- Email: [email protected]
Functional gap junction genes are encoded by insect viruses
Magazine R491
Functional gap junction genes are encoded by insect viruses Matthew W. Turnbull1*, Anne-Nathalie Volkoff2, Bruce A. Webb3 and Pauline Phelan4 Ichnoviruses belong to the virus family Polydnaviridae, whose members are obligately associated with certain endoparasitoid wasps. Expression of ichnovirus genes in parasitized lepidopteran hosts leads to immune suppression and is essential for successful parasitization. To date, the role of specific ichnovirus genes in alteration of host physiology has been unclear, and no cellular homologues have been described. Here, we describe the isolation of a gene family from two ichnoviruses that is homologous to the innexin gene family, which encodes gap junctions in invertebrates. Campoletis sonorensis ichnovirus (CsIV) innexins are expressed in multiple tissues in infected lepidopterans, including haemocytes, the primary immunocytes of the host. Two of the CsIV proteins have been expressed and shown to form functional gap junctions in Xenopus oocytes. To our knowledge this is the first study to describe gap junction genes in any virus. We hypothesize that the virus innexins disrupt cellular immunity in infected insects by altering normal gap junctional intercellular communication. This would represent a novel mechanism of viral alteration of host physiology, and suggests that gap junctions play a crucial role in coordinating cellular immune responses. Gap junctions, encoded by innexins in insects [1], coordinate multicellular processes. In insects, blood cells (haemocytes) play a critical role in immune responses and gap junctions are known to form between those cells [2,3], yet the functions of gap junctions in
insect immunity are unclear. Pathogenic agents, such as polydnaviruses, may provide a tool to investigate their role in nonmodel systems. The virus family Polydnaviridae is unique in that its members, the ichnoviruses and bracoviruses, are obligately associated with parasitoid wasps [4]. Virions are delivered to host caterpillars during parasitization where virus-encoded transcripts and proteins are produced in the absence of virus replication [4]. In the case of ichnoviruses, virusencoded proteins almost invariably suppress the immune system of the caterpillar, and thus are essential for successful parasitization (reviewed in [5]). Polydnavirus genomes are unusual in that they exhibit simultaneous reductive and proliferative evolution, i.e. few, if any, genes associated with virus replication and packaging are encapsidated, and the packaged genome contains genes generally associated with pathogenesis of the caterpillar host [5,6]. Therefore, genes identified in the ichnovirus genome may be assumed a priori both to be necessary for successful parasitization and to induce either immunological or developmental pathologies in the infected caterpillar. We identified five complete open reading frames in the genomes of two polydnaviruses, CsIV and Hyposoter didymator ichnovirus (HdIV), that shared significant (e-values <10−40) global similarity to innexins (see Table S1 in Supplemental data published with this paper online). These virus innexins, termed vinnexins, encode 22 of 39 invariant residues found in insect innexins (Figure S1), but form a well supported virus-specific clade in gene trees (Figure S2). Two CsIV vinnexins, cs-Vnxd and cs-Vnxg, were expressed in paired Xenopus laevis oocytes and assayed for their ability to form functional gap junctions. Cell pairs in which both cells expressed either cs-Vnxd or csVnxg electrically coupled (Figure 1). Average junctional conductance for these homotypic channels was 1.87 µS for cs-Vnxd (n = 15) and 3.62 µS for cs-Vnxg
Figure 1. Vinnexins make functional gap junctions in Xenopus oocytes. Xenopus laevis oocytes were microinjected with in vitro transcribed vinnexin RNAs or water (negative control). The vitelline membrane was removed to expose the plasma membrane and cells were paired. Cell pairs were recorded 24–48 h later using a double voltage clamp technique. Both oocytes of a pair were clamped to a holding potential (Vh) of –80mV. Sequential 10 mV depolarizing and hyperpolarizing voltage steps (V1) were applied to one cell (cell 1 in (A)) to generate transjunctional voltage, Vj (V1–Vh), and the current, Ij (–I2) required to maintain the neighbour cell (cell 2 in (B–D) at the holding potential was µS) is recorded. Junctional conductance (µ Ij/Vj. Transjunctional voltage steps elicit junctional currents in cell pairs injected with (B) cs-vnxd (0.5 ng) and (C) cs-vnxg (0.5 ng) but not in (D) water-injected control pairs.
(n = 32) following expression of 0.5–1.0 ng RNA, which is within the range previously observed for Drosophila innexins. Heterotypic channels also formed between cell pairs in which one cell expressed cs-Vnxd and the other cs-Vnxg (data not shown). Vinnexin channels were not voltage dependent; junctional current (Ij) was linear at all transjunctional voltage steps (Vj, Figure 1A–C). The vinnexins are transcribed in several tissues of the infected
Current Biology Vol 15 No 13 R492
comparisons as well as experimental procedures are available at http://www.current-biology.com/cgi/ content/full/15/13/R491/DC1/
Figure 2. cs-Vnxq2 protein localizes to haemocyte membranes. Isolated haemocytes from the CsIV-susceptible host caterpillar Heliothis virescens were labelled with anti-peptide antisera generated against a unique sequence in the cytoplasmic region of the cs-Vnxq2 protein. (A) Immunofluorescence confocal microscopy detected no specific signal in control haemocytes. (B) Antisera demonstrated localization (red) to cellular membranes in haemocytes isolated from 3 day post-parasitized H. virescens. Green, FITC-phalloidin labeled F-actin. Note the actin cytoskeleton is disrupted in infected haemocytes hence the absence of cell processes in (B). Scale bar = 10 µm.
caterpillar (Figure S3), concurrently with two newly isolated host lepidopteran innexin-2 orthologues (data not shown). Peptide antisera raised against a unique region of cytoplasmic sequence of the CsIV vinnexin cs-Vnxq2 were used to label isolated haemocytes. No signal was observed in control insects using immunofluorescence microscopy (Figure 2A); by contrast, positive signal was observed consistently in haemocytes of parasitized insects (Figure 2B). Western blots confirmed that the antigen was restricted to infected haemocytes with no detectable signal in other tissues (Figure S4). Our data demonstrate that some ichnoviruses have coopted a host gene family controlling intercellular communication, presumably as a novel means to alter host physiology. An intriguing hypothesis, supported by the observed haemocyte specificity and the known role of ichnoviruses in parasitoid–host interactions, is that vinnexins disrupt host immunity, interfering with haemocytic encapsulation, the primary immune response against haemocoelic parasites in insects. The native encapsulation reaction requires two haemocyte populations interacting over several hours to days in a tightly
regulated cascade of behaviours [2,7]. The mechanisms of regulation during intermediate to terminal stages of encapsulation are unknown but gap junctions are probably involved [8,9]. The vinnexins may alter native gapjunction-mediated intercellular communication among haemocytes by modifying the permeability of host innexin channels, via vinnexin–innexin interactions, or by permitting diffusion through vinnexin channels, thereby abrogating encapsulation. The discovery and initial characterization of the vinnexins in combination with the isolation of caterpillar innexins transcribed in haemocytes provides a set of conceptual and molecular tools for future studies of the functions of gap junctions in cellular immunity. Acknowledgements Thanks to C. Holcroft, R. Hilgarth, J. Kroemer, and E. Fleming for helpful comments and assistance. This work was supported by the University of Kentucky (M.W.T.), the Wellcome Trust and the Leverhulme Trust (P.P.), and the National Science Foundation and the United States Department of Agriculture (B.A.W.). This is publication number 0408-103 of the University of Kentucky Experiment Station. Supplemental data Supplemental data including analysis of vinnexin expression and sequence
References 1. Phelan, P., Stebbings, L.A., Baines, R.A., Bacon, J.P., Davies, J.A., and Ford, C. (1998). Drosophila Shaking-B protein forms gap junctions in paired Xenopus oocytes. Nature 391, 181–184. 2. Grimstone, A.V., Rotheram, S., and Salt, G. (1967). An electronmicroscope study of capsule formation by insect blood cells. J. Cell Sci. 2, 281–292. 3. Churchill, D., Coodin, S., Shivers, R.R., and Caveney, S. (1993). Rapid de novo formation of gap junctions between insect hemocytes in vitro: a freeze-fracture, dye-transfer and patch-clamp study. J. Cell Sci. 104, 763–772. 4. Webb, B.A., Beckage, N.E., Hayakawa, Y., Krell, P.J., Lanzrein, B., Stoltz, D.B., Strand, M.R., and Summers, M.D. (2000). Polydnaviridae. In Virus Taxonomy: The Classification and Nomenclature of Viruses. The 7th Report of the International Committee on Taxonomy of Viruses. M.H.V. van Regenmortel, et al., eds (San Diego: Academic Press), pp. 1167. 5. Kroemer, J.A., and Webb, B.A. (2004). Polydnavirus genes and genomes: Emerging gene families and new insights into polydnavirus replication. Annu. Rev. Entomol. 49, 431–456. 6. Espagne, E., Dupuy, C., Huguet, E., Cattolico, L., Provost, B., Martins, N., Poirie, M., Periquet, G., and Drezen, J.M. (2004). Genome sequence of a polydnavirus: insights into symbiotic virus evolution. Science 306, 286–289. 7. Pech, L.L., and Strand, M.R. (1996). Granular cells are required for encapsulation of foreign targets by insect haemocytes. J. Cell Sci. 109, 2053–2060. 8. Gupta, A.P. (1991). Gap cell junctions, cell adhesion molecules, and molecular basis of encapsulation. In Immmunology of Insects and Other Arthropods. A.P. Gupta, ed. (Boca Raton: CRC Press), pp. 133–167. 9. Lackie, A.M. (1988). Haemocyte behaviour. Adv. Insect Physiol. 21, 85–178. 1Clemson
University, 114 Long Hall, Box 340315, Department of Entomology, Soils, and Plant Sciences, Clemson, SC 29634-0315, USA. 2I.N.R.A.-U.M. II, Laboratoire de Pathologie Comparée, 30380 Saint Christol-lez-Alès, France. 3University of Kentucky, Department of Entomology, S-225 Agricultural Science Center, N, University of Kentucky, Lexington, KY 40546-0091, USA. 4University of Kent, Department of Biosciences, Canterbury, Kent, CT2 7NJ, UK. *E-mail: [email protected]
Functional gap junction genes are encoded by insect viruses Matthew W. Turnbull1*, Anne-Nathalie Volkoff2, Bruce A. Webb3 and Pauline Phelan4 Ichnoviruses belong to the virus family Polydnaviridae, whose members are obligately associated with certain endoparasitoid wasps. Expression of ichnovirus genes in parasitized lepidopteran hosts leads to immune suppression and is essential for successful parasitization. To date, the role of specific ichnovirus genes in alteration of host physiology has been unclear, and no cellular homologues have been described. Here, we describe the isolation of a gene family from two ichnoviruses that is homologous to the innexin gene family, which encodes gap junctions in invertebrates. Campoletis sonorensis ichnovirus (CsIV) innexins are expressed in multiple tissues in infected lepidopterans, including haemocytes, the primary immunocytes of the host. Two of the CsIV proteins have been expressed and shown to form functional gap junctions in Xenopus oocytes. To our knowledge this is the first study to describe gap junction genes in any virus. We hypothesize that the virus innexins disrupt cellular immunity in infected insects by altering normal gap junctional intercellular communication. This would represent a novel mechanism of viral alteration of host physiology, and suggests that gap junctions play a crucial role in coordinating cellular immune responses. Gap junctions, encoded by innexins in insects [1], coordinate multicellular processes. In insects, blood cells (haemocytes) play a critical role in immune responses and gap junctions are known to form between those cells [2,3], yet the functions of gap junctions in
insect immunity are unclear. Pathogenic agents, such as polydnaviruses, may provide a tool to investigate their role in nonmodel systems. The virus family Polydnaviridae is unique in that its members, the ichnoviruses and bracoviruses, are obligately associated with parasitoid wasps [4]. Virions are delivered to host caterpillars during parasitization where virus-encoded transcripts and proteins are produced in the absence of virus replication [4]. In the case of ichnoviruses, virusencoded proteins almost invariably suppress the immune system of the caterpillar, and thus are essential for successful parasitization (reviewed in [5]). Polydnavirus genomes are unusual in that they exhibit simultaneous reductive and proliferative evolution, i.e. few, if any, genes associated with virus replication and packaging are encapsidated, and the packaged genome contains genes generally associated with pathogenesis of the caterpillar host [5,6]. Therefore, genes identified in the ichnovirus genome may be assumed a priori both to be necessary for successful parasitization and to induce either immunological or developmental pathologies in the infected caterpillar. We identified five complete open reading frames in the genomes of two polydnaviruses, CsIV and Hyposoter didymator ichnovirus (HdIV), that shared significant (e-values <10−40) global similarity to innexins (see Table S1 in Supplemental data published with this paper online). These virus innexins, termed vinnexins, encode 22 of 39 invariant residues found in insect innexins (Figure S1), but form a well supported virus-specific clade in gene trees (Figure S2). Two CsIV vinnexins, cs-Vnxd and cs-Vnxg, were expressed in paired Xenopus laevis oocytes and assayed for their ability to form functional gap junctions. Cell pairs in which both cells expressed either cs-Vnxd or csVnxg electrically coupled (Figure 1). Average junctional conductance for these homotypic channels was 1.87 µS for cs-Vnxd (n = 15) and 3.62 µS for cs-Vnxg
Figure 1. Vinnexins make functional gap junctions in Xenopus oocytes. Xenopus laevis oocytes were microinjected with in vitro transcribed vinnexin RNAs or water (negative control). The vitelline membrane was removed to expose the plasma membrane and cells were paired. Cell pairs were recorded 24–48 h later using a double voltage clamp technique. Both oocytes of a pair were clamped to a holding potential (Vh) of –80mV. Sequential 10 mV depolarizing and hyperpolarizing voltage steps (V1) were applied to one cell (cell 1 in (A)) to generate transjunctional voltage, Vj (V1–Vh), and the current, Ij (–I2) required to maintain the neighbour cell (cell 2 in (B–D) at the holding potential was µS) is recorded. Junctional conductance (µ Ij/Vj. Transjunctional voltage steps elicit junctional currents in cell pairs injected with (B) cs-vnxd (0.5 ng) and (C) cs-vnxg (0.5 ng) but not in (D) water-injected control pairs.
(n = 32) following expression of 0.5–1.0 ng RNA, which is within the range previously observed for Drosophila innexins. Heterotypic channels also formed between cell pairs in which one cell expressed cs-Vnxd and the other cs-Vnxg (data not shown). Vinnexin channels were not voltage dependent; junctional current (Ij) was linear at all transjunctional voltage steps (Vj, Figure 1A–C). The vinnexins are transcribed in several tissues of the infected
Current Biology Vol 15 No 13 R492
comparisons as well as experimental procedures are available at http://www.current-biology.com/cgi/ content/full/15/13/R491/DC1/
Figure 2. cs-Vnxq2 protein localizes to haemocyte membranes. Isolated haemocytes from the CsIV-susceptible host caterpillar Heliothis virescens were labelled with anti-peptide antisera generated against a unique sequence in the cytoplasmic region of the cs-Vnxq2 protein. (A) Immunofluorescence confocal microscopy detected no specific signal in control haemocytes. (B) Antisera demonstrated localization (red) to cellular membranes in haemocytes isolated from 3 day post-parasitized H. virescens. Green, FITC-phalloidin labeled F-actin. Note the actin cytoskeleton is disrupted in infected haemocytes hence the absence of cell processes in (B). Scale bar = 10 µm.
caterpillar (Figure S3), concurrently with two newly isolated host lepidopteran innexin-2 orthologues (data not shown). Peptide antisera raised against a unique region of cytoplasmic sequence of the CsIV vinnexin cs-Vnxq2 were used to label isolated haemocytes. No signal was observed in control insects using immunofluorescence microscopy (Figure 2A); by contrast, positive signal was observed consistently in haemocytes of parasitized insects (Figure 2B). Western blots confirmed that the antigen was restricted to infected haemocytes with no detectable signal in other tissues (Figure S4). Our data demonstrate that some ichnoviruses have coopted a host gene family controlling intercellular communication, presumably as a novel means to alter host physiology. An intriguing hypothesis, supported by the observed haemocyte specificity and the known role of ichnoviruses in parasitoid–host interactions, is that vinnexins disrupt host immunity, interfering with haemocytic encapsulation, the primary immune response against haemocoelic parasites in insects. The native encapsulation reaction requires two haemocyte populations interacting over several hours to days in a tightly
regulated cascade of behaviours [2,7]. The mechanisms of regulation during intermediate to terminal stages of encapsulation are unknown but gap junctions are probably involved [8,9]. The vinnexins may alter native gapjunction-mediated intercellular communication among haemocytes by modifying the permeability of host innexin channels, via vinnexin–innexin interactions, or by permitting diffusion through vinnexin channels, thereby abrogating encapsulation. The discovery and initial characterization of the vinnexins in combination with the isolation of caterpillar innexins transcribed in haemocytes provides a set of conceptual and molecular tools for future studies of the functions of gap junctions in cellular immunity. Acknowledgements Thanks to C. Holcroft, R. Hilgarth, J. Kroemer, and E. Fleming for helpful comments and assistance. This work was supported by the University of Kentucky (M.W.T.), the Wellcome Trust and the Leverhulme Trust (P.P.), and the National Science Foundation and the United States Department of Agriculture (B.A.W.). This is publication number 0408-103 of the University of Kentucky Experiment Station. Supplemental data Supplemental data including analysis of vinnexin expression and sequence
References 1. Phelan, P., Stebbings, L.A., Baines, R.A., Bacon, J.P., Davies, J.A., and Ford, C. (1998). Drosophila Shaking-B protein forms gap junctions in paired Xenopus oocytes. Nature 391, 181–184. 2. Grimstone, A.V., Rotheram, S., and Salt, G. (1967). An electronmicroscope study of capsule formation by insect blood cells. J. Cell Sci. 2, 281–292. 3. Churchill, D., Coodin, S., Shivers, R.R., and Caveney, S. (1993). Rapid de novo formation of gap junctions between insect hemocytes in vitro: a freeze-fracture, dye-transfer and patch-clamp study. J. Cell Sci. 104, 763–772. 4. Webb, B.A., Beckage, N.E., Hayakawa, Y., Krell, P.J., Lanzrein, B., Stoltz, D.B., Strand, M.R., and Summers, M.D. (2000). Polydnaviridae. In Virus Taxonomy: The Classification and Nomenclature of Viruses. The 7th Report of the International Committee on Taxonomy of Viruses. M.H.V. van Regenmortel, et al., eds (San Diego: Academic Press), pp. 1167. 5. Kroemer, J.A., and Webb, B.A. (2004). Polydnavirus genes and genomes: Emerging gene families and new insights into polydnavirus replication. Annu. Rev. Entomol. 49, 431–456. 6. Espagne, E., Dupuy, C., Huguet, E., Cattolico, L., Provost, B., Martins, N., Poirie, M., Periquet, G., and Drezen, J.M. (2004). Genome sequence of a polydnavirus: insights into symbiotic virus evolution. Science 306, 286–289. 7. Pech, L.L., and Strand, M.R. (1996). Granular cells are required for encapsulation of foreign targets by insect haemocytes. J. Cell Sci. 109, 2053–2060. 8. Gupta, A.P. (1991). Gap cell junctions, cell adhesion molecules, and molecular basis of encapsulation. In Immmunology of Insects and Other Arthropods. A.P. Gupta, ed. (Boca Raton: CRC Press), pp. 133–167. 9. Lackie, A.M. (1988). Haemocyte behaviour. Adv. Insect Physiol. 21, 85–178. 1Clemson
University, 114 Long Hall, Box 340315, Department of Entomology, Soils, and Plant Sciences, Clemson, SC 29634-0315, USA. 2I.N.R.A.-U.M. II, Laboratoire de Pathologie Comparée, 30380 Saint Christol-lez-Alès, France. 3University of Kentucky, Department of Entomology, S-225 Agricultural Science Center, N, University of Kentucky, Lexington, KY 40546-0091, USA. 4University of Kent, Department of Biosciences, Canterbury, Kent, CT2 7NJ, UK. *E-mail: [email protected]