- Email: [email protected]
Interaction of the influenza virus nucleoprotein with F-actin
International Congress Series 1219 (2001) 503 – 512
Interaction of the inf luenza virus nucleoprotein with F-actin Paul Digard* , Debra Elton, Martha Simpson-Holley, Elizabeth Medcalf Division of Virology, Department of Pathology, University of Cambridge, Tennis Court Road, Cambridge CB2 1QP, UK
Abstract Introduction: The influenza virus genome is transcribed in the nucleus of infected cells but assembled into progeny virions in the cytoplasm. This is reflected in the cellular distribution of the virus nucleoprotein (NP), a protein which encapsidates genomic RNA to form ribonucleoprotein (RNP) structures: at the early stage post-infection, NP is found in the nucleus, but later it is found predominantly in the cytoplasm. The purpose of this study was to examine the possibility that cytoplasmic NP interacts with actin microfilaments. Methods: Bacterially expressed NP was tested for the ability to interact with filamentous (F)-actin in a variety of in vitro assays, and the localisation of actin and exogenously expressed NP in mammalian cells was examined microscopically. Results: Purified NP bound actin filaments in vitro and showed partial colocalisation with b-actin in vivo. Electron microscopy showed that NP induced bundling of actin fibres in vitro. In confirmation of this, NP caused a dramatic increase in the low-shear viscosity and light-scattering properties of Factin suspensions. Conclusions: NP binds F-actin in vitro and can alter the mechanical properties of actin filaments. This raises the possibility that influenza virus may use the host-cell cytoskeleton during virus replication. D 2001 Elsevier Science B.V. All rights reserved. Keywords: Microfilaments; Ribonucleoprotein; Bundling
1. Introduction The influenza A virus genome consists of eight segments of negative sense singlestranded RNA (vRNA), which are transcribed in infected cells to yield two types of positive
*
Corresponding author. Tel.: +44-1223-336918; fax: +44-1223-336926. E-mail address: [email protected] (P. Digard).
0531-5131/01/$ – see front matter D 2001 Elsevier Science B.V. All rights reserved. PII: S 0 5 3 1 - 5 1 3 1 ( 0 1 ) 0 0 6 2 8 - 8
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P. Digard et al. / International Congress Series 1219 (2001) 503–512
sense transcripts: capped and polyadenylated mRNAs, and exact complements (cRNA) which serve as replicative intermediate RNAs for the production of further vRNAs [1]. Four viral proteins are necessary and sufficient to carry out this process [2]: the three subunits of an RNA-dependent RNA polymerase, (PB1, PB2 and PA) and a 55-kDa single strand RNA-binding nucleoprotein (NP). Indeed, the v- and cRNA segments are always associated with these polypeptides to form ribonucleoprotein (RNP) structures [1]. Unusually for an RNA virus, influenza transcription occurs in the nuclei of infected cells [3]. Thus, on initiation of infection, incoming RNPs enter the nucleus, and during infection, newly synthesised RNP proteins also undergo nuclear import. However, progeny virions are assembled in the cytoplasm at the apical plasma membrane and at later times, this is reflected in the cytoplasmic accumulation of NP, generally assumed to be in the form of RNPs [1]. This changing pattern of RNP localisation 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, and in the virion is thought to act as the link between the lipid envelope and the packaged RNPs [4]. Moreover, M1 partially localises to the nucleus, where it promotes the export of RNPs [5,6]. NS2, which binds to RNPs via the M1 protein [7], possesses a nuclear export signal, and it has been proposed that NS2 is the viral factor directly responsible for the export of RNPs from the nucleus [8]. In addition, NP possesses the ability to locate to both cytoplasm and nucleus and in the absence of other viral proteins, it shuttles between the two compartments [6], suggesting that it interacts with the cellular nuclear export apparatus in the absence of M1 and NS2. Indeed, we have recently found evidence that NP interacts with the cellular CRM1-mediated nuclear export pathway and that this pathway is important for RNP export during virus infection [9]. However, the question remains: which mechanisms prevent the nuclear re-import of exported RNPs? One well documented method of modulating nuclear import is through interactions with cytoplasmic anchoring proteins [10], and recent work involving cell fractionation and fluorescent staining experiments has suggested that cytoplasmic NP is associated with the cytoskeleton [11,12]. The purpose of the experiments described here was to examine the possibility that NP might directly interact with the actin cytoskeleton.
2. Materials and methods 2.1. Plasmids and protein expression A plasmid containing the native A/PR/8/34 NP gene under the control of a bacteriophage T7 RNA polymerase promoter (pKT5) has previously been described [13]. A plasmid containing the NP gene fused to Escherichia coli maltose-binding protein (pMAL –NP) has also been reported [14]. Maltose-binding protein (MBP) or MBP fused to NP (MBP –NP) were purified from extracts of E. coli cultures containing plasmids pMAL-c2 (New England Biolabs) or pMAL –NP by affinity chromatography on amylose resin columns (New England Biolabs) as previously described [14]. The maltose-binding protein (MBP) moiety of MBP– NP was removed by the addition of 2
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mM CaCl2 and 0.5% (w/w) of factor Xa protease (New England Biolabs) in an overnight incubation at 14 C. NP was then purified by ion-exchange chromatography on a MonoQ column (Pharmacia) or a heparin III agarose column (Affinity Chromatography, Isle of Man). Samples were clarified by centrifugation at 390,000 gav for 15 min before use. For in vitro RNA binding assays, a 178-nucleotide synthetic RNA target was generated by in vitro transcription of plasmid pKT8-D 3050 as previously described [14]. 2.2. Actin-binding assays Purified rabbit muscle actin [14] was polymerised 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 assays contained 3 M 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 l. After mixing, the samples were centrifuged at 200,000 gav for 20 min at 20 C before separation into pellet and supernatant fractions. Equivalent amounts of each fraction were analysed by SDS-PAGE and stained with Coomassie brilliant blue dye. For electron microscopy, actin (1 M) and MBP or MBP– NP were mixed in F-buffer and left on ice for 20 min. Drops (30 l) were applied to carbon-coated grids for 1 min, rinsed with five drops of F-buffer and five drops of 1% uranyl acetate before blotting dry and viewing in a Philips 208S electron microscope at 20K or 30K magnification. Light scattering experiments were performed in an LS50B spectrophotometer (Perkin Elmer) using excitation and emission wavelengths of 520 nm (2.5-nm slit widths), with 4 M actin either in the form of F-actin (in F-buffer), or starting as G-actin (in 5 mM Tris –Cl pH 8.0, 0.1 mM ATP, 0.2 mM CaCl2 [G buffer]) with polymerisation induced by the addition of 100 mM NaCl, 1 mM MgCl2, 1 mM ATP. Reactions were performed at 20 C. For low-shear viscosity assays, 5 M G-actin was induced to polymerise in capillary tubes by the addition of 100 mM NaCl, 1 mM MgCl2, 1 mM ATP in the presence and absence of varying molar ratios of NP polypeptides. After overnight incubation at 4 C, the speed at which a small steel ball fell through the solution was measured [15]. 2.3. Transfection of tissue culture cells and indirect immunofluorescence HeLa cells were infected with a recombinant vaccinia virus encoding T7 RNA polymerase (VTF7; Ref. [16]) at a multiplicity of infection of 5 for 2 h at 37 C. The cells were washed three times with serum-free medium before transfection with plasmid DNA encoding NP using a cationic liposome mixture (Lipofectin; Gibco), as previously described [14]. After 4-h incubation at 37 C, the cells were washed with PBS containing 1% newborn calf serum, fixed in PBS containing 4% formaldehyde, and stained for NP using anti-RNP serum as previously described [14] or for -actin using monoclonal antibody AC-74 (Sigma). Fluorescence was viewed and images captured on an MRC 1024 confocal microscope. Control experiments, where cells were stained with individual fluorescent reagents (as well as untransfected cells), confirmed that the labelling was channel-specific (data not shown).
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3. Results 3.1. Binding of NP to F-actin We tested whether purified NP bound filamentous (F)-actin in vitro using a cosedimentation assay, where F-actin is readily separated from monomeric protein by centrifugation. As shown in Fig. 1a, most of the actin sedimented after centrifugation (lane 2), and only the expected critical monomer concentration of actin (0.1 – 0.2 M) remained in the supernatant (lane 1). The bacterially expressed NP purified as a major band of the expected electrophoretic mobility and three shorter NP polypeptides that presumably resulted from proteolytic cleavage of the authentic molecule (lane 5). In the absence of actin, only trace amounts of NP sedimented (lanes 5 and 6). However, when NP was sedimented after mixing with F-actin, the greater proportion of the NP now partitioned into the pellet fraction (lanes 3 and 4), demonstrating its association with actin filaments. Furthermore, NP fused to MBP or to glutathione-S-transferase (GST) also cosedimented with actin filaments, while MBP and GST alone did not [14]. The binding of NP to actin filaments was also visualised directly by electron microscopy. Actin filaments incubated with two molar equivalents of MBP formed an essentially random distribution of fibres (Fig. 1b). However, the addition of a similar ratio of MBP –NP to actin induced massive bundling of the filaments (Fig. 1c). Thus, we conclude that NP binds F-actin in vitro. Previous work has shown that a proportion of NP in influenza virus infected cells colocalises with peripheral F-actin [11,12]. We therefore tested whether similar colocalisation of NP and actin could be observed in the absence of other influenza virus polypeptides. HeLa cells were infected with vaccinia virus vTF7-3 expressing T7 RNA polymerase [16], transfected with plasmid pKT5 containing the NP gene under the control of a T7 RNA polymerase promoter, and 4 h later, examined for NP and -actin distribution by indirect immunofluorescence. A relatively high dose of PKT5 was transfected ( 0.3
Fig. 1. F-actin-binding activity of NP. (a) Cosedimentation of NP and actin. NP (1.5 M), actin (Act; 3 M) and an NP – actin mixture as labelled were centrifuged and separated into supernatant (S) and pellet (P) fractions before analysis by SDS-PAGE and staining with Coomassie brilliant blue. Arrows indicate the named polypeptides and molecular mass markers (M; kDa) are shown on the right. (b, c) Electron microscopy of actin filaments mixed with three-fold molar excesses of MBP (b) and MBP – NP (c). Scale bar: 100 nm.
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g4 104 cells) to bias NP towards cytoplasmic accumulation [14]. As expected, b-actin was largely peripheral and concentrated in areas of membrane ruffling (Fig. 2b, arrows). NP was diffusely distributed throughout the cytoplasm but also showed areas of concentration at the periphery of the cell in membrane ruffles, which in many cases colocalised with -actin (Fig. 2a, arrows). In addition, colocalisation of transfected NP was also observed with actin stress fibres [14]. This is consistent with the ability of NP to bind F-actin in vivo as well as in vitro. Electron microscopic examination of NP – actin complexes suggested that NP is an actinbundling polypeptide (Fig. 1c). To examine this further, we followed the physical size of actin fibres in solution by light scattering. The ability of F-actin to scatter light is dependent on filament length and is further enhanced by filament bundling [15]. In the absence of actin, 0.5 M MBP– NP showed a low level ( 100 U) of light scattering that was relatively consistent over 20 min (Fig. 3; NP). A similarly stable signal of around 170 U was obtained from a 4 M suspension of F-actin that had previously been allowed to polymerise to equilibrium (data not shown, but see G ! F trace in Fig. 3). However, when 0.5 m MBP – NP was added to this F-actin suspension, a very rapid increase in light scattering to around 500 U was observed (Fig. 3; F + NP), suggesting bundling of the actin filaments. Similar results were obtained when G-actin was allowed to polymerise into F-actin in the presence of NP. In the absence of NP, a 4-M solution of G-actin showed low-level light scattering that after the addition of salt, ATP and MgCl2 to induce polymerisation, increased over time with the characteristic sigmoidal curve (Fig. 3; G ! F), indicative of the lag phase imposed by the initially rate-limiting step of the formation of nucleation centres [15]. However, when G-actin was allowed to polymerise in the presence of 0.5 M MBP –NP, no lag phase was evident and the steady-state scattering intensity reached after 20 min was much higher (Fig. 3; G ! F + NP). This indicates that NP causes the polymerisation of longer and/or bundled actin filaments.
Fig. 2. Colocalisation of NP and -actin. HeLa cells were transfected with 0.3 g of pKT5 and analysed 4 h later by confocal microscopy after fixation and indirect immunofluorescent staining for (a) NP and (b) -actin.
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Fig. 3. Light scattering analysis of actin and NP. The scattering intensity at 520 nm of solutions of MBP – NP and actin (4 M) in the indicated mixtures was followed over 1200 s. In certain cases, actin was added as F-actin (F), in others, as G-actin (G) and induced to polymerise by the addition of NaCl, ATP and MgCl2 to the buffer.
The finding that NP induced the formation of actin bundles suggested that it might change the mechanical properties of microfilaments. To test this hypothesis, we investigated the effects of NP addition on the apparent viscosity of actin filaments. The viscosity of an F-actin solution is proportional to the length and number of filaments and these properties can be altered by the addition of actin-binding proteins [15]. The approach used was to allow the polymerisation of G-actin in capillary tubes in the presence and absence of MBP– NP and then measure the apparent viscosity of the solution by falling ball viscometry [15]. In the absence of NP, relatively low viscosity solutions of F-actin were formed, as assessed by the speed at which a ball bearing fell through the capillary (Fig. 4a). However, the apparent viscosity of the solutions increased as increasing amounts of MBP – NP were included in the polymerisation reactions. Low molar ratios of NP – actin ( < 0.4:1) caused only modest increases in viscosity, but higher ratios caused a dramatic increase (Fig. 4a). As a control, we tested the effects of adding a truncated version of MBP – NP which lacks the C-terminal two-thirds of NP (MBP – NPDC161) and is unable to bind actin (data not shown). No significant increase (or decrease) in apparent viscosity of the actin solution was seen, even with equimolar ratios of this mutant NP polypeptide
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Fig. 4. Low-shear viscosity analysis of actin and NP. (a) Titration of WT MBP – NP and MBP – NPDC161. Solutions (5 M) of G-actin were allowed to polymerise overnight at 4 C in the presence of increasing amounts of MBP – NP polypeptides before analysis by falling-ball viscometry. (b) The indicated mixes of actin (5 M), MBP – NP (3.5 M) and a 178-nucleotide transcript (0.4 M) were analysed as above.
(Fig. 4a). Thus, NP can alter the mechanical properties of an actin gel, again consistent with its ability to bundle actin fibres. Much of the NP in infected cells exists in the form of a complex with viral RNA. Previously, we have shown that the RNA- and actin-binding activities of NP are separate and that ternary complexes of NP, RNA and actin can form [14]. We therefore tested whether NP – RNA complexes also increased the viscosity of F-actin solutions. Polymer-
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isation reactions were set up containing various combinations of actin, MBP –NP and a 178-nucleotide RNA transcript corresponding to influenza virus segment 8 with the coding sequences deleted. Solutions of NP or NP and RNA had no measurable apparent viscosity in the absence of actin (Fig. 4b). Actin alone showed relatively low viscosity and this was not altered by the addition of RNA (Fig. 4b). As before, a mixture of NP and actin showed a dramatic increase in apparent viscosity, but this was abolished by the further addition of RNA (Fig. 4b). This suggests that NP – RNA complexes interact with F-actin in a different manner to free NP.
4. Discussion Two recent studies suggested that influenza virus NP interacts with microfilaments of the host cell cytoskeleton late in virus infection [11,12]. Both reports found that NP partitioned into cytoskeleton-containing pools during biochemical fractionation of infected cells, that NP colocalised 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 showing that NP binds directly to F-actin in vitro and is capable of inducing filament bundling (Figs. 1, 3 and 4). We also provide evidence that NP associates with actin filaments in vivo in the absence of other influenza proteins (Fig. 2). Furthermore, in an extension of the study reported here, we have shown that complexes containing F-actin, NP and RNA could be formed and have identified point mutations in NP, which specifically weaken the NP – actin interaction [14]. The question therefore arises as to what role(s) the NP – actin interaction might play in the virus lifecycle. The finding that NP induces actin bundling in vitro (Figs. 1, 3 and 4) raises the possibility that the virus might specifically modify the actin cytoskeleton during infection. However, influenza virus infection does not cause major alterations in the microfilament network until very late in infection when the cells begin to apoptose (data not shown). More subtle modifications have not yet been ruled out, and this is a worthwhile area for further experimentation. We note though, that most of the NP in infected cells is thought to be bound to RNA [1] and that NP did not cause the dramatic changes in actin viscosity when in the presence of RNA (Fig. 4). From analogy with the ways other viral and intracellular pathogens subvert the actin cytoskeleton [17] roles in intracellular trafficking of viral components, viral transcription and assembly and release of virions are possible. The hypothesis that influenza uses microfilaments for the purpose of directing the intracellular localisation of RNPs is perhaps supported by the behaviour of NP mutants with weakened affinity for actin: we have shown that in the absence of other influenza virus polypeptides, these mutant NP molecules displayed an increased propensity to accumulate in the nucleus. This suggested the possibility that actin-binding might serve to retain RNPs in the cytoplasm [14]. The RNP particles of some paramyxoviruses are associated with the actin cytoskeleton, and in the case of human parainfluenza virus type 3, actin stimulates virus transcription [18]. However, the ability of NP mutants to support transcription and replication of a model influenza virus segment did not correlate with their affinity for F-actin [14], arguing
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against actin playing a significant role in influenza virus transcription. The question of whether actin plays a role during influenza virus budding is an interesting one. Although disruption of microfilaments with cytochalasins does not inhibit the assembly and release of spherical influenza virions, an intact actin cytoskeleton is necessary for the formation of filamentous virus particles [19]. Recent work from our laboratory suggests that actin microfilaments are involved in maintaining the correct organisation of the plasma membrane lipid rafts that the virus is thought to bud from (unpublished experiments). However, it remains to be determined whether microfilaments play other roles during the assembly of filamentous particles. The ultimate determination of the role(s) of the actin cytoskeleton in influenza virus infection requires further experimentation. An interesting avenue will be to try and create mutant viruses containing NP mutants deficient for actin-binding, using the recently described systems for influenza virus reverse genetics [20,21].
Acknowledgements We are grateful to Drs. B. Pope and A. Weeds for the gift of actin and their expertise. This work was supported by grants from the Royal Society, Wellcome Trust (nos. 048911 and 059151) and Medical Research Council (no. G9901213) to PD. PD is a Royal Society University Research fellow.
References [1] R.A. Lamb, R.M. Krug, Orthomyxoviridae: the viruses and their replication, in: B.N. Fields, D.M. Knipe, P.M. Howley (Eds.), Fields Virology, Lippincott-Raven, Philadelphia, 1996, pp. 1353 – 1396. [2] T.-S. Huang, P. Palese, M. Krystal, Determination of influenza virus proteins required for genome replication, J. Virol. 64 (1990) 5669 – 5673. [3] C. Herz, E. Stavnezer, R.M. Krug, Influenza virus, an RNA virus, synthesizes its messenger RNA in the nucleus of infected cells, Cell 26 (1981) 391 – 400. [4] G. Whittaker, M. Bui, A. Helenius, The role of M1 in nuclear import and export in influenza virus infection, Trends Cell Biol. 6 (1996) 67 – 71. [5] K. Martin, A. Helenius, Nuclear transport of influenza virus ribonucleoproteins: the viral matrix protein (M1) promotes export and inhibits import, Cell 67 (1991) 117 – 130. [6] G. Whittaker, M. Bui, A. Helenius, Nuclear trafficking of influenza virus ribonucleoproteins in heterokaryons, J. Virol. 70 (1996) 2743 – 2756. [7] J. Yasuda, S. Nakada, A. Kato, T. Toyoda, A. Ishihama, Molecular assembly of influenza virus: association of the NS2 protein with virion matrix, Virology 196 (1993) 249 – 255. [8] R.E. O’Neill, J. Talon, P. Palese, The influenza virus NEP (NS2 protein) mediates the nuclear export of viral ribonucleoproteins, EMBO J. 17 (1998) 288 – 296. [9] D. Elton, M. Simpson-Holley, K. Archer, L. Medcalf, R. Hallam, J. McCauley, P. Digard, Interaction of the influenza virus nucleoprotein with the cellular CRM1-mediated nuclear export pathway. J. Virol. 75 (2001) 408 – 419. [10] E.A. Nigg, Nucleocytoplasmic transport: signals, mechanisms and regulation, Nature 386 (1997) 779 – 787. [11] R.T. Avalos, Z. Yu, D.P. Nayak, Association of influenza virus NP and M1 proteins with cellular cytoskeletal elements in influenza virus infected cells, J. Virol. 71 (1997) 2947 – 2958. [12] M. Husain, C.M. Gupta, Interactions of viral matrix protein and nucleoprotein with the host cell cytoskeletal actin in influenza viral infection, Curr. Sci. 73 (1997) 40 – 47.
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[13] V. Blok, C. Cianci, K.W. Tibbles, S.C. Inglis, M. Krystal, P. Digard, Inhibition of the influenza virus RNA-dependent RNA polymerase by antisera directed against the carboxy-terminal region of the PB2 subunit, J. Gen. Virol. 77 (1996) 1025 – 1033. [14] P. Digard, D. Elton, K. Bishop, E. Medcalf, A. Weeds, B. Pope, Modulation of nuclear localization of the influenza virus nucleoprotein through interaction with actin filaments, J. Virol. 73 (1999) 2222 – 2231. [15] J.A. Cooper, Actin filament assembly and organization in vitro, in: K.L. Carraway, C.A.C. Carraway (Eds.), The Cytoskeleton: A Practical Approach, IRL Press, Oxford, 1992, pp. 47 – 71. [16] T.R. Fuerst, P.L. Earl, B. Moss, Use of a hybrid vaccinia virus-T7 RNA polymerase system for expression of target genes, Mol. Cell Biol. 7 (1987) 2538 – 2544. [17] S. Cudmore, I. Reckman, M. Way, Viral manipulations of the actin cytoskeleton, Trends Biochem. Sci. 5 (1997) 142 – 148. [18] B.P. De, A. Lesoon, A.K. Banerjee, Human parainfluenza virus type 3 transcription in vitro: role of cellular actin in mRNA synthesis, J. Virol. 65 (1991) 3268 – 3275. [19] P.C. Roberts, W.R. Compans, Host cell dependence of viral morphology, Proc. Natl. Acad. Sci. U. S. A. 95 (1998) 5746 – 5751. [20] G. Neumann, T. Watanabe, H. Ito, H. Watanabe, S. Goto, P. Gao, M. Hughes, D.R. Perez, R. Donis, E. Hoffmann, G. Hobom, Y. Kawaoka, Generation of influenza A viruses entirely from cloned cDNAs, Proc. Natl. Acad. Sci. U. S. A. 96 (1999) 9345 – 9350. [21] E. Fodor, L. Devenish, O.G. Engelhardt, P. Palese, G.G. Brownlee, A. Garcı´a-Sastre, Rescue of influenza A virus from recombinant DNA, J. Virol. 73 (1999) 9679 – 9682.
Interaction of the inf luenza virus nucleoprotein with F-actin Paul Digard* , Debra Elton, Martha Simpson-Holley, Elizabeth Medcalf Division of Virology, Department of Pathology, University of Cambridge, Tennis Court Road, Cambridge CB2 1QP, UK
Abstract Introduction: The influenza virus genome is transcribed in the nucleus of infected cells but assembled into progeny virions in the cytoplasm. This is reflected in the cellular distribution of the virus nucleoprotein (NP), a protein which encapsidates genomic RNA to form ribonucleoprotein (RNP) structures: at the early stage post-infection, NP is found in the nucleus, but later it is found predominantly in the cytoplasm. The purpose of this study was to examine the possibility that cytoplasmic NP interacts with actin microfilaments. Methods: Bacterially expressed NP was tested for the ability to interact with filamentous (F)-actin in a variety of in vitro assays, and the localisation of actin and exogenously expressed NP in mammalian cells was examined microscopically. Results: Purified NP bound actin filaments in vitro and showed partial colocalisation with b-actin in vivo. Electron microscopy showed that NP induced bundling of actin fibres in vitro. In confirmation of this, NP caused a dramatic increase in the low-shear viscosity and light-scattering properties of Factin suspensions. Conclusions: NP binds F-actin in vitro and can alter the mechanical properties of actin filaments. This raises the possibility that influenza virus may use the host-cell cytoskeleton during virus replication. D 2001 Elsevier Science B.V. All rights reserved. Keywords: Microfilaments; Ribonucleoprotein; Bundling
1. Introduction The influenza A virus genome consists of eight segments of negative sense singlestranded RNA (vRNA), which are transcribed in infected cells to yield two types of positive
*
Corresponding author. Tel.: +44-1223-336918; fax: +44-1223-336926. E-mail address: [email protected] (P. Digard).
0531-5131/01/$ – see front matter D 2001 Elsevier Science B.V. All rights reserved. PII: S 0 5 3 1 - 5 1 3 1 ( 0 1 ) 0 0 6 2 8 - 8
504
P. Digard et al. / International Congress Series 1219 (2001) 503–512
sense transcripts: capped and polyadenylated mRNAs, and exact complements (cRNA) which serve as replicative intermediate RNAs for the production of further vRNAs [1]. Four viral proteins are necessary and sufficient to carry out this process [2]: the three subunits of an RNA-dependent RNA polymerase, (PB1, PB2 and PA) and a 55-kDa single strand RNA-binding nucleoprotein (NP). Indeed, the v- and cRNA segments are always associated with these polypeptides to form ribonucleoprotein (RNP) structures [1]. Unusually for an RNA virus, influenza transcription occurs in the nuclei of infected cells [3]. Thus, on initiation of infection, incoming RNPs enter the nucleus, and during infection, newly synthesised RNP proteins also undergo nuclear import. However, progeny virions are assembled in the cytoplasm at the apical plasma membrane and at later times, this is reflected in the cytoplasmic accumulation of NP, generally assumed to be in the form of RNPs [1]. This changing pattern of RNP localisation 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, and in the virion is thought to act as the link between the lipid envelope and the packaged RNPs [4]. Moreover, M1 partially localises to the nucleus, where it promotes the export of RNPs [5,6]. NS2, which binds to RNPs via the M1 protein [7], possesses a nuclear export signal, and it has been proposed that NS2 is the viral factor directly responsible for the export of RNPs from the nucleus [8]. In addition, NP possesses the ability to locate to both cytoplasm and nucleus and in the absence of other viral proteins, it shuttles between the two compartments [6], suggesting that it interacts with the cellular nuclear export apparatus in the absence of M1 and NS2. Indeed, we have recently found evidence that NP interacts with the cellular CRM1-mediated nuclear export pathway and that this pathway is important for RNP export during virus infection [9]. However, the question remains: which mechanisms prevent the nuclear re-import of exported RNPs? One well documented method of modulating nuclear import is through interactions with cytoplasmic anchoring proteins [10], and recent work involving cell fractionation and fluorescent staining experiments has suggested that cytoplasmic NP is associated with the cytoskeleton [11,12]. The purpose of the experiments described here was to examine the possibility that NP might directly interact with the actin cytoskeleton.
2. Materials and methods 2.1. Plasmids and protein expression A plasmid containing the native A/PR/8/34 NP gene under the control of a bacteriophage T7 RNA polymerase promoter (pKT5) has previously been described [13]. A plasmid containing the NP gene fused to Escherichia coli maltose-binding protein (pMAL –NP) has also been reported [14]. Maltose-binding protein (MBP) or MBP fused to NP (MBP –NP) were purified from extracts of E. coli cultures containing plasmids pMAL-c2 (New England Biolabs) or pMAL –NP by affinity chromatography on amylose resin columns (New England Biolabs) as previously described [14]. The maltose-binding protein (MBP) moiety of MBP– NP was removed by the addition of 2
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mM CaCl2 and 0.5% (w/w) of factor Xa protease (New England Biolabs) in an overnight incubation at 14 C. NP was then purified by ion-exchange chromatography on a MonoQ column (Pharmacia) or a heparin III agarose column (Affinity Chromatography, Isle of Man). Samples were clarified by centrifugation at 390,000 gav for 15 min before use. For in vitro RNA binding assays, a 178-nucleotide synthetic RNA target was generated by in vitro transcription of plasmid pKT8-D 3050 as previously described [14]. 2.2. Actin-binding assays Purified rabbit muscle actin [14] was polymerised 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 assays contained 3 M 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 l. After mixing, the samples were centrifuged at 200,000 gav for 20 min at 20 C before separation into pellet and supernatant fractions. Equivalent amounts of each fraction were analysed by SDS-PAGE and stained with Coomassie brilliant blue dye. For electron microscopy, actin (1 M) and MBP or MBP– NP were mixed in F-buffer and left on ice for 20 min. Drops (30 l) were applied to carbon-coated grids for 1 min, rinsed with five drops of F-buffer and five drops of 1% uranyl acetate before blotting dry and viewing in a Philips 208S electron microscope at 20K or 30K magnification. Light scattering experiments were performed in an LS50B spectrophotometer (Perkin Elmer) using excitation and emission wavelengths of 520 nm (2.5-nm slit widths), with 4 M actin either in the form of F-actin (in F-buffer), or starting as G-actin (in 5 mM Tris –Cl pH 8.0, 0.1 mM ATP, 0.2 mM CaCl2 [G buffer]) with polymerisation induced by the addition of 100 mM NaCl, 1 mM MgCl2, 1 mM ATP. Reactions were performed at 20 C. For low-shear viscosity assays, 5 M G-actin was induced to polymerise in capillary tubes by the addition of 100 mM NaCl, 1 mM MgCl2, 1 mM ATP in the presence and absence of varying molar ratios of NP polypeptides. After overnight incubation at 4 C, the speed at which a small steel ball fell through the solution was measured [15]. 2.3. Transfection of tissue culture cells and indirect immunofluorescence HeLa cells were infected with a recombinant vaccinia virus encoding T7 RNA polymerase (VTF7; Ref. [16]) at a multiplicity of infection of 5 for 2 h at 37 C. The cells were washed three times with serum-free medium before transfection with plasmid DNA encoding NP using a cationic liposome mixture (Lipofectin; Gibco), as previously described [14]. After 4-h incubation at 37 C, the cells were washed with PBS containing 1% newborn calf serum, fixed in PBS containing 4% formaldehyde, and stained for NP using anti-RNP serum as previously described [14] or for -actin using monoclonal antibody AC-74 (Sigma). Fluorescence was viewed and images captured on an MRC 1024 confocal microscope. Control experiments, where cells were stained with individual fluorescent reagents (as well as untransfected cells), confirmed that the labelling was channel-specific (data not shown).
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3. Results 3.1. Binding of NP to F-actin We tested whether purified NP bound filamentous (F)-actin in vitro using a cosedimentation assay, where F-actin is readily separated from monomeric protein by centrifugation. As shown in Fig. 1a, most of the actin sedimented after centrifugation (lane 2), and only the expected critical monomer concentration of actin (0.1 – 0.2 M) remained in the supernatant (lane 1). The bacterially expressed NP purified as a major band of the expected electrophoretic mobility and three shorter NP polypeptides that presumably resulted from proteolytic cleavage of the authentic molecule (lane 5). In the absence of actin, only trace amounts of NP sedimented (lanes 5 and 6). However, when NP was sedimented after mixing with F-actin, the greater proportion of the NP now partitioned into the pellet fraction (lanes 3 and 4), demonstrating its association with actin filaments. Furthermore, NP fused to MBP or to glutathione-S-transferase (GST) also cosedimented with actin filaments, while MBP and GST alone did not [14]. The binding of NP to actin filaments was also visualised directly by electron microscopy. Actin filaments incubated with two molar equivalents of MBP formed an essentially random distribution of fibres (Fig. 1b). However, the addition of a similar ratio of MBP –NP to actin induced massive bundling of the filaments (Fig. 1c). Thus, we conclude that NP binds F-actin in vitro. Previous work has shown that a proportion of NP in influenza virus infected cells colocalises with peripheral F-actin [11,12]. We therefore tested whether similar colocalisation of NP and actin could be observed in the absence of other influenza virus polypeptides. HeLa cells were infected with vaccinia virus vTF7-3 expressing T7 RNA polymerase [16], transfected with plasmid pKT5 containing the NP gene under the control of a T7 RNA polymerase promoter, and 4 h later, examined for NP and -actin distribution by indirect immunofluorescence. A relatively high dose of PKT5 was transfected ( 0.3
Fig. 1. F-actin-binding activity of NP. (a) Cosedimentation of NP and actin. NP (1.5 M), actin (Act; 3 M) and an NP – actin mixture as labelled were centrifuged and separated into supernatant (S) and pellet (P) fractions before analysis by SDS-PAGE and staining with Coomassie brilliant blue. Arrows indicate the named polypeptides and molecular mass markers (M; kDa) are shown on the right. (b, c) Electron microscopy of actin filaments mixed with three-fold molar excesses of MBP (b) and MBP – NP (c). Scale bar: 100 nm.
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g4 104 cells) to bias NP towards cytoplasmic accumulation [14]. As expected, b-actin was largely peripheral and concentrated in areas of membrane ruffling (Fig. 2b, arrows). NP was diffusely distributed throughout the cytoplasm but also showed areas of concentration at the periphery of the cell in membrane ruffles, which in many cases colocalised with -actin (Fig. 2a, arrows). In addition, colocalisation of transfected NP was also observed with actin stress fibres [14]. This is consistent with the ability of NP to bind F-actin in vivo as well as in vitro. Electron microscopic examination of NP – actin complexes suggested that NP is an actinbundling polypeptide (Fig. 1c). To examine this further, we followed the physical size of actin fibres in solution by light scattering. The ability of F-actin to scatter light is dependent on filament length and is further enhanced by filament bundling [15]. In the absence of actin, 0.5 M MBP– NP showed a low level ( 100 U) of light scattering that was relatively consistent over 20 min (Fig. 3; NP). A similarly stable signal of around 170 U was obtained from a 4 M suspension of F-actin that had previously been allowed to polymerise to equilibrium (data not shown, but see G ! F trace in Fig. 3). However, when 0.5 m MBP – NP was added to this F-actin suspension, a very rapid increase in light scattering to around 500 U was observed (Fig. 3; F + NP), suggesting bundling of the actin filaments. Similar results were obtained when G-actin was allowed to polymerise into F-actin in the presence of NP. In the absence of NP, a 4-M solution of G-actin showed low-level light scattering that after the addition of salt, ATP and MgCl2 to induce polymerisation, increased over time with the characteristic sigmoidal curve (Fig. 3; G ! F), indicative of the lag phase imposed by the initially rate-limiting step of the formation of nucleation centres [15]. However, when G-actin was allowed to polymerise in the presence of 0.5 M MBP –NP, no lag phase was evident and the steady-state scattering intensity reached after 20 min was much higher (Fig. 3; G ! F + NP). This indicates that NP causes the polymerisation of longer and/or bundled actin filaments.
Fig. 2. Colocalisation of NP and -actin. HeLa cells were transfected with 0.3 g of pKT5 and analysed 4 h later by confocal microscopy after fixation and indirect immunofluorescent staining for (a) NP and (b) -actin.
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Fig. 3. Light scattering analysis of actin and NP. The scattering intensity at 520 nm of solutions of MBP – NP and actin (4 M) in the indicated mixtures was followed over 1200 s. In certain cases, actin was added as F-actin (F), in others, as G-actin (G) and induced to polymerise by the addition of NaCl, ATP and MgCl2 to the buffer.
The finding that NP induced the formation of actin bundles suggested that it might change the mechanical properties of microfilaments. To test this hypothesis, we investigated the effects of NP addition on the apparent viscosity of actin filaments. The viscosity of an F-actin solution is proportional to the length and number of filaments and these properties can be altered by the addition of actin-binding proteins [15]. The approach used was to allow the polymerisation of G-actin in capillary tubes in the presence and absence of MBP– NP and then measure the apparent viscosity of the solution by falling ball viscometry [15]. In the absence of NP, relatively low viscosity solutions of F-actin were formed, as assessed by the speed at which a ball bearing fell through the capillary (Fig. 4a). However, the apparent viscosity of the solutions increased as increasing amounts of MBP – NP were included in the polymerisation reactions. Low molar ratios of NP – actin ( < 0.4:1) caused only modest increases in viscosity, but higher ratios caused a dramatic increase (Fig. 4a). As a control, we tested the effects of adding a truncated version of MBP – NP which lacks the C-terminal two-thirds of NP (MBP – NPDC161) and is unable to bind actin (data not shown). No significant increase (or decrease) in apparent viscosity of the actin solution was seen, even with equimolar ratios of this mutant NP polypeptide
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Fig. 4. Low-shear viscosity analysis of actin and NP. (a) Titration of WT MBP – NP and MBP – NPDC161. Solutions (5 M) of G-actin were allowed to polymerise overnight at 4 C in the presence of increasing amounts of MBP – NP polypeptides before analysis by falling-ball viscometry. (b) The indicated mixes of actin (5 M), MBP – NP (3.5 M) and a 178-nucleotide transcript (0.4 M) were analysed as above.
(Fig. 4a). Thus, NP can alter the mechanical properties of an actin gel, again consistent with its ability to bundle actin fibres. Much of the NP in infected cells exists in the form of a complex with viral RNA. Previously, we have shown that the RNA- and actin-binding activities of NP are separate and that ternary complexes of NP, RNA and actin can form [14]. We therefore tested whether NP – RNA complexes also increased the viscosity of F-actin solutions. Polymer-
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isation reactions were set up containing various combinations of actin, MBP –NP and a 178-nucleotide RNA transcript corresponding to influenza virus segment 8 with the coding sequences deleted. Solutions of NP or NP and RNA had no measurable apparent viscosity in the absence of actin (Fig. 4b). Actin alone showed relatively low viscosity and this was not altered by the addition of RNA (Fig. 4b). As before, a mixture of NP and actin showed a dramatic increase in apparent viscosity, but this was abolished by the further addition of RNA (Fig. 4b). This suggests that NP – RNA complexes interact with F-actin in a different manner to free NP.
4. Discussion Two recent studies suggested that influenza virus NP interacts with microfilaments of the host cell cytoskeleton late in virus infection [11,12]. Both reports found that NP partitioned into cytoskeleton-containing pools during biochemical fractionation of infected cells, that NP colocalised 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 showing that NP binds directly to F-actin in vitro and is capable of inducing filament bundling (Figs. 1, 3 and 4). We also provide evidence that NP associates with actin filaments in vivo in the absence of other influenza proteins (Fig. 2). Furthermore, in an extension of the study reported here, we have shown that complexes containing F-actin, NP and RNA could be formed and have identified point mutations in NP, which specifically weaken the NP – actin interaction [14]. The question therefore arises as to what role(s) the NP – actin interaction might play in the virus lifecycle. The finding that NP induces actin bundling in vitro (Figs. 1, 3 and 4) raises the possibility that the virus might specifically modify the actin cytoskeleton during infection. However, influenza virus infection does not cause major alterations in the microfilament network until very late in infection when the cells begin to apoptose (data not shown). More subtle modifications have not yet been ruled out, and this is a worthwhile area for further experimentation. We note though, that most of the NP in infected cells is thought to be bound to RNA [1] and that NP did not cause the dramatic changes in actin viscosity when in the presence of RNA (Fig. 4). From analogy with the ways other viral and intracellular pathogens subvert the actin cytoskeleton [17] roles in intracellular trafficking of viral components, viral transcription and assembly and release of virions are possible. The hypothesis that influenza uses microfilaments for the purpose of directing the intracellular localisation of RNPs is perhaps supported by the behaviour of NP mutants with weakened affinity for actin: we have shown that in the absence of other influenza virus polypeptides, these mutant NP molecules displayed an increased propensity to accumulate in the nucleus. This suggested the possibility that actin-binding might serve to retain RNPs in the cytoplasm [14]. The RNP particles of some paramyxoviruses are associated with the actin cytoskeleton, and in the case of human parainfluenza virus type 3, actin stimulates virus transcription [18]. However, the ability of NP mutants to support transcription and replication of a model influenza virus segment did not correlate with their affinity for F-actin [14], arguing
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against actin playing a significant role in influenza virus transcription. The question of whether actin plays a role during influenza virus budding is an interesting one. Although disruption of microfilaments with cytochalasins does not inhibit the assembly and release of spherical influenza virions, an intact actin cytoskeleton is necessary for the formation of filamentous virus particles [19]. Recent work from our laboratory suggests that actin microfilaments are involved in maintaining the correct organisation of the plasma membrane lipid rafts that the virus is thought to bud from (unpublished experiments). However, it remains to be determined whether microfilaments play other roles during the assembly of filamentous particles. The ultimate determination of the role(s) of the actin cytoskeleton in influenza virus infection requires further experimentation. An interesting avenue will be to try and create mutant viruses containing NP mutants deficient for actin-binding, using the recently described systems for influenza virus reverse genetics [20,21].
Acknowledgements We are grateful to Drs. B. Pope and A. Weeds for the gift of actin and their expertise. This work was supported by grants from the Royal Society, Wellcome Trust (nos. 048911 and 059151) and Medical Research Council (no. G9901213) to PD. PD is a Royal Society University Research fellow.
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