INTRODUCTION

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The influenza A virus genome consists of eight segments of negative-sense single-stranded RNA (vRNA) which are transcribed in infected cells to yield two types of positive-sense transcripts: capped and polyadenylated mRNAs and exact complements (cRNA) which serve as replicative intermediate RNAs for the production of further vRNAs (18). Four viral proteins are necessary and sufficient to carry out this process (16): the three subunits of an RNA-dependent RNA polymerase (PB1, PB2, and PA) and a 55-kDa single-stranded RNA-binding nucleoprotein (NP). Indeed, the v- and cRNA segments are always associated with these polypeptides to form ribonucleoprotein (RNP) structures, thought to comprise one copy of the trimeric polymerase and approximately one NP polypeptide per 20 bases of RNA (18).

Unusually for a virus with no DNA coding stage, influenza virus transcription occurs in the nuclei of infected cells (15). Thus, on initiation of infection the incoming RNPs are targeted to the nucleus, and during infection newly synthesized RNP proteins also undergo nuclear import. Consistent with this, nuclear localization signals (NLSs) have been identified in all four transcriptase proteins (9, 25, 26-28, 41, 41a), including three in NP, the major protein component of RNPs. At early times postinfection, the polymerase subunits and NP accumulate in the nucleus (reviewed in reference 18), and NP has been shown to be necessary for the nuclear import of vRNA in permeabilized cells (30). However, virion assembly, including packaging of progeny RNPs, occurs in the cytoplasm and at later times this is reflected in the cytoplasmic accumulation of the four transcriptase proteins, generally assumed to be in the form of RNPs (18). This differential localization of RNPs necessitates regulatory mechanisms, and evidence points to the involvement of at least three viral polypeptides: the virion matrix (M1) protein, the minor virion component NS2, and NP itself. M1 is capable of binding to membranes, RNA, NS2, and RNPs. In the virion, it is thought to act as the link between the lipid envelope and the packaged genome segments (44). Moreover, M1 itself can localize to the nucleus, where it is thought to promote the export of RNPs (22, 43). NS2, which binds to RNPs via the M1 protein (47), has recently been shown to possess a nuclear export signal (31). Microinjected anti-NS2 antibodies inhibit export of RNPs, and it has therefore been proposed that NS2 is the viral factor directly responsible for the export of RNPs from the nucleus (31). Accordingly, import of NP (and RNPs early in infection) can be postulated to occur because of the presence of NLSs and export can be postulated to occur because of the subsequent expression of components (M1 and NS2) that target RNPs to an export pathway (31, 44). However, this hypothesis does not fully explain why RNPs accumulate in the cytoplasm in the absence of M1 export (33, 42) or why a mutant influenza virus which synthesizes very little or no NS2 is fully replication competent (36, 45). Moreover, since vRNA synthesis is not temporally separated from RNP export, it is not clear how import of newly synthesized NP is maintained in the presence of active export of the same polypeptide in the form of RNP complexes, or why cytoplasmic RNPs are not simply reimported back into the nucleus by virtue of their polyvalent NLS sequences. Therefore, there must be other elements involved in the control of RNP localization.

NP possesses the ability to locate to both cytoplasm and nucleus. In the absence of other viral proteins, it shuttles between the two compartments (43), which suggests that it interacts with the cellular nuclear export apparatus in the absence of M1 and NS2. NP can also accumulate in the cytoplasm at "late" times postexpression (27), indicating that the activities of the NP NLSs are modulatable. Therefore, RNP localization may also be regulated by mechanisms acting through NP. One well-documented way of modulating nuclear import is through interactions with cytoplasmic anchoring proteins (29, 40), and recent work involving cell fractionation and fluorescent staining experiments has suggested that cytoplasmic NP is associated with the cytoskeleton (3, 17).

Here we demonstrate that purified NP shows high-affinity binding to filamentous actin (F-actin) in vitro with a K_d on the order of 1 µM and a stoichiometry of 1 per actin subunit. Additionally, there is a further interaction of lower affinity which may reflect oligomerization of NP itself. We also show the formation of complexes containing NP, RNA, and actin and have identified a cluster of amino acid residues in NP important for the interaction with F-actin. Localization of exogenously expressed NP was found to depend on the amount expressed, with small amounts entering the nucleus and large amounts accumulating in the cytoplasm, where the protein colocalizes with F-actin. Mutations in NP which decrease F-actin binding change the cellular distribution of the protein, favoring accumulation in the nuclei even when the expression level is high. We propose that F-actin binding acts as a cytoplasmic retention signal for NP.

	MATERIALS AND METHODS

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Plasmids, antisera, and viruses. A cDNA copy of A/PR8/34 segment 5 was excised from plasmid pVB5+ (6) with EcoRI and ligated into similarly linearized plasmid pMAL-c2 (New England Biolabs). The NP and maltose binding protein (MBP) open reading frames (ORFs) were then fused by oligonucleotide-directed mutagenesis. The resulting plasmid (pMAL-NP) contains the lac promoter, the MBP gene, a factor Xa protease recognition site, an alanine codon, and the NP ORF (with an NcoI restriction enzyme site flanking the initiation codon). Point mutations were introduced into this plasmid by standard procedures. All mutations were confirmed by nucleotide sequencing of the appropriate region of the NP gene. To construct a glutathione S-transferase (GST)-NP fusion protein, plasmid pMAL-NP was digested with NcoI, end filled with the Klenow fragment of DNA polymerase, and digested with EcoRI. The DNA fragment containing the NP ORF was then ligated into SmaI-EcoRI-cut plasmid pGEX-2T (Pharmacia) to create plasmid pGEX-NP. To obtain eukaryotic expression of nonfused NP, NP genes were excised from the pMAL constructs by digestion with NcoI and XbaI and inserted into similarly digested plasmid pKT0 (39). The resulting plasmids (pKT5 series) contain the NP gene under control of a T7 RNA polymerase promoter.

Expression and purification of NP. Exponentially growing cultures of Escherichia coli TG1 containing plasmid pMAL-NP, pGEX-NP, or pGEX-2T were induced by the addition of isopropyl-thiol--galactosidase and incubated with shaking for a further 4 h. Bacteria were pelleted by centrifugation and resuspended in a 1/5 culture volume of amylose column buffer (ACB) (500 mM NaCl, 20 mM Tris-Cl [pH 7.6], 1 mM EDTA, 0.02% sodium azide) (pMAL) or phosphate-buffered saline (PBS) (pGEX), containing 0.25 µg of lysozyme per ml. After incubation for 30 min on ice and overnight at 20°C, the suspension was thawed and sonicated before centrifugation at an average of 9,000 × g for 30 min. The supernatants were sequentially filtered through Whatman no. 1 filter paper and a 0.45-µm-pore-size cellulose acetate filter before chromatography over amylose resin (New England Biolabs) or glutathione Sepharose (Pharmacia) as appropriate. Columns were washed sequentially with 10 volumes of ACB or PBS, 10 volumes of buffer containing 2 M NaCl, and another 10 volumes of ACB or PBS. Bound protein was eluted with buffer supplemented with 10 mM maltose or 50 mM reduced glutathione.

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Actin-binding assays. Purified rabbit muscle actin (38) was polymerized in 100 mM NaCl-10 mM Tris-Cl (pH 8.0)-1 mM MgCl₂-0.1 mM ATP-0.2 mM EGTA-1 mM sodium azide (F buffer). Sedimentation assay mixtures contained 3 µM actin in 50 mM NaCl-10 mM HEPES (pH 7.6)-1 mM MgCl₂-10% glycerol-0.1 mM EDTA-0.1 mM ATP in a final volume of 100 µl. After being mixed, the samples were centrifuged at an average of 200,000 × g for 20 min at 20°C before separation into pellet and supernatant fractions. Equivalent amounts of the two fractions were analyzed by SDS-PAGE and stained with Coomassie brilliant blue dye. Quantitation was performed by using a Molecular Dynamics computing densitometer (32). For electron microscopy, actin (1 µM) and MBP or MBP-NP were mixed in F buffer and left on ice for 20 min. Drops (30 µl) were applied to carbon-coated grids for 1 min and then grids were rinsed with 5 drops of F buffer and 5 drops of 1% uranyl acetate before being blotted dry and viewed in a Philips 208S electron microscope at a magnification of 20,000 or 30,000.

RNA transcription and binding assays. For in vitro RNA binding assays, a radiolabelled 178-nucleotide synthetic RNA target was generated by in vitro transcription of plasmid pKT8-3'5' (generous gift of K. Tibbles) with bacteriophage T7 RNA polymerase in the presence of -[³²P]CTP (100 µM; specific activity, 2 Ci/mmol) according to standard procedures. The transcript corresponds to influenza virus A/PR8/34 segment 8 but with nucleotides 84 to 795 (inclusive) deleted and a C-to-A transversion of the penultimate base. Shorter transcripts corresponding to the minimal terminal repeat ("panhandle") sequences or only the 3' end of segment 8 were generated by in vitro transcription of the appropriate oligonucleotide templates cloned into plasmid pNEB193 (New England Biolabs). For in vivo RNA transcription assays, a synthetic influenza virus genome segment containing an antisense chloramphenicol acetyltransferase (CAT) gene was produced by in vitro transcription of plasmid pPB2CAT (generous gift of Mark Krystal). Filter binding assays were performed by incubating protein samples with 20 fmol of RNA (around 5,000 cpm) in 25 mM Tris-Cl (pH 7.6)-50 mM NaCl-5 mM MgCl₂-0.5 mM dithiothreitol-5% glycerol at room temperature for 20 min. The reaction mixtures were passed through nitrocellulose filters equilibrated in 20 mM Tris-Cl (pH 7.6)-50 mM NaCl and washed three times with 200 µl of the same buffer. Bound radioactivity was quantified by liquid scintillation counting.

Transfection of tissue culture cells, transient replication assays, and indirect immunofluorescence. BHK cells (in 35-mm-diameter dishes) were infected with recombinant vaccinia viruses at a multiplicity of infection of 5 of each virus per cell for 2 h at 37°C. The cells were washed three times with serum-free medium before transfection with 1 µg of plasmid DNA encoding NP and 0.5 µg of pPB2CAT9 in vitro-transcribed RNA and 10 µg of a cationic liposome mixture (Lipofectin [GIBCO-BRL] or Escort [Sigma-Aldrich]), according to the manufacturers' instructions. The cells were incubated at 37°C for 20 h, washed three times with ice-cold PBS, and solubilized in 1 ml of CAT ELISA lysis buffer (Boehringer Mannheim). The lysate was clarified by centrifugation at 14,000 × g for 10 min at 4°C, and CAT expression was quantified relative to known standards by a commercial enzyme-linked immunosorbent assay (Boehringer Mannheim). Replication activities of mutants were calculated as percentages of the CAT synthesis seen with the wild-type (WT) gene after subtraction of the background obtained in the absence of NP. Values were calculated from a minimum of three separate experiments and two independent plasmid DNA preparations.

	RESULTS

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Binding of NP to F-actin. We tested whether purified NP bound F-actin in vitro by a cosedimentation assay, in which F-actin readily separates from monomeric protein by centrifugation. As shown in Fig. 1, most of the actin sedimented after centrifugation (lane 2) and only the expected critical actin monomer concentration (0.1 to 0.2 µM) remained in the supernatant (lane 1). Control assays showed that only trace amounts of the expressed proteins pelleted in the absence of actin (Fig. 1, lanes 4, 8, and 12). However, when GST, GST-NP, or MBP-NP was sedimented after mixing with F-actin, the NP-containing polypeptides partitioned approximately evenly between the supernatant and pellet fractions (Fig. 1, lanes 9, 10, 13, 14) while GST did not (lanes 5 and 6), demonstrating association of NP with actin filaments. To test the ability of nonfused NP to bind F-actin, MBP-NP was cleaved into MBP and NP moieties by digestion with factor Xa protease and sedimented with or without F-actin. Since MBP and actin comigrate on SDS-PAGE, the samples were analyzed in parallel by Western blotting with an anti-MBP serum, confirming that MBP alone did not pellet whether or not actin was present (Fig. 1, inset, lanes 15 to 18). However, the NP polypeptide pelleted only in the presence of actin (Fig. 1, lanes 15 to 18). Thus, NP binds F-actin in vitro.

View larger version (28K): [in this window] [in a new window]   FIG. 1.   F-actin-binding activities of GST, GST-NP, and MBP-NP. The proteins (MBP/NP corresponds to MBP-NP digested with factor Xa protease) were centrifuged in the presence (+) or absence () of 3 µM F-actin. Supernatant (S) and pellet (P) fractions were analyzed by SDS-PAGE and staining with Coomassie blue or by Western blotting with anti-MBP serum (inset, lanes 15 to 18). Arrows indicate the named polypeptides (right), and molecular mass markers (in kilodaltons) are indicated on the left.

View larger version (12K): [in this window] [in a new window]   FIG. 2.   Titration binding curves of WT and R156-A MBP-NP for F-actin. Sedimentation assays were performed by using 3 µM F-actin and WT (open circles) or mutant R156-A (closed circles) MBP-NP. The amount of bound MBP-NP (expressed as the molar ratio relative to actin) is plotted against the concentration of free MBP-NP.

View larger version (139K): [in this window] [in a new window]   FIG. 3.   Electron microscopy of actin filaments mixed with equimolar concentrations of MBP (A) and MBP-NP (B and D) and a threefold excess of MBP-NP (C). Scale bars, 300 nm.

RNA and the NP-actin interaction. Since NP is a single-stranded-RNA-binding protein, we investigated the relationship between actin binding and RNA binding. First, we examined the effect of actin on the ability of NP to bind RNA in solution by titrating a radiolabelled 178-nucleotide RNA corresponding to the 5' and 3' noncoding sequences from vRNA segment 8 with increasing concentrations of MBP, MBP-NP, and MBP-NP plus 3 µM actin. Addition of MBP up to a concentration of 300 nM did not result in significant retention of the labelled probe, indicating that MBP itself does not bind RNA (Fig. 4a). However, titration with increasing amounts of MBP-NP resulted in the retention of 50% of the input RNA with around 20 nM protein (Fig. 4a). In replicate experiments, the affinity of MBP-NP for RNA was determined to be 16.3 ± 3.8 nM¹ (n = 5), which is in good agreement with that estimated for NP purified from influenza virus virions (20 nM¹ [4]). Therefore, the bacterially expressed NP behaved similarly to native NP and was fully competent to bind RNA. Moreover, addition of F-actin to the reaction mixtures had no significant effect on this RNA-binding activity (Fig. 4a).

View larger version (14K): [in this window] [in a new window]   FIG. 4.   RNA and the interaction between NP and F-actin. (a) Effect of actin on the RNA-binding activity of NP. The amounts of radiolabelled RNA (1 nM) bound by MBP (circles), MBP-NP (squares), and MBP-NP plus 3 µM F-actin (triangles) were determined by a filter assay. (b) Effect of RNA on actin binding by NP. MBP-NP (0.75 µM) was cosedimented in the presence (+) or absence () of 3 µM F-actin with the indicated molar ratio of a 34-mer RNA. S, supernatant; P, pellet. (c) Cosedimentation of RNA with actin and NP. Radiolabelled 34-mer (100 nM in 100 µl; open bars) or 54-mer (50 nM; shaded bars) was cosedimented in the presence of 3 µM MBP-NP with or without 3 µM F-actin.

Mutational analysis of actin binding. To further delineate the NP-F-actin interaction, we performed a mutational analysis of NP. On the grounds that an interaction with a relatively invariant protein such as actin might be reflected by sequence conservation within NP, we altered residues in regions conserved among the NPs of influenza A, B, C, and Dhori viruses (12) (summarized in Fig. 5b) and tested the effects of the mutations on actin-binding activity. Many of the mutations had no effect on the interaction. Quantitative analysis of the R156-A mutant produced a binding curve indistinguishable from that of the WT polypeptide (Fig. 2). Similarly, when tested at concentrations in the low micromolar range, the W104-F and R391-A mutants bound efficiently to F-actin (Fig. 5a, panels ii and vi, respectively; cf. panel i [WT]). The binding affinities for F-actin of these and the R8-A, W120-A, R199-A, K236-A, and R267-A mutants were found to vary from that of the WT protein by less than threefold (Fig. 5b). However, the F338-A, E339/D340-AA, R342-A, and Q405-A polypeptides showed markedly different behavior, with the majority of each protein remaining in the supernatant fractions at each concentration tested (Fig. 5a, panels iii, iv, v, and vii). Quantification produced estimated binding constants for F-actin some 10- to 35-fold lower than for the WT protein (Fig. 5b). Thus, four mutations that disrupted the ability of the protein to bind F-actin were identified in the C-terminal one-third of NP.

View larger version (28K): [in this window] [in a new window]   FIG. 5.   Identification of NP point mutations which disrupt actin binding. (a) WT or mutant MBP-NP polypeptides were cosedimented in the absence () or presence (+) of 3 µM F-actin, and supernatant (S) and pellet (P) fractions were examined by SDS-PAGE. (b) Summary of point mutations and cellular localization signals in NP. Arrows indicate the positions of amino acid changes introduced into NP. White arrows indicate mutations with no major effect on F-actin binding and black arrows indicate those that substantially reduced binding. Tabulated values are the fold decrease in binding affinity for F-actin relative to values for the WT protein. Also shown are the positions of the three postulated cellular localization signals that have been identified in NP.

Ability of actin-binding mutants to support viral transcription. Although we had identified mutations that reduced the ability of NP to bind actin, it was necessary to examine whether the mutations affected other functional properties of NP. Many lines of evidence indicate that NP is essential for virus RNA synthesis (18). This has been confirmed in a recombinant system where NP and the three subunits of the influenza virus RNA polymerase are necessary and sufficient to drive transcription and replication of a synthetic influenza virus genome segment containing an antisense CAT (flu-CAT) reporter gene (16). We used an adaptation of this assay to apply the stringent test of whether the mutants defective in actin binding could support virus RNA synthesis.

Actin and the intracellular localization of NP. Since F-actin is cytoplasmic and NP can also accumulate in the cytoplasm, we examined the relevance of actin binding to the localization of NP. Because exogenously expressed NP shuttles between nucleus and cytoplasm and has been shown to accumulate in the cytoplasm under certain circumstances (27, 43), we first established the localization parameters of the WT polypeptide in our expression system by testing the effect of expression levels on the compartmentalization of NP. BHK cells were infected with a recombinant vaccinia virus expressing T7 RNA polymerase and transfected with various amounts of plasmid pKT5, and the subsequent expression and cellular localization of NP were examined by indirect immunofluorescence at 4 h posttransfection. Nontransfected cells revealed only a diffuse, low-level fluorescence (data not shown). In contrast, cells transfected with a plasmid containing the WT NP gene exhibited strong fluorescence, the location of which depended on the amount of plasmid transfected. In cells transfected with a high dose of plasmid (0.3 µg/10⁵ cells), virtually all the fluorescence was cytoplasmic, often with apparent sparing of the nucleus (Fig. 6a). However, cells transfected with a low dose of plasmid (3 ng/10⁶ cells) showed almost exclusively nuclear fluorescence (Fig. 6c). Cells transfected with an intermediate amount of plasmid (30 ng/10⁵ cells) showed an intermediate pattern, with many cells containing substantial amounts of cytoplasmic NP but with nuclear accumulation as well (Fig. 6b). When lysates from cells transfected in parallel were subjected to SDS-PAGE and Western blot analysis to assess the quantity of NP expressed, the amount correlated with the plasmid dose (data not shown). The transfection efficiency varied by less than twofold (generally ranging between 20 and 40%; data not shown) over the range of plasmid concentrations tested. This indicates that the expression levels depended primarily on the amount of plasmid received per cell and not the number of cells transfected. Thus, the cellular distribution of NP depended on the amount (or rate) of synthesis of protein within any one cell.

View larger version (47K): [in this window] [in a new window]   FIG. 6.   Cellular localization of WT and mutant NP polypeptides. BHK cells (105) were infected with a recombinant vaccinia virus expressing T7 RNA polymerase, transfected with the indicated amounts of plasmids, and examined 4 h later by indirect immunofluorescence with anti-NP serum.

View larger version (80K): [in this window] [in a new window]   FIG. 7.   Colocalization of NP and F-actin. Cells were transfected with 0.3 µg of pKT5 and examined 4 h later by confocal laser microscopy after staining for NP (red) or actin (green). Panels b, e, and h are images resulting from the merger of the red and green channels. Panels a through f are from HeLa cells fixed before detergent extraction. Panels g through i are from a BHK cell extracted with detergent before fixation.

View larger version (85K): [in this window] [in a new window]   FIG. 8.   Cellular location of NP mutants bearing amino acid substitutions in NLS I. Cells transfected with 0.3 µg (except for those shown in panel b, which were transfected with 3 ng) of plasmids encoding the indicated mutants were examined by indirect immunofluorescence with anti-NP serum.

	DISCUSSION

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Two recent studies suggested that influenza virus NP interacts with microfilaments of the host cell cytoskeleton late in infection (3, 17). Both reports found that NP partitioned into cytoskeleton-containing pools during biochemical fractionation of infected cells; that NP colocalized with microfilaments, especially at the cell periphery; and that disruption of microfilaments with cytochalasins altered the distribution of NP. Here, we confirm and extend these observations by using two methods to show that NP binds directly to F-actin in vitro (Fig. 1 to 3) with an affinity comparable to that of many cellular F-actin-binding proteins (35). We also show that NP associates with actin filaments in vivo in the absence of other influenza virus proteins (Fig. 7). Furthermore, complexes containing F-actin, NP, and RNA could be formed (Fig. 4). As the in vitro actin-binding assay employed high-speed centrifugation, we could not directly test whether authentic RNP particles bound F-actin because they sediment under these conditions, but we predict that RNP structures bind F-actin.

We have identified point mutations in NP which weaken its affinity for actin in vitro (Fig. 5) and decrease the extent of colocalization of the proteins in vivo (Fig. 6 and 8). These mutations were largely localized to a region of NP previously identified as important for the accumulation of the protein in the nuclei of Xenopus oocytes (9). The signal mapped by Davey and colleagues was one of the first NLSs proposed, but as subsequent NLSs are identified, it appears increasingly atypical. In sequence composition, it does not possess the small stretch of basic amino acids found in other viral proteins such as simian virus 40 large T antigen, or the bipartite basic NLS contained in many cellular proteins (10). In oocytes it acts as a sequence which causes the retention of NP that has diffused into the nucleus, rather than as a signal for the active uptake of the protein; hence it was called a "nuclear accumulation" signal (NAS) (9). Subsequently, the study of Wang et al. (41) found that this NAS was not essential for nuclear localization of NP in mammalian cells. Here we show that mutation of residues within the NAS actually increases the ability of NP to localize to the nuclei of mammalian cells (Fig. 6 and 8). The reason for the discrepancy between the activities of the NAS in mammalian and amphibian cells is not clear, but the fact that, unlike most somatic cells, the nuclei of amphibian oocytes have been shown to contain actin may be relevant (8, 34). Nevertheless, the region of NP defined as the NAS, which we find important for actin binding, is one of the most highly conserved blocks of amino acids between influenza A, B, C, and Dhori virus NPs (12), suggesting that it is functionally important.

The most plausible role for the NP-actin interaction is during the late stages of viral infection, when NP is also found in the cytoplasm. In this study, we show that the cellular localization of exogenously expressed NP depends on the amount of protein synthesized. Low levels of NP were targeted efficiently to the nucleus, but in a dramatic switch, large amounts localized almost exclusively to the cytoplasm (Fig. 6), where in part NP associated with actin (Fig. 7). These phenomena could perhaps be explained by saturation of the nuclear import machinery leading to cytoplasmic accumulation of NP and subsequent F-actin binding. However, the fact that all the mutations that weakened F-actin binding restored nuclear import, even under conditions of high-level expression (Fig. 6) or in the absence of a functional N-terminal NLS (Fig. 8), strongly implies a causative link between F-actin binding and cytoplasmic retention of NP. We propose that the interaction between NP and actin plays a role in regulating the localization of RNPs by causing their cytoplasmic retention late in infection. Previous work has shown that nuclear import of NP and RNPs is driven by NLSs in NP and that export of RNPs depends at least in part on the subsequent expression of M1 and NS2. Mechanistically, retention of RNPs on cytosolic actin filaments would act in opposition to the import pathway, but in concert with the export pathway, to prevent reimport of exported RNPs destined for packaging into progeny virions.

Cytosolic anchoring as a means of regulating nuclear localization is well established for several cellular proteins, such as the transcription factors of the NF-B family, or Xenopus xnf7 (29, 40). In the case of NF-B, retention is thought to result from masking of the NLS by the cytoplasmic anchor, since the addition of an extra NLS overrides the retention mechanism (5). In contrast, Xenopus xnf7 remains cytoplasmic even after addition of another NLS, leading to the proposal that cytoplasmic retention results from a physical interaction with a fixed anchor, possibly the cytoskeleton (20). Here we show that direct interaction with actin microfilaments mediates cytoplasmic retention of an NLS-containing protein. This represents a novel mechanism for controlling the cellular location of a ribonucleoprotein complex and may be applicable to a wider range of cellular proteins and RNAs.

It seems likely that the interaction between NP and actin is subject to regulation. The quantity of actin in the cell is such that the switch in NP localization between low- and high-dose transfections cannot be explained purely in terms of the protein's affinity for actin. In addition, the nonuniform colocalization of NP with actin (Fig. 7) also suggests a regulated process. Since the microfilament network in cells consists of a polarized distribution of the various actin isoforms in complex with many different cellular actin-binding proteins (14), this could reflect a largely passive process, resulting from the occlusion of potential actin-binding sites and/or preferential binding to others. However, influenza virus may also have evolved positive mechanisms for regulating the NP-actin interaction, a likely candidate being phosphorylation of NP (2, 27). This is consistent with the effect of protein kinase inhibitors on NP localization noted by Neumann et al. (27), while phosphorylation is a well-documented mechanism for the control of the subcellular localization of cellular proteins, by modulation of both nuclear import (19, 21, 23) and actin binding (1, 24).