- Email: [email protected]
Membrane changes associated with assembly of visna virus
VIROLOGY
74, 520-530
Membrane
(1976)
Changes
Associated
M. DUBOIS-DALCQ,’
with Assembly
T. S. REESE,
AND
of Visna Virus
0. NARAYAN
Infectious Diseases Branch and Laboratory of Neuropathology and Neuroanatomical Sciences, Institute of Neurological and Communicative Disorders and Stroke, National Institutes of Health, Maryland 20014; and Department of Neurology, Johns Hopkins University School of Medicine, Maryland 21205 Accepted
June
National Bethesda, Baltimore,
21,1976
The maturation of visna virus at the surface membranes of sheep choroid plexus cells has been studied by freeze-fracture, deep-etching, surface replication, and scanning EM techniques. The results were integrated with data obtained from thin sections. The first changes detectable in the membranes of infected cells consist of flat regions which lack intramembrane particles and show clusters of globular units on corresponding areas of the surface. Protrusion of the cell membrane, characteristic of the budding process, appears to be initiated after a capsid attaches to these modified membrane regions. During the growth of the viral bud, the areas lacking intramembrane particles, the crescent capsids, and the surface covered with globular units increase in size simultaneously. The surface units become more widely spaced as if they were anchored to the bending capsid. At the end of this development, viruses bud off singly or in rows from the soma and long processes of the infected cells. The capsid detaches from the envelope and forms a central core, the surface units become larger and more closely apposed, and free viruses become smaller than viral buds. Thus, visna virus formation requires insertion and growth of surface units probably representing viral-coded proteins, reorganization of host cell membrane proteins under these units, and, initially, attachment of a coiled capsid to the modified membrane.
mately 150 nm in diameter, with a dense, crescent-shaped capsid closely apposed to the membrane. The free viruses are spheres, approximately 100 nm in diameter, with a dense central core. These viral structures, which are able to absorb to uninfected cells (Chippaux-Hyppolite et al., 19721, presumably correspond to the infectious agent obtained by gradient purification (Coward et al., 1970). Both viral buds and particles are covered with spikes (Takemoto et al., 1973; Thormar, 1961). In the present study, the combination of freeze-fracturing, deep-etching, and surface replication technique, used in our previous study of measles virus (Dubois-Dalcq and Reese, 19751, has been applied to sheep choroid plexus cells productively infected with visna virus in order to explore membrane changes occurring during maturation of this virus. Infected cells were
INTRODUCTION
Visna is a slow virus which causes a demyelinating disease in sheep with a prolonged incubation period and protracted clinical course (Haase, 1975; Nathanson et al., 1976; Sigurdsson et al., 1957). This agent is closely related to C-type RNA tumor viruses on the basis of it morphogenesis and the presence of an RNA-directed DNA polymerase (Haase, 1975). The maturation of visna virus has been characterized in several electron microscopic studies using thin sections (Chippaux-Hyppolite et al., 1972; Coward et al., 1970, 1972; Friedmann et al., 1974; MacIntyre et al., 1973, 1974; Takemoto et al., 1971, 1973; Thormar, 1961). Viral buds on the plasmalemma of host cells are approxi’ Author addressed.
to whom
requests
for reprints
should
be 520
Copyright All rights
0 1976 by Academic Press, of reproduction in any form
Inc. reserved.
VISNA
VIRUS-ASSOCIATED
also examined in a scanning electron microscope to determine the distribution of budding. Comparison of these data with data from thin sections makes it possible to elaborate on certain features of the assembly of visna virus. MATERIALS
AND
METHODS
Virus and cells. Visna virus VV, was kindly provided by Dr. D. H. Harter (Columbia University, New York). Dilutions of this virus or a virus pool obtained after 5 days growth in sheep choroid plexus cells were used in some experiments. In others, visna strain K1514 was used. The K1514 strain was obtained from Dr. N. Nathanson (Johns Hopkins University, Baltimore) and was derived from strain K796 passed several times in Icelandic sheep brain. Sheep choroid plexus cells were obtained commercially from Microbiological Associates, Inc., Bethesda, Md. Cells were trypsinized and grown in large Falcon plastic flasks or on round glass coverslips 12 mm in diameter. The growth medium consisted of Eagle’s Minimal Essential Medium (MEM) supplemented with 10 to 15% calf serum, 100 u of penicillin G, and 100 pug of streptomycin sulfate/ml. Confluent monolayers were inoculated with visna virus at a multiplicity of infection of approximately one virus per cell. The virus was allowed to absorb for 2 hr at 37” and the cultures were subsequently refed with a maintenance medium containing 2 to 5% horse or sheep serum and incubated at 37” in 5% CO,. Monolayers were fixed 2 to 9 days postinoculation, as the number of cells showing cytopathic changes increased (Coward et al., 1970; Narayan et al., 1974; Thormar, 1961). Freeze-fracturing and freeze-etching. Cells grown in Falcon flasks were fixed in situ with 1.25% purified glutaraldehyde and 1% paraformaldehyde in 0.08 M cacodylate buffer for 30 min at pH 7.2. Cells were then scraped off and centrifuged in the fixative at 3000 rpm for 30 min. The pellets were fixed again in 4% glutaraldehyde for 1 hr, and rinsed in buffer. For freeze-fracturing, pellets of cells were progressively equilibrated with 25% glycerol
MEMBRANE
CHANGES
521
in water, cut into small squares, and placed on gold alloy disks. They were then rapidly frozen in monochlorodifluoromethane (Freon 22) at - 146” and stored in liquid nitrogen (Dubois-Dalcq and Reese, 1975). For deep-etching, the cell pellets were progressively equilibrated with distilled water and then frozen in a Van Harreveld apparatus modified to prepare specimens for freeze-fracturing (Heuser et al., 1976). When the vacuum was greater than 10Ffi Torr, cells were fractured at -118” in a Balzer 360 M apparatus equipped with an electron beam gun for platinum shadowing. Cells frozen in water were fractured only through the superficial layer since good freezing was usually obtained in the first 20 pm at the surface of the cell layer. Etching was performed for 11/2 to 3 min at -100” prior to replication. Surface replicas. After several rinses in MEM, cells grown on glass coverslips were fixed in 1% glutaraldehyde in 0.1 M cacodylate buffer for 30 min and postfixed in 1% osmium in the same buffer. The coverslips were then quickly passed through increasing concentrations of methanol and critical point-dried in carbon dioxide. The coverslips covered with the dried monolayer were then fitted on the top of the Balzer’s specimen stage and a platinum replica of the cell surface was made at -105” with a vacuum greater than lo-” Torr (Dubois-Dalcq and Reese, 1975). All replicas were cleaned in methanol and then sodium hypochlorite (Chlorox), and mounted on Formvar and carboncoated grids. Positive photographic prints were mounted with origin of platinum shadowing below unless otherwise stated. Viral structures, all from the same surface replica, were measured at 120,000 final magnification using a Hewlett Packard Digitizer. Between 60 to 150 measurements were performed for each parameter. The mean size from each group was compared to other groups using Student’s t test. Scanning electron microscopy. Cells fixed in the same manner as for surface replicas were dehydrated in methanol and critical point-dried in CO,. They were then coated with gold-palladium just before ex-
522
DUBOIS-DALCQ,
REESE
amining them in an ETEC scanning electron microscope. Thin sections. Cells fixed in a manner identical to that used for freeze-fracturing and surface replication were postfixed in 1% osmium tetroxide for 1 hr, stained with uranyl acetate at pH 5, dehydrated in graded alcohols, and embedded in Epon. In some instances, serial thin sections were cut and picked up on Formvar-coated slot grids. Sections were examined in a Philips 201 electron microscopy. RESULTS
From 2 to 9 days after inoculation, an increasing number of abnormal cells were found with the phase microscope. Cells showing these cytopathic changes consisted of spindle- or stellate-shaped cells and multinucleated cells (5 to 10 nuclei). The ultrastructural features of both strains of virus were identical. From 2 days on, cells with many viral buds scattered over their surfaces could be detected. Some of these cells had no alterations in shape while others showed the cytopathic changes described above (Fig. 8). In thin sections, dense crescents, thought to be the viral capsids, were found apposed to the plasmalemma (Fig. 1). Some capsids were small crescents, with diameters less than 100 nm, inserted into small bulges in the membrane (Fig. 1, small arrows), whereas others were larger hemispheres lining the inside of buds up to 150 nm in diameter (Fig. 1, large arrow), The small bulges, on the top of the smallest capsids, might be the result of shrinkage occurring during embedding. Freeze-fracturing the membranes of infected cells at this stage of infection revealed the internal structure of the bud membrane (Fig. 2). The inner leaflet of hemispherical buds was characterized by a paucity of the intramembrane particles 813 nm in diameter which cover the rest of the membrane (Fig. 2, inset). Flat or slightly bowed areas, less than 100 nm in diameter which had few particles (Fig. 2, arrows), were also common. These presumably correspond to the smaller buds which have smaller capsids underlying them. The outer leaflet of the membrane
AND
NARAYAN
covering the viral buds had only a few intramembrane particles but these were also scarce on the outer leaflet of the surrounding cell membrane (Fig. 3). Deep-etching infected cells disclosed a thin rim of their true outer surface which could be compared to adjacent regions of the freeze-fractured membrane. The surface of buds was covered with globular units where the inner leaflet had few intramembrane particles (Fig. 4). Large expanses of the true outer surface of many infected cells could be studied with the surface replica and scanning techniques. Numerous viral buds of varying size were found on a single cell (Figs. 5 and 8). Occasionally, several viral buds of similar size were aligned in a row (Fig. 8, black arrow) sometimes along a bundle of fine extracellular filaments (Fig. 5, between the 2 large arrows) (McIntyre et al., 1974). Structural details of the bud surface were also clear in surface replicas, and were characterized by globular units similar in size to those exposed by deep-etching. Globular units of various sizes were also seen on the rest of the cell surface (Figs. 5 to 7) but their number was not greater than that in control cells. In some cells, the only changes detectable in surface replicas were aggregates of globular units on flat regions of the cell surface (Fig. 6). The small bulges (Fig. 5, number l), which probably were associated with the smaller capsids, were covered with approximately 12 globular units. The units in these small aggregates were relatively heterogeneous in size, with a mean diameter of 13 t 0.6 nm (SEM) and spacing of 20 2 0.4 nm. Sometimes they were more spaced as if they were still being assembled (Fig. 6, left). In thin sections, similar buds were covered with aggregates of surface projections 8 nm in height and with a spacing of approximately 20 nm. A large number of measurements was not performed on sectioned buds because of the poorly defined limits and weak electron density of their projections. The shadow of the larger buds varied with respect to size and shape, presumably depending on the stage of viral bud forma-
FIG. 1. Different types of viral buds in a cell with no detectable cytopathic changes. The buds are characterized by an electron dense plate of capsid material closely apposed to the cell membrane. Smaller capsids (small arrows) are associated with small protrusions of the membrane whereas larger membrane protrusions contain larger hemispherical capsids (large arrow). x 57,000. FIG. 2. Freeze-fractured plasmalemma from an infected cell at a stage similar to that in Fig. 1. On the exposed inner leaflet of the membrane are numerous 8-13 nm particles, except at viral buds where they are infrequent. The smaller circular regions with few particles (arrows) are also less elevated and probably correspond to the smaller buds shown in Fig. 1. Inset shows one of these protrusions at higher magnification. X 76,000; inset x 114,000. 523
524
DUBOIS-DALCQ,
REESE
AND
NARAYAN
FIG. 3. Outer leaflet of the plasmalemma of an infected cell. This view of the viral buds is complementary to that shown in Fig. 2. Particles are infrequent on the outer leaflet of both the cell membrane and the bud membrane. x 120,000. FIG. 4. Surface membrane of an infected cell which was freeze-fractured and then deep-etched to expose the true outer surfaces of adjacent unfractured membranes. Arrows mark the boundaries between areas of the membrane exposed by fracturing and areas of true outer surface of the cell and viral membranes which are exposed by etching. The true outer surfaces of the two viral buds are covered with globular units but the interior of the bud membrane exposed by fracturing has no particles. Conversely, the interior of the cell membrane (asterisk) is marked by particles. Particles at arrowheads are larger on the true outer surface of the cell membrane below (star). x 120,000.
FIG. 5. Replica of the surface of an infected cell showing numerous buds of various sizes scattered over the cell surface. Small buds (between the two large arrows) are aligned next to a bundle of extracellular filaments (small arrow). The smallest bud formations cast almost no shadow (at 1). As they protrude further, a conical shadow appears (at 2). Further protrusion of the bud with narrowing of its neck, or a complete pinching off of the virus, results in a shadow with an ellipsoidal shape (at 3). Numerous globular units are seen on the viral buds and are similar in size and distribution to those revealed by deep-etching (Fig. 4). Views of the bud surface at higher magnification are shown in Fig. 6 and 7. x 36,000. FIGS. 6 and 7. Details of the surfaces of small and large viral buds such as those shown in Fig. 5. A few globular units of various sizes are scattered on the cell surface, while viral buds are covered with large globular units. In Fig. 6, these aggregates of globular units are on a flat area of the cell surface. The bud on the left has globular units that are sparse and loosely packed. In Fig. 7, similar small buds on the right, covered with globular units, protrude slightly. The two large buds with ellipsoidal shadows have larger globular units than those on the small buds indicated by the arrow. A medium-size bud with a conical shadow is seen in the left lower corner. Inset shows a thin section through a small viral bud at about twice the magnification of the surface replicas. Clusters of faintly stained projections on the surface membrane (arrow) probably correspond to the globular units. x 90,000; inset x 175,000.
VISNA
VIRUS-ASSOCIATED
MEMBRANE
FIGURES 5-l
CHANGES
525
526
DUBOIS-DALCQ,
REESE
tion (Figs. 5 and 7). Viral buds which protruded from the cell membrane produced shadows and the shape of the bud was deduced from the shape of the shadow. Medium-size buds (Fig. 5, number 2, and Fig. 7, lower left corner) had a conical shadow because of the absence of a narrow neck. The globular units on these buds had a diameter similar to that of the units on the flat buds (13 1+_0.3 nm), but their spacing was larger, 25 t 0.7 nm as compared to 20 * 0.4 nm (p < 0.001). When the shadow had an ellipsoidal shape (Fig. 5, number 3; Fig. 7), the bud was presumably connected to the surface by a narrow neck or was possibly a detached virion on the surface of the cell. The globular units in these buds were larger (16 + 0.6 nm) than those in the smaller buds (p < O.OOl), but their spacing (24 ? 0.6 nm) was similar to that on the medium-size buds (25 ? 0.7 nm). We em-
AND
NARAYAN
phasize that these are measurements of the platinum coating and are not the actual size of the globular units. However, differences in size and spacing observed in the replica probably reflect true differences in the size and spacing of globular units. Cells showing marked changes in shape, as well as budding viruses, could be identified in the scanning electron microscope (Fig. 8). Some cells were stellate and had long processes spreading on the glass or over other cell surfaces. Their processes contained filaments, which were identified as such in thin sections, and often had several budding viruses along them (Figs. B-10). Some viruses also budded from the edges of cells near their point of attachment to the substrate (Fig. 8, white and black arrows). Stellate cells were often surrounded by
FIG. 8. Scanning electron micrograph of a stellate cell 8 days after inoculation. Viral buds or free viruses cover the surface of the soma and one of the elongated processes (P) of this cell. Occasionally, buds are aligned in straight rows (black arrow). Similar spherical bodies, thought to be free viruses, lie on the glass substrate (white arrow). The shapes of some of the viral buds are apparent on the lateral border of this cell where they are seen in profile (black and white arrows) and may be compared to the buds in Fig. 1. F is probably a bundle of filaments, although at this level of resolution it is not possible to distinguish them clearly from thin processes such as those shown in Fig. 9. x 16,150.
VISNA
VIRUS-ASSOCIATED
regions of substrate free of other cells and by detached viruses. These were 100 nm in diameter and could be clearly identified on the substrate surfaces. They were either isolated (Fig. 8, white arrow) or in clumps (Fig. 9, inset) and approximately one-third smaller than the large viral buds which were 150 nm in diameter. Similar clusters of viruses of various sizes, presumably detached, lay on unmodified cell surfaces adjacent to stellate cells (Fig. 9). Measurement of globular units on the smallest free viruses showed that they were larger (18 + 0.5 nm) than those on small buds (13 -+ 0.3, p < 0.001) and somewhat larger than those on the larger buds with ellipsoidal shadows (p < 0.021. In contrast, the spacing between units on the free virus was reduced (21 + 0.4 nm) when compared to the large buds (24 & 0.6, p < 0.001). In serial thin sections, small detached viruses had a dense central core (Fig. 10, large arrow) and, after freeze-fracturing, their membrane was found to be devoid of particles. Large (up to 200 nm) and irregularly shaped detached viral structures were also identified in surface replicas (Fig. 9) and serial thin sections. Some had small membrane excrescences and many contained membrane vesicles. Others contained one or several cores of various shapes or, sometimes, several crescent capsids (Dubois-Dalcq, 1976; Haase, 1975). DISCUSSION
A series of viral-induced changes occurs at the surface membrane of visna-infected cells. The end results of these changes is the release of virions from the cell by budding. All these changes can coexist on a single cell and are thought to represent a series of stages in viral bud formation, for reasons discussed below. The most subtle changes detected in the membrane consist of flat circular regions which lack intramembrane particles and show clusters of globular units on corresponding areas of the membrane surface. Since these modified regions are sometimes the only changed regions on the plasmalemma of an infected cell, they are thought to be early changes preceding the formation of viral buds. Whether these changes precede the attachment of nucleocapsids to the
MEMBRANE
CHANGES
527
membrane is not clear. Indeed, retraction of the surface membrane in cells embedded for thin sections could account for the observation that capsids attached to the surface membrane are always curved. Nevertheless, the early attachment of capsids to modified regions of the cell membrane suggests that they play a role in the initiation of the budding process. No intramembrane particles, such as the small ones seen in maturation of SSPE measles virus (Dubois-Dalcq and Reese, 19751, were found at sites of capsid attachment. Curved sites of visna capsid attachments were almost devoid of particles. During the growth of the viral bud, the areas lacking intramembrane particles, the crescent capsids, and the surface covered with globular units increased in size simultaneously. These three changes, which characterize viral bud formation in visna infection, can be interpreted in a manner similar to observations on RNA tumor viruses. An increasing number of host membrane proteins, at least the ones represented by the intramembrane particles, are excluded from the viral bud (Sheffield, 19741, the capsids develop an increasing number of coils (Sarkar et al., 19711, and a number of surface units, probably composed of viral-coded proteins (Demsey and Stackpole, 19751, are added to the bud periphery. Despite the lack of visible attachment sites, the surface units may be anchored to the capsid and both elements might then form membrane complexes similar to those observed in paramyxovirus budding (Dubois-Dalcq and Reese, 1975). The increase in size of the attached capsid, which would require stretching of the surface membrane in order for it to assume a spherical shape, could explain the increased distance between surface globular units on the viral buds. At the end of development, the viruses presumably bud off singly and randomly from the infected cell soma, as seen in other types of infections (Holmes, 1975; Panem and Kirsten, 1975). Sometimes groups of aligned buds appear to form synchronously and budding sites are also seen along the processes of stellate cells as in Sindbis and vesicular stomatitis virus-infected cells (Birdwell et aE., 1973; Birdwell
FIG. 9. Replica of the cell surface 8 days after inoculation, showing an elongated process from a stellate cell (the cell body is outside of the field illustrated) lying on the surface of another cell. Numerous protrusions of various sizes mark the surface of the elongated process and these protrusions have on their surfaces the globular units characteristic of viral buds (arrows). Clusters of viral particles are scattered over the rest of the cell surface. Some of these are irregularly shaped and larger than others. Similar clusters of free virus occasionally adhere to the glass, as shown in the inset. These free viruses are smaller than the large viral buds but the globular units on their surfaces are larger so that they appear almost confluent. (Compare inset to Fig. 7.) Origin of platinum shadowing is on the right. x 72,200; inset x 85,500. FIG. 10. Thin section through an elongated process from a stellate cell 8 days after inoculation. These processes are characterized by their content of filaments (F) and the numerous viral buds on their surface (small arrows). Viral particles are smaller then the buds and characterized by a dense core (large arrow). X 71,250.
528
VISNA
VIRUS-ASSOCIATED
and Strauss, 1974). The mechanism by which these events are controlled was not evident in our study. The final step in viral maturation occurs after the virus has been released from the cell, and is characterized by a reduction in size of viral particles. The capsid detaches from the envelope, as if it were uncoiling, and forms a central core. The membrane complexes, assumed to be responsible for capsid attachment, must consequently be broken, allowing greater plasticity of the viral membrane and possibly permitting the loss of envelope membrane as the virus gets smaller. How the membrane is removed from the viral envelope has not been clarified in this study although the existence of intermediate stages between the large crescent buds and the small free viruses was observed. The free viruses are covered with large and closely apposed surface globular units that make a continuous coat on the detached virus. Since the surface globular units are present throughout viral maturation, they constitute a marker for viral structures. They resemble structures described on the surface of various oncornaviruses (de Harven et al., 1973; Demsey and Stackpole, 1975; Nermut et al., 1972; Panem and Kirsten, 1975; Sheffield, 1973, 1974) and presumably correspond to glycoproteins organized as clusters of spikes on the surface of visna virus (Coward et al., 1972; Haase, 1975; Takemoto et al., 1973; Thormar, 1961). The increase in size of the surface units observed during viral formation could be due to continuous addition of viral proteins to the bud surface, in a manner analogous to the widening of surface strands in productive measles infection (Dubois-Dalcq and Reese, 1975). ACKNOWLEDGMENTS The technical help of Kathy Worthington and Frank Nolan is gratefully acknowledged. We are also grateful to Barbara Reese for taking the Scanning electron micrographs, and to Drs. H. Panitch and H. McFarland for reviewing the manuscript. REFERENCES BIRDWELL, C. R., STRAUSS, E. G., and STRAUSS, J. H. (1973). Replication of sindbis virus. III. An electron microscopic study of virus maturation using
MEMBRANE
CHANGES
529
the surface replica technique. Virology 56, 429438. BIRDWELL, C. R., and STRAUSS, J. H. (1974). Maturation of vesicular stomatitis virus: Electron microscopy of surface replicas of infected cells. Virology 59, 587-590. CHIPPAUX-HYPPOLITE, C., TARANGER, C., TAMALET, J., PAUTRAT, G., and BRAHIC, M. (1972). Aspects ultrastructuraux du virus visna en cultures cellulaires. Ann. Inst. Pasteur 123, 409-420. COWARD, J. E., HARTER, D. H., and MORGAN, C. (1970). Electron microscopic observations of visna virus-infected cell cultures. Virology 40, 10301038. COWARD, J. E., HARTER, D. H., Hsu, K. C., and MORGAN, C. (1972). Ferritin-conjugated antibody labeling of visna virus. Virology 50, 925-930. DEHARVEN, E., LAMPEN, N., SATO, T., and FRIEND, C. (1973). Scanning electron microscopy of cells infected with a murine leukemia virus. Virology 51, 240-243. DEMSEY, A., and STACKPOLE, C. W. (19751. Cell surface membrane alterations during murine leukemia virus morphogenesis. J. Cell Biol. 67, 92a. DUBOIS-DALCQ, M., and REESE, T. S. (1975). Structural changes in the membrane of vero cells infected with a paramyxovirus. J. Cell Biol. 67,551565. DUBOIS-DALCQ, M. (1976). In “Workshop on the Application of Modern Biological Research to Investigation of Slow Virus Infections of the CNS.” Wurzburg. (Ter Meulen and Katz, eds.). Springer Verlag, New York/Berlin, In press. FRIEDMANN, A., COWARD, J. E., HARTER, D. H., LIPSET, J. S., and MORGAN, C. (1974). Electron microscopic studies of visna virus ribonucleic acid. J. Gen. Virol. 25, 93-104. HAASE, A. T. (1975). The slow infection caused by visna virus. Cur-r. Top. Microbial. Immunol. 72, 102-156. HEUSER, J. E., LANDIS, S. M. D., and REESE, T. S. (1976). Preservation of rapidly changing structures in synapses by rapid freezing. Cold Spring Harbor Symposium: The Synapse. HOLMES, K. V. (1975). Scanning electron microscopic studies of virus-infected cells. I. Cytopathic effects and maturation of vesicular stomatitis virus in L2 cells. J. Viral. 15, 355-362. MACINTYRE, E. H. WINTERSGILL, C. J., and VATTER, A. E. (1973). Visna virus infection of sheep and human cells in vitro: An ultrastructural study. J. Cell Sci. 13, 1733191. MACINTYRE, E. H., WINTERGILL, C. J., and VATTER, A. E. (1974). Prolonged culture of visna virus in human astrocytes. Beitr. Pathol. 152, 163-178. NATHANSON, N., PANITCH, H., and PETURSSON, G. (19761. Pathogenesis of visna: Review and speculation. In “Slow Virus Diseases of Animals and Man” (R. H. Kimberlin, ed.), pp. 115-131. North
530 Holland
DUBOIS-DALCQ,
REESE
Publishing Co., Amsterdam. O., SILVERSTEIN, A. M., PRICE, D., and R. T. (1974). Visna virus infection of American lambs. Science 183, 1202-1203. NERMUT, M. V., FRANK, H., and SCHAFER, W. (1972). Properties of mouse leukemia viruses. III. EM appearance as revealed after conventional preparation techniques as well as freeze-drying and freeze-etching. Virology 49, 345-358. PANEM, S., and KIRSTEN, W. H. (1975). Secondary scanning electron microscopy of cells infected with murine oncornaviruses. Virology 63, 447-458. SHEFFIELD, J. B. (1973). Envelope of mouse mammary tumor virus studied by freeze-etching and freeze-fracture techniques. J. Viral. 12, 616-624. SHEFFIELD, J. B. (1974). Membrane alterations which accompany MuMTV maturation. I. Studies by freeze-cleave techniques. Virology 57, 287-290. NARAYAN, JOHNSON,
AND
NARAYAN
B., PALSSON, P. A., and GRIMSSON, H. (1957). Visna, a demyelinating transmissible disease of sheep. J. Neuropathal. Exp. Neural. 16, 389-403. TAKEMOTO, K. K., MATTERN, C. F. T., STONE, L. B., COE, J. E., and LAVELLE, G. (19711. Antigenic and morphological similarities of progressive pneumonia virus, a recently isolated “slow virus” of sheep, to visna and Maedi Viruses. J. Virol. 7, 301-308. TAKEMOTO, K. K., AOKI, T., GARON, C., and STURM, M. M. (1973). Comparative studies of visna, progressive pneumonia and Rous sarcoma viruses by electron microscopy. J. Nat. Cancer Inst. 50, 543545. THORMAR, H. (1961). An electron microscope study of tissue cultures infected with visna virus. Virology 14, 463-475.
SIGURD~SON,
74, 520-530
Membrane
(1976)
Changes
Associated
M. DUBOIS-DALCQ,’
with Assembly
T. S. REESE,
AND
of Visna Virus
0. NARAYAN
Infectious Diseases Branch and Laboratory of Neuropathology and Neuroanatomical Sciences, Institute of Neurological and Communicative Disorders and Stroke, National Institutes of Health, Maryland 20014; and Department of Neurology, Johns Hopkins University School of Medicine, Maryland 21205 Accepted
June
National Bethesda, Baltimore,
21,1976
The maturation of visna virus at the surface membranes of sheep choroid plexus cells has been studied by freeze-fracture, deep-etching, surface replication, and scanning EM techniques. The results were integrated with data obtained from thin sections. The first changes detectable in the membranes of infected cells consist of flat regions which lack intramembrane particles and show clusters of globular units on corresponding areas of the surface. Protrusion of the cell membrane, characteristic of the budding process, appears to be initiated after a capsid attaches to these modified membrane regions. During the growth of the viral bud, the areas lacking intramembrane particles, the crescent capsids, and the surface covered with globular units increase in size simultaneously. The surface units become more widely spaced as if they were anchored to the bending capsid. At the end of this development, viruses bud off singly or in rows from the soma and long processes of the infected cells. The capsid detaches from the envelope and forms a central core, the surface units become larger and more closely apposed, and free viruses become smaller than viral buds. Thus, visna virus formation requires insertion and growth of surface units probably representing viral-coded proteins, reorganization of host cell membrane proteins under these units, and, initially, attachment of a coiled capsid to the modified membrane.
mately 150 nm in diameter, with a dense, crescent-shaped capsid closely apposed to the membrane. The free viruses are spheres, approximately 100 nm in diameter, with a dense central core. These viral structures, which are able to absorb to uninfected cells (Chippaux-Hyppolite et al., 19721, presumably correspond to the infectious agent obtained by gradient purification (Coward et al., 1970). Both viral buds and particles are covered with spikes (Takemoto et al., 1973; Thormar, 1961). In the present study, the combination of freeze-fracturing, deep-etching, and surface replication technique, used in our previous study of measles virus (Dubois-Dalcq and Reese, 19751, has been applied to sheep choroid plexus cells productively infected with visna virus in order to explore membrane changes occurring during maturation of this virus. Infected cells were
INTRODUCTION
Visna is a slow virus which causes a demyelinating disease in sheep with a prolonged incubation period and protracted clinical course (Haase, 1975; Nathanson et al., 1976; Sigurdsson et al., 1957). This agent is closely related to C-type RNA tumor viruses on the basis of it morphogenesis and the presence of an RNA-directed DNA polymerase (Haase, 1975). The maturation of visna virus has been characterized in several electron microscopic studies using thin sections (Chippaux-Hyppolite et al., 1972; Coward et al., 1970, 1972; Friedmann et al., 1974; MacIntyre et al., 1973, 1974; Takemoto et al., 1971, 1973; Thormar, 1961). Viral buds on the plasmalemma of host cells are approxi’ Author addressed.
to whom
requests
for reprints
should
be 520
Copyright All rights
0 1976 by Academic Press, of reproduction in any form
Inc. reserved.
VISNA
VIRUS-ASSOCIATED
also examined in a scanning electron microscope to determine the distribution of budding. Comparison of these data with data from thin sections makes it possible to elaborate on certain features of the assembly of visna virus. MATERIALS
AND
METHODS
Virus and cells. Visna virus VV, was kindly provided by Dr. D. H. Harter (Columbia University, New York). Dilutions of this virus or a virus pool obtained after 5 days growth in sheep choroid plexus cells were used in some experiments. In others, visna strain K1514 was used. The K1514 strain was obtained from Dr. N. Nathanson (Johns Hopkins University, Baltimore) and was derived from strain K796 passed several times in Icelandic sheep brain. Sheep choroid plexus cells were obtained commercially from Microbiological Associates, Inc., Bethesda, Md. Cells were trypsinized and grown in large Falcon plastic flasks or on round glass coverslips 12 mm in diameter. The growth medium consisted of Eagle’s Minimal Essential Medium (MEM) supplemented with 10 to 15% calf serum, 100 u of penicillin G, and 100 pug of streptomycin sulfate/ml. Confluent monolayers were inoculated with visna virus at a multiplicity of infection of approximately one virus per cell. The virus was allowed to absorb for 2 hr at 37” and the cultures were subsequently refed with a maintenance medium containing 2 to 5% horse or sheep serum and incubated at 37” in 5% CO,. Monolayers were fixed 2 to 9 days postinoculation, as the number of cells showing cytopathic changes increased (Coward et al., 1970; Narayan et al., 1974; Thormar, 1961). Freeze-fracturing and freeze-etching. Cells grown in Falcon flasks were fixed in situ with 1.25% purified glutaraldehyde and 1% paraformaldehyde in 0.08 M cacodylate buffer for 30 min at pH 7.2. Cells were then scraped off and centrifuged in the fixative at 3000 rpm for 30 min. The pellets were fixed again in 4% glutaraldehyde for 1 hr, and rinsed in buffer. For freeze-fracturing, pellets of cells were progressively equilibrated with 25% glycerol
MEMBRANE
CHANGES
521
in water, cut into small squares, and placed on gold alloy disks. They were then rapidly frozen in monochlorodifluoromethane (Freon 22) at - 146” and stored in liquid nitrogen (Dubois-Dalcq and Reese, 1975). For deep-etching, the cell pellets were progressively equilibrated with distilled water and then frozen in a Van Harreveld apparatus modified to prepare specimens for freeze-fracturing (Heuser et al., 1976). When the vacuum was greater than 10Ffi Torr, cells were fractured at -118” in a Balzer 360 M apparatus equipped with an electron beam gun for platinum shadowing. Cells frozen in water were fractured only through the superficial layer since good freezing was usually obtained in the first 20 pm at the surface of the cell layer. Etching was performed for 11/2 to 3 min at -100” prior to replication. Surface replicas. After several rinses in MEM, cells grown on glass coverslips were fixed in 1% glutaraldehyde in 0.1 M cacodylate buffer for 30 min and postfixed in 1% osmium in the same buffer. The coverslips were then quickly passed through increasing concentrations of methanol and critical point-dried in carbon dioxide. The coverslips covered with the dried monolayer were then fitted on the top of the Balzer’s specimen stage and a platinum replica of the cell surface was made at -105” with a vacuum greater than lo-” Torr (Dubois-Dalcq and Reese, 1975). All replicas were cleaned in methanol and then sodium hypochlorite (Chlorox), and mounted on Formvar and carboncoated grids. Positive photographic prints were mounted with origin of platinum shadowing below unless otherwise stated. Viral structures, all from the same surface replica, were measured at 120,000 final magnification using a Hewlett Packard Digitizer. Between 60 to 150 measurements were performed for each parameter. The mean size from each group was compared to other groups using Student’s t test. Scanning electron microscopy. Cells fixed in the same manner as for surface replicas were dehydrated in methanol and critical point-dried in CO,. They were then coated with gold-palladium just before ex-
522
DUBOIS-DALCQ,
REESE
amining them in an ETEC scanning electron microscope. Thin sections. Cells fixed in a manner identical to that used for freeze-fracturing and surface replication were postfixed in 1% osmium tetroxide for 1 hr, stained with uranyl acetate at pH 5, dehydrated in graded alcohols, and embedded in Epon. In some instances, serial thin sections were cut and picked up on Formvar-coated slot grids. Sections were examined in a Philips 201 electron microscopy. RESULTS
From 2 to 9 days after inoculation, an increasing number of abnormal cells were found with the phase microscope. Cells showing these cytopathic changes consisted of spindle- or stellate-shaped cells and multinucleated cells (5 to 10 nuclei). The ultrastructural features of both strains of virus were identical. From 2 days on, cells with many viral buds scattered over their surfaces could be detected. Some of these cells had no alterations in shape while others showed the cytopathic changes described above (Fig. 8). In thin sections, dense crescents, thought to be the viral capsids, were found apposed to the plasmalemma (Fig. 1). Some capsids were small crescents, with diameters less than 100 nm, inserted into small bulges in the membrane (Fig. 1, small arrows), whereas others were larger hemispheres lining the inside of buds up to 150 nm in diameter (Fig. 1, large arrow), The small bulges, on the top of the smallest capsids, might be the result of shrinkage occurring during embedding. Freeze-fracturing the membranes of infected cells at this stage of infection revealed the internal structure of the bud membrane (Fig. 2). The inner leaflet of hemispherical buds was characterized by a paucity of the intramembrane particles 813 nm in diameter which cover the rest of the membrane (Fig. 2, inset). Flat or slightly bowed areas, less than 100 nm in diameter which had few particles (Fig. 2, arrows), were also common. These presumably correspond to the smaller buds which have smaller capsids underlying them. The outer leaflet of the membrane
AND
NARAYAN
covering the viral buds had only a few intramembrane particles but these were also scarce on the outer leaflet of the surrounding cell membrane (Fig. 3). Deep-etching infected cells disclosed a thin rim of their true outer surface which could be compared to adjacent regions of the freeze-fractured membrane. The surface of buds was covered with globular units where the inner leaflet had few intramembrane particles (Fig. 4). Large expanses of the true outer surface of many infected cells could be studied with the surface replica and scanning techniques. Numerous viral buds of varying size were found on a single cell (Figs. 5 and 8). Occasionally, several viral buds of similar size were aligned in a row (Fig. 8, black arrow) sometimes along a bundle of fine extracellular filaments (Fig. 5, between the 2 large arrows) (McIntyre et al., 1974). Structural details of the bud surface were also clear in surface replicas, and were characterized by globular units similar in size to those exposed by deep-etching. Globular units of various sizes were also seen on the rest of the cell surface (Figs. 5 to 7) but their number was not greater than that in control cells. In some cells, the only changes detectable in surface replicas were aggregates of globular units on flat regions of the cell surface (Fig. 6). The small bulges (Fig. 5, number l), which probably were associated with the smaller capsids, were covered with approximately 12 globular units. The units in these small aggregates were relatively heterogeneous in size, with a mean diameter of 13 t 0.6 nm (SEM) and spacing of 20 2 0.4 nm. Sometimes they were more spaced as if they were still being assembled (Fig. 6, left). In thin sections, similar buds were covered with aggregates of surface projections 8 nm in height and with a spacing of approximately 20 nm. A large number of measurements was not performed on sectioned buds because of the poorly defined limits and weak electron density of their projections. The shadow of the larger buds varied with respect to size and shape, presumably depending on the stage of viral bud forma-
FIG. 1. Different types of viral buds in a cell with no detectable cytopathic changes. The buds are characterized by an electron dense plate of capsid material closely apposed to the cell membrane. Smaller capsids (small arrows) are associated with small protrusions of the membrane whereas larger membrane protrusions contain larger hemispherical capsids (large arrow). x 57,000. FIG. 2. Freeze-fractured plasmalemma from an infected cell at a stage similar to that in Fig. 1. On the exposed inner leaflet of the membrane are numerous 8-13 nm particles, except at viral buds where they are infrequent. The smaller circular regions with few particles (arrows) are also less elevated and probably correspond to the smaller buds shown in Fig. 1. Inset shows one of these protrusions at higher magnification. X 76,000; inset x 114,000. 523
524
DUBOIS-DALCQ,
REESE
AND
NARAYAN
FIG. 3. Outer leaflet of the plasmalemma of an infected cell. This view of the viral buds is complementary to that shown in Fig. 2. Particles are infrequent on the outer leaflet of both the cell membrane and the bud membrane. x 120,000. FIG. 4. Surface membrane of an infected cell which was freeze-fractured and then deep-etched to expose the true outer surfaces of adjacent unfractured membranes. Arrows mark the boundaries between areas of the membrane exposed by fracturing and areas of true outer surface of the cell and viral membranes which are exposed by etching. The true outer surfaces of the two viral buds are covered with globular units but the interior of the bud membrane exposed by fracturing has no particles. Conversely, the interior of the cell membrane (asterisk) is marked by particles. Particles at arrowheads are larger on the true outer surface of the cell membrane below (star). x 120,000.
FIG. 5. Replica of the surface of an infected cell showing numerous buds of various sizes scattered over the cell surface. Small buds (between the two large arrows) are aligned next to a bundle of extracellular filaments (small arrow). The smallest bud formations cast almost no shadow (at 1). As they protrude further, a conical shadow appears (at 2). Further protrusion of the bud with narrowing of its neck, or a complete pinching off of the virus, results in a shadow with an ellipsoidal shape (at 3). Numerous globular units are seen on the viral buds and are similar in size and distribution to those revealed by deep-etching (Fig. 4). Views of the bud surface at higher magnification are shown in Fig. 6 and 7. x 36,000. FIGS. 6 and 7. Details of the surfaces of small and large viral buds such as those shown in Fig. 5. A few globular units of various sizes are scattered on the cell surface, while viral buds are covered with large globular units. In Fig. 6, these aggregates of globular units are on a flat area of the cell surface. The bud on the left has globular units that are sparse and loosely packed. In Fig. 7, similar small buds on the right, covered with globular units, protrude slightly. The two large buds with ellipsoidal shadows have larger globular units than those on the small buds indicated by the arrow. A medium-size bud with a conical shadow is seen in the left lower corner. Inset shows a thin section through a small viral bud at about twice the magnification of the surface replicas. Clusters of faintly stained projections on the surface membrane (arrow) probably correspond to the globular units. x 90,000; inset x 175,000.
VISNA
VIRUS-ASSOCIATED
MEMBRANE
FIGURES 5-l
CHANGES
525
526
DUBOIS-DALCQ,
REESE
tion (Figs. 5 and 7). Viral buds which protruded from the cell membrane produced shadows and the shape of the bud was deduced from the shape of the shadow. Medium-size buds (Fig. 5, number 2, and Fig. 7, lower left corner) had a conical shadow because of the absence of a narrow neck. The globular units on these buds had a diameter similar to that of the units on the flat buds (13 1+_0.3 nm), but their spacing was larger, 25 t 0.7 nm as compared to 20 * 0.4 nm (p < 0.001). When the shadow had an ellipsoidal shape (Fig. 5, number 3; Fig. 7), the bud was presumably connected to the surface by a narrow neck or was possibly a detached virion on the surface of the cell. The globular units in these buds were larger (16 + 0.6 nm) than those in the smaller buds (p < O.OOl), but their spacing (24 ? 0.6 nm) was similar to that on the medium-size buds (25 ? 0.7 nm). We em-
AND
NARAYAN
phasize that these are measurements of the platinum coating and are not the actual size of the globular units. However, differences in size and spacing observed in the replica probably reflect true differences in the size and spacing of globular units. Cells showing marked changes in shape, as well as budding viruses, could be identified in the scanning electron microscope (Fig. 8). Some cells were stellate and had long processes spreading on the glass or over other cell surfaces. Their processes contained filaments, which were identified as such in thin sections, and often had several budding viruses along them (Figs. B-10). Some viruses also budded from the edges of cells near their point of attachment to the substrate (Fig. 8, white and black arrows). Stellate cells were often surrounded by
FIG. 8. Scanning electron micrograph of a stellate cell 8 days after inoculation. Viral buds or free viruses cover the surface of the soma and one of the elongated processes (P) of this cell. Occasionally, buds are aligned in straight rows (black arrow). Similar spherical bodies, thought to be free viruses, lie on the glass substrate (white arrow). The shapes of some of the viral buds are apparent on the lateral border of this cell where they are seen in profile (black and white arrows) and may be compared to the buds in Fig. 1. F is probably a bundle of filaments, although at this level of resolution it is not possible to distinguish them clearly from thin processes such as those shown in Fig. 9. x 16,150.
VISNA
VIRUS-ASSOCIATED
regions of substrate free of other cells and by detached viruses. These were 100 nm in diameter and could be clearly identified on the substrate surfaces. They were either isolated (Fig. 8, white arrow) or in clumps (Fig. 9, inset) and approximately one-third smaller than the large viral buds which were 150 nm in diameter. Similar clusters of viruses of various sizes, presumably detached, lay on unmodified cell surfaces adjacent to stellate cells (Fig. 9). Measurement of globular units on the smallest free viruses showed that they were larger (18 + 0.5 nm) than those on small buds (13 -+ 0.3, p < 0.001) and somewhat larger than those on the larger buds with ellipsoidal shadows (p < 0.021. In contrast, the spacing between units on the free virus was reduced (21 + 0.4 nm) when compared to the large buds (24 & 0.6, p < 0.001). In serial thin sections, small detached viruses had a dense central core (Fig. 10, large arrow) and, after freeze-fracturing, their membrane was found to be devoid of particles. Large (up to 200 nm) and irregularly shaped detached viral structures were also identified in surface replicas (Fig. 9) and serial thin sections. Some had small membrane excrescences and many contained membrane vesicles. Others contained one or several cores of various shapes or, sometimes, several crescent capsids (Dubois-Dalcq, 1976; Haase, 1975). DISCUSSION
A series of viral-induced changes occurs at the surface membrane of visna-infected cells. The end results of these changes is the release of virions from the cell by budding. All these changes can coexist on a single cell and are thought to represent a series of stages in viral bud formation, for reasons discussed below. The most subtle changes detected in the membrane consist of flat circular regions which lack intramembrane particles and show clusters of globular units on corresponding areas of the membrane surface. Since these modified regions are sometimes the only changed regions on the plasmalemma of an infected cell, they are thought to be early changes preceding the formation of viral buds. Whether these changes precede the attachment of nucleocapsids to the
MEMBRANE
CHANGES
527
membrane is not clear. Indeed, retraction of the surface membrane in cells embedded for thin sections could account for the observation that capsids attached to the surface membrane are always curved. Nevertheless, the early attachment of capsids to modified regions of the cell membrane suggests that they play a role in the initiation of the budding process. No intramembrane particles, such as the small ones seen in maturation of SSPE measles virus (Dubois-Dalcq and Reese, 19751, were found at sites of capsid attachment. Curved sites of visna capsid attachments were almost devoid of particles. During the growth of the viral bud, the areas lacking intramembrane particles, the crescent capsids, and the surface covered with globular units increased in size simultaneously. These three changes, which characterize viral bud formation in visna infection, can be interpreted in a manner similar to observations on RNA tumor viruses. An increasing number of host membrane proteins, at least the ones represented by the intramembrane particles, are excluded from the viral bud (Sheffield, 19741, the capsids develop an increasing number of coils (Sarkar et al., 19711, and a number of surface units, probably composed of viral-coded proteins (Demsey and Stackpole, 19751, are added to the bud periphery. Despite the lack of visible attachment sites, the surface units may be anchored to the capsid and both elements might then form membrane complexes similar to those observed in paramyxovirus budding (Dubois-Dalcq and Reese, 1975). The increase in size of the attached capsid, which would require stretching of the surface membrane in order for it to assume a spherical shape, could explain the increased distance between surface globular units on the viral buds. At the end of development, the viruses presumably bud off singly and randomly from the infected cell soma, as seen in other types of infections (Holmes, 1975; Panem and Kirsten, 1975). Sometimes groups of aligned buds appear to form synchronously and budding sites are also seen along the processes of stellate cells as in Sindbis and vesicular stomatitis virus-infected cells (Birdwell et aE., 1973; Birdwell
FIG. 9. Replica of the cell surface 8 days after inoculation, showing an elongated process from a stellate cell (the cell body is outside of the field illustrated) lying on the surface of another cell. Numerous protrusions of various sizes mark the surface of the elongated process and these protrusions have on their surfaces the globular units characteristic of viral buds (arrows). Clusters of viral particles are scattered over the rest of the cell surface. Some of these are irregularly shaped and larger than others. Similar clusters of free virus occasionally adhere to the glass, as shown in the inset. These free viruses are smaller than the large viral buds but the globular units on their surfaces are larger so that they appear almost confluent. (Compare inset to Fig. 7.) Origin of platinum shadowing is on the right. x 72,200; inset x 85,500. FIG. 10. Thin section through an elongated process from a stellate cell 8 days after inoculation. These processes are characterized by their content of filaments (F) and the numerous viral buds on their surface (small arrows). Viral particles are smaller then the buds and characterized by a dense core (large arrow). X 71,250.
528
VISNA
VIRUS-ASSOCIATED
and Strauss, 1974). The mechanism by which these events are controlled was not evident in our study. The final step in viral maturation occurs after the virus has been released from the cell, and is characterized by a reduction in size of viral particles. The capsid detaches from the envelope, as if it were uncoiling, and forms a central core. The membrane complexes, assumed to be responsible for capsid attachment, must consequently be broken, allowing greater plasticity of the viral membrane and possibly permitting the loss of envelope membrane as the virus gets smaller. How the membrane is removed from the viral envelope has not been clarified in this study although the existence of intermediate stages between the large crescent buds and the small free viruses was observed. The free viruses are covered with large and closely apposed surface globular units that make a continuous coat on the detached virus. Since the surface globular units are present throughout viral maturation, they constitute a marker for viral structures. They resemble structures described on the surface of various oncornaviruses (de Harven et al., 1973; Demsey and Stackpole, 1975; Nermut et al., 1972; Panem and Kirsten, 1975; Sheffield, 1973, 1974) and presumably correspond to glycoproteins organized as clusters of spikes on the surface of visna virus (Coward et al., 1972; Haase, 1975; Takemoto et al., 1973; Thormar, 1961). The increase in size of the surface units observed during viral formation could be due to continuous addition of viral proteins to the bud surface, in a manner analogous to the widening of surface strands in productive measles infection (Dubois-Dalcq and Reese, 1975). ACKNOWLEDGMENTS The technical help of Kathy Worthington and Frank Nolan is gratefully acknowledged. We are also grateful to Barbara Reese for taking the Scanning electron micrographs, and to Drs. H. Panitch and H. McFarland for reviewing the manuscript. REFERENCES BIRDWELL, C. R., STRAUSS, E. G., and STRAUSS, J. H. (1973). Replication of sindbis virus. III. An electron microscopic study of virus maturation using
MEMBRANE
CHANGES
529
the surface replica technique. Virology 56, 429438. BIRDWELL, C. R., and STRAUSS, J. H. (1974). Maturation of vesicular stomatitis virus: Electron microscopy of surface replicas of infected cells. Virology 59, 587-590. CHIPPAUX-HYPPOLITE, C., TARANGER, C., TAMALET, J., PAUTRAT, G., and BRAHIC, M. (1972). Aspects ultrastructuraux du virus visna en cultures cellulaires. Ann. Inst. Pasteur 123, 409-420. COWARD, J. E., HARTER, D. H., and MORGAN, C. (1970). Electron microscopic observations of visna virus-infected cell cultures. Virology 40, 10301038. COWARD, J. E., HARTER, D. H., Hsu, K. C., and MORGAN, C. (1972). Ferritin-conjugated antibody labeling of visna virus. Virology 50, 925-930. DEHARVEN, E., LAMPEN, N., SATO, T., and FRIEND, C. (1973). Scanning electron microscopy of cells infected with a murine leukemia virus. Virology 51, 240-243. DEMSEY, A., and STACKPOLE, C. W. (19751. Cell surface membrane alterations during murine leukemia virus morphogenesis. J. Cell Biol. 67, 92a. DUBOIS-DALCQ, M., and REESE, T. S. (1975). Structural changes in the membrane of vero cells infected with a paramyxovirus. J. Cell Biol. 67,551565. DUBOIS-DALCQ, M. (1976). In “Workshop on the Application of Modern Biological Research to Investigation of Slow Virus Infections of the CNS.” Wurzburg. (Ter Meulen and Katz, eds.). Springer Verlag, New York/Berlin, In press. FRIEDMANN, A., COWARD, J. E., HARTER, D. H., LIPSET, J. S., and MORGAN, C. (1974). Electron microscopic studies of visna virus ribonucleic acid. J. Gen. Virol. 25, 93-104. HAASE, A. T. (1975). The slow infection caused by visna virus. Cur-r. Top. Microbial. Immunol. 72, 102-156. HEUSER, J. E., LANDIS, S. M. D., and REESE, T. S. (1976). Preservation of rapidly changing structures in synapses by rapid freezing. Cold Spring Harbor Symposium: The Synapse. HOLMES, K. V. (1975). Scanning electron microscopic studies of virus-infected cells. I. Cytopathic effects and maturation of vesicular stomatitis virus in L2 cells. J. Viral. 15, 355-362. MACINTYRE, E. H. WINTERSGILL, C. J., and VATTER, A. E. (1973). Visna virus infection of sheep and human cells in vitro: An ultrastructural study. J. Cell Sci. 13, 1733191. MACINTYRE, E. H., WINTERGILL, C. J., and VATTER, A. E. (1974). Prolonged culture of visna virus in human astrocytes. Beitr. Pathol. 152, 163-178. NATHANSON, N., PANITCH, H., and PETURSSON, G. (19761. Pathogenesis of visna: Review and speculation. In “Slow Virus Diseases of Animals and Man” (R. H. Kimberlin, ed.), pp. 115-131. North
530 Holland
DUBOIS-DALCQ,
REESE
Publishing Co., Amsterdam. O., SILVERSTEIN, A. M., PRICE, D., and R. T. (1974). Visna virus infection of American lambs. Science 183, 1202-1203. NERMUT, M. V., FRANK, H., and SCHAFER, W. (1972). Properties of mouse leukemia viruses. III. EM appearance as revealed after conventional preparation techniques as well as freeze-drying and freeze-etching. Virology 49, 345-358. PANEM, S., and KIRSTEN, W. H. (1975). Secondary scanning electron microscopy of cells infected with murine oncornaviruses. Virology 63, 447-458. SHEFFIELD, J. B. (1973). Envelope of mouse mammary tumor virus studied by freeze-etching and freeze-fracture techniques. J. Viral. 12, 616-624. SHEFFIELD, J. B. (1974). Membrane alterations which accompany MuMTV maturation. I. Studies by freeze-cleave techniques. Virology 57, 287-290. NARAYAN, JOHNSON,
AND
NARAYAN
B., PALSSON, P. A., and GRIMSSON, H. (1957). Visna, a demyelinating transmissible disease of sheep. J. Neuropathal. Exp. Neural. 16, 389-403. TAKEMOTO, K. K., MATTERN, C. F. T., STONE, L. B., COE, J. E., and LAVELLE, G. (19711. Antigenic and morphological similarities of progressive pneumonia virus, a recently isolated “slow virus” of sheep, to visna and Maedi Viruses. J. Virol. 7, 301-308. TAKEMOTO, K. K., AOKI, T., GARON, C., and STURM, M. M. (1973). Comparative studies of visna, progressive pneumonia and Rous sarcoma viruses by electron microscopy. J. Nat. Cancer Inst. 50, 543545. THORMAR, H. (1961). An electron microscope study of tissue cultures infected with visna virus. Virology 14, 463-475.
SIGURD~SON,