.8 kb was amplified with the use of the oligonucleotide TCCCCCGGGAGCTAGTTAAAGAAAATC to introduce aSmaI site at −67 bp from the start codon and the oligonucleotide CCTTCACCTCTCTTAACA. After cutting with SmaI and BstEII, the PCR fragment was used to replace the corresponding fragments in pWL5 and pWL6 to generate pWL9 and pWL10, respectively.

Protein Induction

To detect newly synthesized Pma1, plasmids were used in which HA-tagged mutant or wild-type Pma1 was placed under the control of theMET25 promoter. Cells were grown under repressing conditions in minimal medium containing 600 μM methionine. To induce synthesis of Pma1, cells were washed once with water and resuspended in methionine-free medium. At the same time, cells were shifted to 37°C. Synthesis of HA-tagged Pma1 was shut off by adding 2 mM methionine alone or in the presence of 100 μg/ml cycloheximide.

To study Ste3, cells were grown to midlog phase at 30°C in synthetic complete minus uracil medium with 2% galactose. Glucose (3%) was added to stop synthesis of Ste3. For detection of Ste3 by Western blot, anti-Ste3 mAb (provided by G. Sprague, University of Oregon) was used. For Ste3 detection by indirect immunofluorescence, cells were transformed with a GAL1-STE3 construct in which a c-myc epitope is fused to the carboxyl terminus of STE3 (pSL2015; provided by N. Davis, Wayne State University).

Indirect Immunofluorescence, Cell Fractionation, Western Blotting, and Metabolic Labeling

For indirect immunofluorescence, cells were spheroplasted with oxalyticase (Enzogenetics, Corvallis, OR) and permeabilized with methanol and acetone, as described (Rose et al., 1990). Cells were stained with anti-HA (BABCO, Berkeley, CA) or anti-myc mAb (Santa Cruz Biotechnology, Santa Cruz, CA) followed by Cy3-conjugated secondary antibody (Jackson ImmunoResearch, West Grove, PA). Pulse-chase experiments were visualized with the use of an Olympus(Lake Success, NY) IX70 microscope, and the images were collected digitally (with the same exposure for each time point within an experiment) and adjusted at the same settings with Adobe Photoshop 4.0 (Adobe Systems, Mountain View, CA). All other fluorescent microscopy experiments were photographed with the use of a Zeiss Axiophot microscope (Carl Zeiss, Thornwood, NY).

Cell fractionation on Renografin (gift of L. Marsh, Albert Einstein College of Medicine) density gradients was performed essentially as described (Schandel and Jenness, 1994; Jenness et al., 1997). Harvested samples were placed on ice in the presence of 10 mM azide. Cell lysates were prepared by vortexing cells with glass beads in the presence of a protease inhibitor cocktail including 1 mM PMSF (Chang and Slayman, 1991). After centrifugation at 400 × g for 5 min to remove unbroken cells, lysate (0.5 ml) was mixed with 0.5 ml of Renografin-76, placed at the bottom of a centrifuge tube, and overlaid with 1 ml of 34, 30, 26, and 22% Renografin solutions. For pulse-chase experiments, samples from different time points were loaded on gradients after normalization to lysate protein by the Bradford assay (Bio-Rad Laboratories, Hercules, CA). Gradients were centrifuged in an SW50.1 rotor overnight at 150,000 × g at 4°C. Fourteen fractions (350 μl) were collected from the top of each gradient. To prevent Renografin from interfering with subsequent Western blotting, membranes were diluted with Tris-EDTA buffer and pelleted by centrifugation at 100,000 × g for 1 h. Intracellular membranes and plasma membrane were consistently contained in fractions 6–7 and 10–11, respectively. Therefore, quantitation of newly synthesized Pma1 was performed by pooling fractions 1–8 and 9–14. Distribution of mutant Pma1 after induction and chase was determined by Western blotting and calculated after subtraction of the background signal obtained at time 0.

For Western blotting of cell lysate, samples were prepared as described previously (Chang and Slayman, 1991) and normalized to lysate protein. After separation by SDS-PAGE, proteins were transferred to nitrocellulose. Anti-HA Western blotting was performed with the use of mAb. Antibodies against Kex2, Pep12, and Gas1 were provided by S. Nothwehr (University of Missouri, Columbus), H. Pelham (Medical Research Council Laboratories, Cambridge, UK), and T. Doering (Washington University, St. Louis, MO), respectively. Immune complexes were visualized by chemiluminescence detection reagents (ECL Western blotting detection system; Amersham, Arlington Heights, IL) or125I-protein A (Amersham). Quantitation of Western blots with the use of 125I-protein A was performed with the use of a Molecular Dynamics (Sunnyvale, CA) phosphorimager.

Metabolic labeling was performed with the use of cultures grown to midlog phase in synthetic complete medium without methionine and cysteine. Cells were resuspended at 1 OD600/ml, labeled with Expre35S35S (New England Nuclear, Boston, MA) (2 mCi/25 OD600cells) for 5 min at room temperature, and chased in the presence of 10 mM methionine and cysteine. At various times during the chase, aliquots were placed on ice in the presence of 10 mM Na azide. Lysates were prepared and resuspended in RIPA buffer (10 mM Tris, pH 7.5, 150 nM NaCl, 2 mM EDTA, 1% NP40, 1% deoxycholate, 0.1% SDS) for immunoprecipitation.

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RESULTS

Inducible Synthesis of Mutant Pma1

To visualize movement of newly synthesized Pma1, we used constructs in which wild-type PMA1 and mutantpma1-7 were tagged with an HA epitope and placed under the control of the MET25 promoter. Although a low level of synthesis is detected under repressing conditions, the Western blot in Figure 1 shows a large increase in synthesis of Pma1 upon activation of the MET25 promoter (by removal of methionine from the medium). Quantitation reveals that the level of wild-type Pma1 upon induction is approximately fivefold greater than that of mutant Pma1, probably because of concurrent degradation of the mutant protein. After stopping synthesis, degradation of newly synthesized mutant Pma1 is readily apparent after 90 min, whereas wild-type Pma1 remains stable (Figure 1).

Fig. 1.

Stability of newly synthesized wild-type and mutant Pma1. Pma1 levels in wild-type cells bearing eitherpMET-HA-pma1-7 or pMET-HA-PMA1 (WLY103 and WLY104). Exponentially growing cells were washed, transferred to methionine-free medium to induce synthesis of epitope-tagged protein, and shifted to 37°C. After 90 min, methionine was added to prevent further Pma1 synthesis, and incubation continued for an additional 90 min. Lysates were prepared and analyzed by Western blot with anti-HA antibody. Note that newly synthesized wild-type Pma1 is stable, whereas mutant Pma1 is degraded during chase.

Previously, we demonstrated that degradation of mutant Pma1 occurs at 37°C upon trafficking of the newly synthesized protein to the vacuole (Chang and Fink, 1995; Luo and Chang, 1997). To confirm that epitope-tagged Pma1 behaves in the same manner, indirect immunofluorescence localization was performed in pep4 cells defective in vacuolar protease activity. Synthesis of HA-tagged Pma1 was induced for 1 h at 37°C, and the cells were then stained with anti-HA antibody. As shown in Figure2, newly synthesized Pma1-7 is accumulated at the vacuole (bottom left), which is recognized as pale regions under phase contrast optics (bottom right). In contrast, staining of cells expressing wild-type Pma1 is at the cell surface (top).

Fig. 2.

Vacuolar delivery of mutant Pma1. Indirect immunofluorescence localization of newly synthesized Pma1.pep4 cells carrying pMET-HA-pma1-7(WLY152) or pMET-HA-PMA1 (WLY153) were washed free of methionine to induce synthesis of epitope-tagged Pma1 and shifted to 37°C for 1 h. Samples were then fixed, spheroplasted, and permeabilized for indirect immunofluorescence staining with anti-HA antibody followed by Cy3-conjugated secondary antibody. Left panels show indirect immunofluorescence images; right panels show the corresponding phase images. Newly synthesized wild-type Pma1 is localized at the cell surface, whereas mutant Pma1 is delivered to the vacuole.

Newly Synthesized Mutant Pma1 Reaches the Prevacuolar Compartment of vps36 Cells before Arrival at the Cell Surface

Mutant Pma1 is delivered to the cell surface in two class E vps mutants, vps36 and vps27(Luo and Chang, 1997). Class E vps mutants are characterized by defective trafficking from the endosome to the vacuole as well as from the endosome back to the Golgi (Piper et al., 1995). In both vps36 and vps27 mutants, some proteins traversing endocytic and biosynthetic pathways are trapped in a novel prevacuolar compartment that is visualized as a characteristic large spot next to the vacuole; on the other hand, newly synthesized CPY is missorted in these cells and travels directly from the Golgi to the cell surface (Raymond et al., 1992; Piper et al., 1995). To determine the route by which mutant Pma1 travels to the plasma membrane in the class E vps mutants, the localization of newly synthesized Pma1 was determined by staining with anti-HA antibody. As shown in Figure 3A (0′, top), no staining is seen in vps36Δ cells before induction of mutant Pma1 synthesis. After a 90-min induction period, staining of newly synthesized Pma1-7 appears predominantly in large perivacuolar spots; little cell surface staining is apparent (Figure 3A, middle, arrowheads). Previously, we established that this staining pattern is characteristic of mutant Pma1 accumulating in the prevacuolar compartment (Luo and Chang, 1997). The same pattern of Pma1-7 accumulation was also seen in vps27Δ cells (data not shown). These observations confirm that newly synthesized mutant Pma1 enters the endosomal system in class E vps mutants. In contrast, wild-type Pma1 does not detectably accumulate in the prevacuolar compartment or other intracellular compartments (Figure4).

Fig. 3.

Localization of newly synthesized mutant Pma1 invps36 cells. Synthesis of HA-tagged mutant Pma1 was induced at 37°C, followed by a 90-min chase in the presence of cycloheximide, as described in MATERIALS AND METHODS. (A) Indirect immunofluorescence localization was performed before induction of synthesis (0′), after a 90-min induction (90′ on), and after a 90-min chase in vps36 cells (WLY156) (90′ off). Cells were stained with anti-HA antibody followed by Cy3-conjugated secondary antibody. In vps36 cells, newly synthesized mutant Pma1 is accumulated in the prevacuolar compartment after induction (middle panel, arrowheads). A slight increase in staining at the plasma membrane is apparent after chase (bottom panel, arrows). (B) Quantitative distribution of newly synthesized Pma1-7 in vps36. Distribution in intracellular and plasma membrane fractions was quantitated after fractionation on Renografin density gradients, as described in MATERIALS AND METHODS. Data are expressed as absolute arbitrary units.

Fig. 4.

Localization of newly synthesized wild-type Pma1 in vps cells. Synthesis of HA-tagged wild-type Pma1 was induced at 37°C in vps1 (WLY145),vps8 (WLY128), and vps36 (WLY157), as described in MATERIALS AND METHODS. Indirect immunofluorescence staining was performed on cells after 90 min of induction. Cells were stained with anti-HA antibody followed by Cy3-conjugated secondary antibody.

To determine whether mutant Pma1 can move to the cell surface from the prevacuolar compartment, vps36Δ cells were reexamined after an additional 90-min chase period. Cycloheximide was included during the chase to ensure that no additional Pma1 synthesis could occur. As shown in Figure 3A (bottom), staining of the prevacuolar compartment declines after the chase. Much of the decrease is likely due to degradation in the proteolytically active prevacuolar compartment (Cereghino et al., 1995). In addition, staining of the plasma membrane appears to increase slightly (Figure 3A, bottom, arrows). A similar pattern was observed in vps27Δ cells (data not shown). Quantitation of newly synthesized Pma1-7 at the cell surface was performed by fractionation on Renografin density gradients, which efficiently separate plasma membranes from intracellular membranes (Jenness et al., 1997) (see below). Figure 3B shows the results of such an experiment in which the level of mutant Pma1 in intracellular and plasma membranes in vps36 cells was quantitated after induction and chase and plotted in arbitrary units. In this experiment, the plasma membrane fraction contains 31% of total Pma1-7 present during the induction period; after chase, the plasma membrane fraction represents 41% of Pma1-7 remaining. In three independent experiments, an ∼9% fractional increase at the cell surface was observed after chase. Thus, relocation of Pma1-7 from the endosomal system to the cell surface appears to occur, albeit inefficiently.

Cell Surface Delivery of Newly Synthesized Mutant Pma1 in vps1 and vps8

To compare a possible endosome-to-surface route with a Golgi-to-surface route, mutant Pma1 trafficking was followed invps1 cells. Vps1 is a dynamin-like protein required for formation of endosome-bound vesicles from the Golgi, and all endosome-directed traffic is diverted to the cell surface invps1 mutants (Nothwehr et al., 1995). Figure5 (left middle panel) shows mutant Pma1 localization in vps1Δ cells after a 90-min induction period. Both bright punctate cytoplasmic staining and some surface staining (arrows) are apparent. After 90 min of chase (left bottom panel), the punctate staining has disappeared and there is exclusive staining of the cell surface. These observations are consistent with direct transport of newly synthesized mutant Pma1 from the Golgi to the plasma membrane.

Fig. 5.

Localization of newly synthesized mutant Pma1 invps1 and vps8 cells. Synthesis of HA-tagged mutant Pma1 was induced for 90 min at 37°C. Indirect immunofluorescence localization of mutant Pma1 was performed invps1 (WLY144; left panel) and vps8(WLY127; right panel) at time 0 (0′), after a 90-min induction (90′ on), and after a 90-min chase (90′ off). Upon induction, Pma1-7 is seen in small punctate structures in vps1 andvps8 cells. At this time in vps1, surface staining is also apparent (arrows). After chase, mutant Pma1 is distributed predominantly at the cell surface in vps1and vps8 cells.

vps8 mutation also allows Pma1-7 to travel to the plasma membrane (Luo and Chang, 1997). Because vps8 cells accumulate the bulk membrane marker FM 4-64 in an endocytic intermediate compartment distinct from the prevacuolar compartment (Luo and Chang, 1997) (see below), it was of interest to examine the pathway to the cell surface taken by Pma1-7 in vps8. As shown in Figure 5, after induction for 90 min, mutant Pma1 localizes to punctate cytoplasmic structures (right middle panel). After a 90-min chase, staining is predominantly at the plasma membrane, indicating that newly synthesized Pma1-7 moves to the cell surface (right bottom panel). Movement to the plasma membrane in both vps8 andvps1 cells is unaffected by the presence of cycloheximide (data not shown). Compared with vps36, cell surface delivery of mutant Pma1 appears more efficient in vps8 andvps1 (compare bottom panels of Figures 3A and 5).

Although the immunofluorescence localization pattern of Pma1-7 invps8Δ resembles that in vps1Δ cells, careful comparison suggests slightly more surface staining after induction invps1 (Figure 5, left middle panel, arrows). To compare further vps1 and vps8 mutants, cell fractionation was performed on Renografin density gradients. Figure6A shows the distribution of marker proteins after fractionation of wild-type cells on a Renografin gradient. The plasma membrane protein Gas1 is found predominantly in fractions 10 and 11. Newly synthesized wild-type Pma1 is also localized in these fractions. In contrast, Kex2 and Pep12, membrane proteins that recycle between the Golgi and the endosome (Wilcox et al., 1992; Becherer et al., 1996), are predominantly distributed in fractions 5–7. Figure 6B shows the fractionation pattern of Pma1-7 in vps1Δ and vps8Δ cells. After 90 min of induction at 37°C in vps1Δ cells, the majority of newly synthesized Pma1-7 cofractionates with plasma membrane in fractions 10 and 11. In contrast, in vps8Δ cells, a larger fraction of newly synthesized Pma1-7 is distributed in intracellular membrane fractions. Because there is distinct and consistent separation between intracellular and plasma membranes on Renografin density gradients, fractions 1–8 and 9–14 were pooled for analysis by quantitative Western blotting. As shown in Figure 6C, after 90 min of induction, >80% of mutant Pma1 has reached the cell surface in vps1, whereas in vps8, Pma1-7 is mostly intracellular. After 90 min of chase, the fraction of mutant Pma1 at the cell surface is increased in vps8Δ cells. These data reveal that Pma1-7 is delivered to the plasma membrane with different kinetics invps1 and vps8.

Fig. 6.

Kinetics of mutant Pma1 movement to the cell surface in vps1 and vps8 cells. Wild-type cells bearing pMET-HA-PMA1 and vps1 andvps8 cells bearing pMET-HA-pma1-7 were resuspended in methionine-free medium and shifted to 37°C for 90 min to induce synthesis of epitope-tagged Pma1. Lysates were prepared and fractionation on Renografin density gradients was performed as described in MATERIALS AND METHODS. Fractions were collected and assayed for marker proteins. (A) Western blot showing distribution of newly synthesized wild-type Pma1, the plasma membrane marker Gas1, and the Golgi/endosome markers Kex2 and Pep12 in wild-type cells (WLY104). (B) Gradient distribution of Pma1-7 in vps1 andvps8 cells (WLY144 and WLY127) . Synthesis of HA-tagged mutant Pma1 was induced for 90 min at 37°C. At this time invps1 cells, newly synthesized Pma1 is predominantly in plasma membrane–containing fractions (lanes 10 and 11); invps8, mutant Pma1 is found in intracellular fractions (lanes 6 and 7) as well as in plasma membrane–containing fractions (lanes 10 and 11). (C) Quantitative distribution of newly synthesized mutant Pma1 in vps1 and vps8 cells. Distribution of Pma1-7 in intracellular and plasma membrane fractions was quantitated after a 90-min induction at 37°C (on) and after a 90-min chase (off), as described in MATERIALS AND METHODS. Data from a representative experiment are expressed as absolute arbitrary units. There is a decrease in the total amount of Pma1 after the chase, likely attributable to a fraction of newly synthesized Pma1 moving to and being degraded in the vacuole. (Inset) Mutant Pma1 levels are expressed as a percentage of the total at each time point. Data are means ± SD of three or four independent experiments. Delivery to the plasma membrane is more rapid in vps1 cells than invps8 cells.

Trafficking of Newly Synthesized Vps10 in vps1 and vps8 Mutants

The kinetic differences in cell surface arrival is consistent with the possibility that Pma1-7 may take different routes to the plasma membrane in vps1 and vps8. To examine this possibility in greater detail, trafficking of Vps10 was compared invps1 and vps8 mutants. In wild-type cells, Vps10, the CPY-sorting receptor, is a stable protein that recycles between thetrans-Golgi and the endosomes (Marcusson et al., 1994; Cooper and Stevens, 1996). Previous reports have shown that recycling of trans-Golgi membrane proteins, including Vps10, is disrupted in vps1 mutants; Vps10 travels instead to the plasma membrane, where it undergoes rapid internalization, delivery to the vacuole, and degradation (Wilsbach and Payne, 1993; Cereghinoet al., 1995; Nothwehr et al., 1995). Figure7 shows analysis of newly synthesized Vps10 by pulse labeling of cells with [35S]methionine and [35S]cysteine followed by chase for various times. In vps1Δ cells, Vps10 undergoes proteolytic cleavage at 2 h of chase (arrow), whereas it appears stable in wild-type and vps8Δ cells. Similarly, vps1, but not vps8, perturbed the stability of thetrans-Golgi membrane protein Kex2 (data not shown). These data support the idea that biosynthetic membrane traffic follows different routes in vps8 and vps1.

Fig. 7.

Trafficking of Vps10 in vps1 andvps8 cells. Pulse-chase experiments were used to analyze the stability of Vps10 in wild-type (L3852), vps8(WLX16-1A), and vps1 (ACX58-3C) cells. Cells were radiolabeled at room temperature with Expre35S35S for 5 min and chased for various times. Vps10 was immunoprecipitated and analyzed by SDS-PAGE and fluorography. Newly synthesized Vps10 undergoes degradation invps1 (arrow) but remains stable in vps8and wild-type cells.

Endocytosis of the Bulk Membrane Marker FM 4-64

Of relevance to defining the trafficking route of Pma1-7 is the report that there is defective endocytosis of the fluorescent membrane marker FM 4-64 in vps8 (Luo and Chang, 1997). To characterize further the endocytic defect in vps8, a time course of FM 4-64 endocytosis was examined. Figure8A shows that FM 4-64 staining occurs predominantly at the plasma membrane when wild-type orvps8 cells are incubated with the dye at 0°C, as described previously (Vida and Emr, 1995). (The small bright fluorescent spots that are also seen [Figure 8A, 0′] are likely endocytic structures formed during photography of the cells.) After warming the cells to permit endocytosis to proceed, cell surface staining disappears at similar rates in both wild-type and vps8 cells, indicating that internalization from the cell surface is not affected invps8. At 20 min after warming, FM 4-64 is seen in the cytoplasm as well as in the vacuolar membrane in wild-type andvps8 cells. By 60 min after internalization, FM 4-64 has been cleared from the cytoplasm and is exclusively at the vacuole membrane in wild-type cells (Figure 8A, top right panel), whereas much of the internalized dye remains in vesicular intermediates in the cytoplasm of vps8Δ cells (Figure 8A, middle right panel).

Fig. 8.

An early endocytic intermediate is accumulated in vps8 but not vps1 cells. (A) Time course of FM 4-64 endocytosis. Exponentially growing cells were incubated on ice for 30 min with 40 μM FM 4-64. Cells were then washed and incubated for an additional 20 or 60 min at 30°C. At 60 min in vps8 (WLX16-1A) and ypt51 (WLY65) cells, FM 4-64 is accumulated in an endocytic intermediate similar to that seen at 20 min in wild-type cells (L3852). (B) FM 4-64 endocytosis is delayed in vps8 at a step before the prevacuolar compartment. After labeling for 15 min with FM 4-64, cells were washed, resuspended, and incubated for 1 h at 30°C before visualization and photography. vps8 (ACY76), vps27(ACY33), vps8 vps27 (WLX20-5D), and vps1(ACX58-3C) cells are shown. The pattern of FM 4-64 accumulation invps8 vps27 double mutants resembles that seen invps8 cells.

In vps1 cells, FM 4-64 labels multiple vacuolar compartments (Figure 8B), reflecting the fragmented vacuolar morphology of the cells (Raymond et al., 1992). Nevertheless, no defect in endocytic delivery to the vacuole was detected, in agreement with previous reports (Wilsbach and Payne, 1993; Nothwehr et al., 1995).

FM 4-64 endocytosis was also performed in double mutants in whichvps8 mutation was combined with a class E vpsmutation. After 1 h of internalization in vps27 cells, accumulation of FM 4-64 in the prevacuolar compartment is seen as a large bright spot next to the vacuole (Figure 8B) (Vida and Emr, 1995). However, in vps8 vps27 double mutants, the pattern of FM 4-64 accumulation largely resembles that seen in vps8 cells, with much of the fluorescence signal in vesicular intermediates in the cytoplasm (Figure 8B). The same result was obtained in vps8 vps36 double mutants (data not shown). In contrast, in cells in which the class E vps mutation is combined withvps1, there is FM 4-64 accumulation in the prevacuolar compartment but not at an early endocytic step (data not shown). These data indicate that the endocytic defect seen in the vps8mutant occurs before the prevacuolar compartment of class Evps cells.

Consistent with delayed transport through an early endocytic intermediate in vps8, a similar pattern of dye accumulation was observed in ypt51Δ cells (Figure 8A, bottom panels). Ypt51 is a small GTPase and homologue of mammalian Rab5, and previous work has shown that ypt51 cells accumulate internalized α factor in an early endocytic intermediate (Sambrook et al., 1989; Singer-Kruger et al., 1994, 1995).

Endocytosis of the Mating Receptor Ste3

To analyze further the endocytic defect of vps8, the behavior of the cell surface receptor Ste3 was examined. In wild-type cells, Ste3 undergoes constitutive endocytosis and vacuolar degradation (Davis et al., 1993). To follow the fate of cell surface Ste3 in the absence of new receptor synthesis, STE3 was placed under the control of the GAL1 promoter. Figure9A shows Western blot analysis of Ste3 stability at various times after glucose addition to repressSTE3 expression. In wild-type cells, Ste3 is rapidly degraded upon glucose addition. In contrast, in vps8, the steady-state level of Ste3 (at time 0) is increased and the rate of Ste3 degradation is decreased (Figure 9A). These results are in agreement with the kinetic delay in delivery to the vacuole observed by FM 4-64 endocytosis (Figure 8). The pep4mutation stabilizes Ste3 in both wild-type and vps8 cells, indicating that Ste3 degradation is due to vacuolar delivery (Figure9A) (Davis et al., 1993).

Fig. 9.

Effect of vps8 on endocytosis of Ste3. Inhibition of vacuolar degradation occurs concomitantly with an increase in cell surface Ste3 in vps8 cells. (A) Western blot after preventing new synthesis of Ste3 in wild-type (ACY72) andvps8 (ACY81) cells. To stop new Ste3 synthesis, glucose (3%) was added to GAL1-STE3 cells exponentially growing at 30°C in the presence of galactose. At the indicated times after glucose addition, cells were harvested and lysate was prepared for analysis of Ste3 protein level (top). Time 0 represents the steady-state level of Ste3 before the addition of glucose. The bottom panel shows that degradation of Ste3 upon glucose addition is dependent on PEP4; stabilization of Ste3 is seen inpep4 (ACY84) and vps8 pep4 (ACY85) cells. (B) Indirect immunofluorescence localization of Ste3 after inhibition of new Ste3 synthesis. Wild-type (ACY72) and vps8(ACY81) cells expressing a GAL1-STE3-myc plasmid (pSL2015) were grown to midlog phase in galactose-containing medium. At 5 and 60 min after the addition of glucose (3%), cells were fixed, permeabilized, and stained with anti-myc antibody followed by Cy3-conjugated secondary antibody. Photographs showing the 5- and 60-min time points represent 4- and 8-s exposures, respectively. Arrowheads indicate cell surface staining in vps8cells.

Indirect immunofluorescence was used to observe Ste3 endocytosis (Figure 9B). Within 5 min after glucose addition, the distribution of Ste3 in wild-type and vps8 cells appears similar (Figure 9B, top panels). At this time, Ste3 is seen predominantly at the plasma membrane, although some Ste3 is also found in small intracellular spots, likely reflecting endocytic intermediates. By 1 h after stopping further receptor synthesis, Ste3 staining in wild-type cells is markedly diminished; although some punctate staining is visible, staining at the cell surface is not readily apparent. In contrast to that in wild-type cells, plasma membrane Ste3 staining persists invps8 cells, consistent with receptor recycling to the plasma membrane.

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DISCUSSION

We have analyzed trafficking pathways in the endosomal system by monitoring the movement of newly synthesized Pma1-7. In several vps mutants, a fraction of Pma1-7 is rescued from vacuolar degradation and delivered to the cell surface. We show that invps1 cells, newly synthesized mutant Pma1 can move to the plasma membrane after appearing briefly in punctate intracellular compartment(s) (Figure 5). Consistent with previous reports (Wilsbach and Payne, 1993; Nothwehr et al., 1995), it seems likely that mutant Pma1 in vps1 cells is routed directly to the plasma membrane from the Golgi complex.

In class E vps mutants, newly synthesized Pma1-7 accumulates first in the prevacuolar compartment (Figure 3). After chase in the presence of cycloheximide to ensure that no further synthesis could occur, a small fraction (∼10%) of newly synthesized Pma1-7 was observed to move to the plasma membrane. These data support a model in which protein traffic can flow from the prevacuolar compartment to the plasma membrane, albeit inefficiently. Because there is defective retrograde transport from the prevacuolar compartment to the Golgi in class E vps mutants (Piper et al., 1995), it is unlikely that transport of mutant Pma1 to the cell surface occurs via a Golgi intermediate. It is possible, however, that movement of Pma1-7 in these cells from the prevacuolar compartment to the surface occurs via an early endosomal intermediate (Figure10) (see below).

Fig. 10.

Model showing trafficking step(s) impaired byvps mutations. Proposed impaired step(s) in transport from the Golgi to the vacuole are labeled with the correspondingvps mutations. Transport from the Golgi to the endosomal system is blocked in vps1Δ cells (Nothwehr et al., 1995); endosome-bound traffic, including Pma1-7, is redirected to the surface from the Golgi. In class E vpsmutants, such as vps36 and vps27, there is defective anterograde transport from the prevacuolar compartment to the vacuole as well as defective retrograde transport to the Golgi (Piper et al., 1995; Nothwehr et al., 1996). In class E vps mutants, Pma1-7 is proposed to travel to the plasma membrane from the prevacuolar compartment via an early endosome. In vps8 cells, there is defective transport from the early endosome to the vacuole; movement of mutant Pma1 to the plasma membrane in vps8 is proposed to occur as a consequence of accumulation in an early endosome.

Our data have relevance to understanding the mechanism by which misfolded proteins are delivered to the vacuole. A model has been proposed in which vacuolar delivery of misfolded proteins occurs by receptor-mediated transport (Chang and Fink, 1995; Hong et al., 1996; Li et al., 1999). Our observation that mutant Pma1 enters the endosomal system of class E vpsmutants argues that such a quality control receptor probably does not recycle routinely through the prevacuolar compartment.

Mutant Pma1 is also routed to the cell surface in vps8cells (Figure 6). Our immunofluorescence and cell fractionation experiments do not have sufficient resolution to prove unequivocally that Pma1-7 enters the endosomal system of vps8 mutants before reaching the surface. Nevertheless, several observations prompt us to hypothesize that mutant Pma1 moves to the plasma membrane from an early endosomal compartment in vps8 cells (Figure 10). First, cell surface arrival of mutant Pma1 occurs more slowly invps8 compared with vps1 (Figure 6). Second, pulse-chase analysis shows that Vps10 undergoes rapid degradation invps1 cells but not in vps8 cells. These data are consistent with vps8 and vps1 mutations having different effects on membrane traffic. Third, FM 4-64 endocytosis experiments indicate defective transport through an early endocytic compartment in vps8, in contrast to vps1 cells, which have no apparent endocytic defect (Figure 8). Finally, although plasma membrane internalization is not impaired (Figure 8), down-regulation of Ste3 from the cell surface is impaired byvps8 (Figure 9). These observations suggest persistent receptor recycling to the surface from an endosomal compartment. (Note that cell surface Ste3 is also increased in the class E vpsmutant ren1/vps2, in which there is a block in traffic from the prevacuolar compartment to the vacuole [Davis et al., 1993].)

Enhanced endosome-to-surface trafficking in vps8represents the simplest model that can account for all of the observations we have reported. Previous work on Vps8p reveals that it is a large membrane-associated protein containing a RING finger zinc-binding motif (Chen and Stevens, 1996; Horazdovsky et al., 1996). A role for Vps8 in the endosomal system is supported by genetic interactions between VPS8 and YPT51(Horazdovsky et al., 1996) as well as betweenVPS8 and PEP5/END1 (Woolford et al., 1998). Nevertheless, we cannot formally rule out the possibility that mutant Pma1 moves to the surface directly from the Golgi invps8 mutants. Similarly, we cannot exclude the possibility that mutant Pma1 undergoes retrograde transport from the endosome to the Golgi followed by delivery to the surface. (On the other hand, because CPY is missorted in vps8Δ cells [Chen and Stevens, 1996; Horazdovsky et al., 1996], recycling of Vps10 back to the Golgi is likely impaired.)

Our hypothesis is summarized in Figure 10, in which proposed impaired step(s) in transport from the Golgi to the vacuole are labeled with the corresponding vps mutations. We propose that invps8 cells, there is an accumulation within early endosomes of proteins entering either from the cell surface (Ste3) or from the biosynthetic pathway (Vps10 and mutant Pma1). As a consequence, there is increased transport to the cell surface. Because plasma membrane delivery of Pma1-7 in class E vps mutants appears less efficient than in vps8 cells, it is possible that movement to the surface from the prevacuolar or late endosome compartment occurs via an early endosome intermediate.

In mammalian cells, an endosome-to-surface traffic pathway is taken constitutively by many internalized membrane proteins and lipids. In some cell types, the pathway is specialized for recycling synaptic vesicle components, antigen presentation, insulin-dependent control of glucose transport, and transcytosis. Indeed, some newly synthesized proteins are normally delivered to the plasma membrane via the endosome (Futter et al., 1995; Leitinger et al., 1995). Recent observations suggest that there is possibly analogous endosome-to-surface trafficking in yeast. Analysis of the chitin synthase Chs3 has led to the proposal that its distribution between the plasma membrane and the “chitosome,” an endosomal compartment, is regulated by recycling (Chuang and Schekman, 1996; Ziman et al., 1998). Work on copper entry into the yeast secretory pathway has suggested that copper loading of the surface enzyme ceruloplasmin occurs within an endosomal compartment (Yuan et al., 1997). Thus, selected plasma membrane proteins may pass through endosomes to fulfill specific processing requirements. Our finding that mutant Pma1 can move from the endosomal system to the plasma membrane represents a first step toward characterizing an endosome-to-surface trafficking pathway in yeast. Future work should focus on elucidating the physiological significance of this pathway.

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ACKNOWLEDGMENTS

We thank Jim Haber, Nick Davis, George Sprague, Hugh Pelham, Steve Nothwehr, Tamara Doering, Birgit Singer-Kruger, Carolyn Slayman, and Scott Emr for strains, plasmids, antibodies, and advice. Thanks to Peter Arvan for reading the manuscript. This work was supported by grant GM58212 from the National Institutes of Health.

REFERENCES

Most viruses are either helical or icosahedral in structure. The two highly symmetric shapes permit viruses to use the same component protein multiple times to create large structures from a minimum number of distinct protein species. The strategy conserves the amount of genetic material viruses need to encode structural proteins.

Although the two basic shapes serve the needs of viruses more or less equally well, structural biologists have had a much easier time determining the structures of the icosahedra. For example, while more than one hundred high resolution structures of icosahedral viruses are now available, the number of comparable helical virus structures is limited to helical plant viruses such as tobacco mosaic virus and filamentous bacteriophage such as E. coli phage f1.

It’s not as though helical animal and human viruses are of limited interest. Just the opposite. They include influenza virus plus members of the paramyxo-, rhabdo- bunya-, corona, filo- and arenavirus families, all of which contain important human pathogens. The problem is that structural analysis of these viruses is unusually difficult. The protein-RNA complex is often disordered or weakly ordered in the virion, and the viruses have a membrane, a structure that complicates both crystallization and electron microscopic analysis.

To advance our knowledge of helical virus structure, investigators have focused their attention on the rhabdoviruses, a family of bullet-shaped viruses that includes rabies and vesicular stomatitis viruses (VSV). Rhabdoviruses have a helical nucleocapsid that is well ordered over most of the virion length. Although a membrane is present, it is tightly wrapped around the nucleocapsid, and does not obscure the helix in electron micrographs of the virion (see images of VSV in Figures 1a and 1b). With such excellent images, one would think it would be a simple matter to compute a three-dimensional reconstruction by standard Fourier-Bessel-based methods. No structure has been forthcoming, however, despite the best efforts of many highly talented structural biologists—until now.

Figure 1. Electron micrographs illustrating VSV structure and assembly. (a) and (b) show virions in negative stained and thin section preparations, respectively. Note that individual turns of the nucleocapsid helix can be seen in both cases. (c) shows virions in the process of assembling at the host cell cytoplasmic membrane. Note that nascent virions bud beginning at the domed end.

Figure 1. Electron micrographs illustrating VSV structure and assembly. (a) and (b) show virions in negative stained and thin section preparations, respectively. Note that individual turns of the nucleocapsid helix can be seen in both cases. (c) shows virions in the process of assembling at the host cell cytoplasmic membrane. Note that nascent virions bud beginning at the domed end.

Using electron micrographs of VSV preserved in the frozen-hydrated state and a real space reconstruction method, Ge et al. [1] have computed the VSV structure at 10.6Å resolution. The results are wonderful. The structure shows a wealth of detail about the nucleocapsid helix, interaction of the nucleocapsid protein (N) with the overlying layer of M protein and contacts between M and the membrane glycoprotein. Unique features at the virion ends are described and there are implications for rhabdovirus assembly. Here we briefly discuss the methods used to compute the reconstruction, the new features revealed about VSV morphology and what this singular accomplishment may mean for future analyses of helical virus structures.

Prior knowledge of VSV composition and assembly [2]

VSV is a typical rhabdovirus with a bullet shape, a length of ~190 nm and a diameter of 85 nm. At the core of the virion is the minus sense ssRNA genome bound to N protein. The RNA is 11,161 nucleotides in length, and it is tightly but non-covalently attached to N protein (MW 47.5 kDa; 422 amino acids) creating the nucleocapsid. Each N molecule encapsidates nine nucleotides of the RNA with 1240 N molecules expected in the overall helical structure. In the intact virion, the nucleocapsid helix is organized into ~35 helical turns with ~38 N protein subunits/turn in the trunk region (but smaller numbers in the domed end).

The entire nucleocapsid is enclosed in a mono-molecular layer of the matrix protein, M (29 kDa; 229 amino acids; 1826 molecules/virion). M protein is thought to create the precise structure of the nucleocapsid helix in the virion as the nucleocapsid is un-structured in the absence of M [3]. In addition to wrapping the nucleocapsid helix, M also makes contact with the virion envelope membrane. Embedded within the membrane are ~400 trimers of the virion glycoprotein required for entry of the virus into a host cell.

VSV assembles by budding at the host cell cytoplasmic membrane. Assembly is initiated by interaction of the nucleocapsid with a specialized region of membrane containing M and G proteins [4]. M and the membrane then bind to the nucleocapsid progressively creating helical turns beginning at the domed virion end. As helical turns are created, the overall structure projects progressively further outward from the host cell (Figure 1c). Assembly is terminated with formation of the blunt end and detachment of the complete virion from the host cell.

Method of reconstruction

Ge et al. [1] determined the VSV structure beginning with cryo-electron micrographs of virions imaged at a magnification of 98,000X. Reconstruction focused on the virion trunk, and was computed in two steps. First, the helical parameters were determined by: (a) measuring the pitch of the helix from layer lines in the Fourier transform of the trunk; and (b) two-dimensional classification of trunk images to determine the number of N protein molecules per helical turn. The latter method demonstrated that each N molecule is located between two others in the turn above and in the turn below, an observation showing there are two helical turns in each repeat unit (see Figure 2a). The exact number of N molecules per turn was determined by testing candidate numbers in the reconstruction as described below.

Second, the reconstruction was computed using the iterative helical real space reconstruction (IHRSR) method developed by our colleague Ed Egelman at the University of Virginia [5]. A starting model was created by classifying trunk images according to their mutual similarity. An average of the class averages was then used to center and align individual particles for determination of their rotational angle, the key step in creation of the starting model. Starting angles were identified by the degree of fit with templates calculated at 4° intervals. Angular classes were then averaged, back projected and the helical parameters applied to create an initial model. The initial model was then refined iteratively until it converged on a solution. A total of 644 trunk images were used in the reconstruction reported.

Figure 2. Interpretive drawings illustrating the new VSV structure. (a) shows an external view of the N (red) and M (blue) helices as they are found in the trunk region of the mature virion. A portion of the M helix has been removed to illustrate the underlying N helix. The membrane and the glycoproteins are not illustrated. Note that N molecules in one helical turn lie between two N molecules in the turn above. Note also that: (i) the M helix lies between turns of the N helix linking the two N turns together; (ii) there is one M molecule for each N; (iii) M molecules make lateral connections with each other providing stability to the overall structure; and (iii) N protein molecules are tilted slightly upward (27°) with respect to horizontal plane. (b) shows a cross-sectional view of the virion in the trunk region. Note that there are 37.5 N molecules in each helical turn and N protein molecules are connected externally by M and in the middle by the virus RNA (black).

Figure 2. Interpretive drawings illustrating the new VSV structure. (a) shows an external view of the N (red) and M (blue) helices as they are found in the trunk region of the mature virion. A portion of the M helix has been removed to illustrate the underlying N helix. The membrane and the glycoproteins are not illustrated. Note that N molecules in one helical turn lie between two N molecules in the turn above. Note also that: (i) the M helix lies between turns of the N helix linking the two N turns together; (ii) there is one M molecule for each N; (iii) M molecules make lateral connections with each other providing stability to the overall structure; and (iii) N protein molecules are tilted slightly upward (27°) with respect to horizontal plane. (b) shows a cross-sectional view of the virion in the trunk region. Note that there are 37.5 N molecules in each helical turn and N protein molecules are connected externally by M and in the middle by the virus RNA (black).

The reliability of the reconstruction was tested by docking two available X-ray crystallographic structures, the N protein-RNA complex [6] and the C-terminal domain of M [7,8]. Both were found to fit well with candidate regions of the cryo-EM volume, a result that authenticates the structure and at the same time allows the N and M proteins to be identified reliably. Docking of the N-RNA complex permitted the directionality of the RNA to be established; the 5’ RNA end was found to be located at the domed end of the virion.

The structure

VSV structure was shown to consist of two, single, concentric helices, one composed of N protein and RNA (the nucleocapsid) and the other a helically arrayed layer of M. The nucleocapsid helix has outer and inner diameters of 45.0 nm and 30.8 nm, respectively, with layers spaced 5.08 nm apart along the helical axis. There are 37.5 N protein molecules per turn with 75 in the repeating unit (Figure 2b). Each N protein is tilted 27° with respect to the horizontal plane, an un-anticipated feature of the new structure. Individual turns of the nucleocapsid are not tightly bound to each other, a finding that rationalizes the observation that the N-RNA helix is un-structured in the absence of M protein [9].

Each turn of the M protein helix lies in the space between two turns of the nucleocapsid helix making contact with both and holding them together (Figure 2a). There is one M molecule for each N in the overall M protein helix. Individual M protein molecules are found in a U shape with arms called the M-hub and the M-protein-C-terminal domain (MCTD), respectively. The M-hub makes contact with the upper and lower turns of the nucleocapsid helix as illustrated in Figure 2a. The MCTD extends outward and at an angle from the ribonucleocapsid helix in a position to make contact with the virus membrane. Thin projections from the membrane are found to contact the MCTD, and these are interpreted as C-terminal tails from the virus glycoprotein.

In order for rhabdovirus RNA synthesis to take place in vitro, virions need to be disrupted with detergent to solubilize the membrane [10]. The new VSV structure suggests disruption may be required to loosen the M protein helix. This would allow the ribonucleocapsid to flex permitting the RNA-dependent RNA polymerase (L protein) to gain access to the template RNA in the nucleocapsid.

A long-standing enigma of rhabdovirus biology is how the viral RNA can function as a template for RNA synthesis despite being tightly bound by N protein. Available evidence indicates that in the nucleocapsid, RNA is resistant to nuclease digestion, even when transcription is in progress [11]. The N structure in complex with RNA shows that N is bi-lobed and that the RNA is sequestered in a 2.0 X 1.0 nm cavity between the two lobes [6]. It is reasonable to expect that the RNA would be protected from nucleases inside the cavity. The nucleocapsid structure, however, does not provide sufficient space to accommodate the large RNA-dependent RNA polymerase molecule. Domain movement to ”open” the two lobes of the N protein has been proposed as a means by which the polymerase may gain access to RNA [12]. Removal of the M protein holding the ribonucleocapsid in place may facilitate N domain movement and allow the polymerase to gain access to the RNA template.

Questions remaining to be answered

The location of the G protein was not revealed in the reconstruction reported. This result was expected, and suggests that the glycoprotein trimers are not arranged with the same helical symmetry as the nucleocapsid-M complex. Further analysis will be required to localize the glycoprotein. Although the new structure accounts for one M protein bound to each N, there is additional M protein in the virion. The experimental copy number for M is 1826 [3], so the number of un-located M molecules is 1826 minus 1240 or 586 molecules, approximately 1/3 of the total M. Possible locations for the missing M protein include the axial channel of the virion and the virion ends, places where M may not be helically arranged.

In addition to N, M and G, VS virions contain P and L proteins, the remaining two proteins encoded in the genome. L is the RNA-dependent RNA polymerase responsible for replication and transcription of the VSV genome while P tethers L to the nucleocapsid. Both proteins are considered to be located in the axial channel of the VS virion, but no candidate features were seen in the current reconstructed volume. There is good evidence for binding of P to the N protein [12], and L is thought to be bound to P. The two may not have been seen in the reconstruction because they are not localized at the same place in each virion or perhaps because of the low copy numbers of L (60/virion) and P (466/virion). Localizing L and P is an important, but challenging problem for future studies.

Future prospects

In view of the success with VSV, one can expect that the methods employed by Ge et al. [1] will soon be focused on other helical viruses. Filoviruses such as Ebola virus are attractive candidates because of their extended helical regions. Domains of ordered nucleocapsid are often seen in influenza virus, and these suggest themselves as candidates for reconstruction. Other possibilities include the paramyxo-, bunya-, corona- and arena- viruses mentioned above.

Like other important structural advances, the work reported here makes very specific predictions about the amino acid sequences involved in protein-protein contacts. M-N, M-M and M-G contacts are examples. Such predictions are amenable to experimental testing with mutants, and one can anticipate that relevant mutational analyses will be forthcoming; they have the potential to authenticate the structure and add to its usefulness. In fact, a start on this effort is reported by Ge et al. [1].

For now it is worth taking time to enjoy the new VSV structure and to congratulate the authors for their skill and ingenuity in computing it. Such reflection may allow us to hope that at last we have the tools to make further progress into understanding the structures of helical viruses.

Acknowledgments

We gratefully acknowledge Anna Maria Copeland for comments on the manuscript. Our work is supported by NIH awards R01AI041644 (JCB), R01AI12464 (GWW), and R37AI12464 (GWW).

References

  1. Ge, P.; Tsao, J.; Schein, S.; Green, T.J.; Luo, M.; Zhou, Z.H. Cryo-EM model of the bullet-shaped vesicular stomatitis virus . Science2010, 327, 689–693. [Google Scholar] [CrossRef] [PubMed]
  2. Rose, J.K.; Whitt, M.A. Rhabdoviruses: The Viruses and their Replication. In Fields Virology; Knipe, D.M., Howley, P.M., Eds.; Lippincott Williams and Wilkins: PA, USA, 2001; pp. 1221–1244. [Google Scholar]
  3. Thomas, D.; Newcomb, W.W.; Brown, J.C.; Wall, J.S.; Hainfeld, J.F.; Trus, B.L.; Steven, A.C. Mass and molecular composition of vesicular stomatitis virus: a scanning transmission electron microscopy analysis. J. Virol.1985, 54, 598–607. [Google Scholar] [PubMed]
  4. Swinteck, B.D.; Lyles, D.S. Plasma membrane microdomains containing vesicular stomatitis virus M protein are separate from microdomains containing G protein and nucleocapsids. J. Virol.2008, 82, 5536–5547. [Google Scholar] [CrossRef] [PubMed]
  5. Egelman, E.H. The iterative helical real space reconstruction method: surmounting the problems posed by real polymers. J. Struct. Biol.2007, 157, 83–94. [Google Scholar] [CrossRef] [PubMed]
  6. Green, T.J.; Zhang, X.; Wertz, G.W.; Luo, M. Structure of the vesicular stomatitis virus nucleoprotein-RNA complex. Science2006, 313, 357–360. [Google Scholar] [CrossRef] [PubMed]
  7. Graham, S.C.; Assenberg, R.; Delmas, O.; Verma, A.; Gholami, A.; Talbi, C.; Owens, R.J.; Stuart, D.I.; Grimes, J.M.; Bourhy, H. Rhabdovirus matrix protein structures reveal a novel mode of self-association . PLoS Pathog.2008, 4, e1000251. [Google Scholar] [CrossRef] [PubMed]
  8. Gaudier, M.; Gaudin, Y.; Knossow, M. Crystal structure of vesicular stomatitis virus matrix protein. EMBO J.2002, 21, 2886–2892. [Google Scholar] [CrossRef] [PubMed]
  9. Newcomb, W.W.; Brown, J.C. Role of the vesicular stomatitis virus matrix protein in maintaining the viral nucleocapsid in the condensed form found in native virions. J. Virol.1981, 39, 295–299. [Google Scholar] [PubMed]
  10. Baltimore, D.; Huang, A.S.; Stampfer, M. Ribonucleic acid synthesis of vesicular stomatitis virus, II. An RNA polymerase in the virion. Proc. Natl. Acad. Sci. USA1970, 66, 572–576. [Google Scholar] [CrossRef]
  11. Hill, V.M.; Simonsen, C.C.; Summers, D.F. Characterization of vesicular stomatitis virus replicating complexes isolated in renografin gradients. Virology1979, 99, 75–83. [Google Scholar

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