American Journal of Transplantation 2003; 3: 1383–1392 Blackwell Munksgaard
C Blackwell Munksgaard 2003 Copyright
ISSN 1600-6135 doi: 10.1046/j.1600-6135.2003.00237.x
BK Polyoma Virus Allograft Nephropathy: Ultrastructural Features from Viral Cell Entry to Lysis Cinthia B. Drachenberga , John C. Papadimitrioua, ∗ , Ravinder Walib , Christopher L. Cubittb,c and Emilio Ramosb Departments of a Pathology and b Medicine, University of Maryland School of Medicine, Baltimore, MD, USA c National Institute of Neurological Disorders and Stroke (NINDS), National Institutes of Health, Bethesda, MD, USA ∗Corresponding author: John C. Papadimitriou,
[email protected] BK virions must enter the host cell and target their genome to the nucleus in order to complete their life cycle. The mechanisms by which the virions accomplish these tasks are not known. In this morphological study we found that BK virions localized beneath the host cell cytoplasmic membrane in 60– 70-nm, smooth (non-coated) monopinocytotic vesicles similar to, or consistent with, caveolae. In the cytoplasm, the monopinocytotic vesicles carrying virions appeared to fuse with a system of smooth, vesicles and tubules that communicated with the rough endoplasmic reticulum and was continuous with the Golgi system. Membrane-bound single virions and large tubuloreticular complexes loaded with virions accumulated in paranuclear locations. Occasional nuclei displayed virions within the perinuclear cisterna in association to the perinuclear viral accumulations. Tubular cells with mature productive infection had large nuclei, distended by daughter virions, whereas they lacked significant numbers of cytoplasmic virions. In addition to virally induced cell necrosis, there was extensive tubular cell damage (apoptosis and necrosis) in morphologically non-infected tubules. The observed ultrastructural interactions between the BK virions and host cells are remarkably similar to viral cell entry and nuclear targeting described for SV40 virus. Key words: Caveolae, cell injury, cytopathic changes, kidney, SV40, transplantation Received 9 February 2003, revised and accepted for publication 20 May 2003
Introduction Two of the 13 known polyomaviruses, BK and JC, are human pathogens causing nephritis and progressive multi-
focal leukoencephalopathy (PML), respectively. In normal individuals, subclinical BK and JC primary infections are associated with seroconversion in more than 90% of cases by the age of 20 (1). JC and/or BK viral reactivation is typically observed in immunosuppressed individuals (2). With the AIDS epidemic, there has been a marked increase in the incidence of progressive multifocal leukoencephalopathy (2). In recent years, an increasing incidence of BK virus nephropathy has been reported in renal transplant patients (3). In general, polyoma viruses are species-specific. The simian polyomavirus (SV40) is endemic in monkeys and in association with immunosuppression causes simian PML (4). SV40 has been reported to co-infect renal transplant recipients presenting with BK allograft nephropathy (5). SV40 has been extensively studied since 1960, when it was first reported. These studies have led to extremely important insights into the molecular biology of mammalian cells and oncogenesis (6). In contrast, very little is known on the pathogenesis of diseases associated with the BK and JC viruses. Similarities between BK, JC and SV40 at the DNA and protein level result in identical morphology. These three viruses are considered to have similar epidemiology and biological behavior in their respective natural hosts as well (1,2). Transmission of infection is most likely by the oral route (1,2). After multiplication at the site of entry, the viruses reach their target organs. The principal target organ for all three viruses is the kidney (4). The primary infection is followed by latency in the urinary tract epithelium, lymphoid cells and central nervous system (1). The polyomaviruses are a family of small, non-enveloped, DNA viruses. The viral capsid is icosahedral and has a diameter of 40–44 nm (7). The genome consists of a closed circular double-stranded DNA molecule with approximately 5 kb, that encodes the early (regulatory) and late (structural) proteins (6). For the life-cycle of the virus to be completed, the virions must attach to the host cell plasma membrane and target their genome to the nucleus. In the cell nucleus, the uncoated mini-chromosome is transcribed. Transcription of the early genes results in the production of the T antigens that cause quiescent cells to re-enter the cell cycle and thus begin replication of cellular DNA. In permissive host cells the T antigens, acting as regulatory proteins, direct the remaining events, resulting in a productive infection (8). The completion of the process consists of viral 1383
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DNA replication and transcription of late genes for the production of the structural proteins (VP1,VP2 and VP3) that will constitute the capsid. Viral capsomeres assemble around the daughter mini-chromosomes in the nucleus, to form stable viral particles (6). SV40 uses an ‘atypical’ endocytic mechanism to enter the host cells. This virus is internalized in non-clathrin-coated vesicles called caveolae. From caveolae the virions reach the nucleus through an intermediate vesicular and tubular system (9–13). This is in contrast to many other viruses that enter the host cells by the usual endocytic pathway. The latter is a constitutive pathway that utilizes clathrin-coated vesicles targeted to the endosomal/lysosomal system. In these organelles the viruses are disassembled through acid-activated hydrolytic enzymes (10,12). Disassembly of SV40, however, is not dependent on low pH (14). Several studies have described the interactions between the murine polyoma virus and the host cells. It appears that this virus follows the same pathway as SV40, using caveolae to enter the host cell (15–18). On the other hand, one study found that the murine polyoma virus entered the host cells using non-clathrin-coated, non-caveolar vesicles (19). For the BK and JC viruses, little or nothing is known about the mechanisms of cellular entry, cytoplasmic transport and nuclear targeting. While there have been no studies specifically addressing these aspects of the BK virus infection, a well-conducted study has shown that JC virions use for cell entry the typical endocytic pathway with clathrincoated vesicles (20). Despite their important similarities at the molecular and morphological level, it appears that the polyoma viruses use divergent pathways for cellular entry. Understanding the interactions between virions and host cells at the ultrastructural and molecular level has important implications in the treatment and prevention of the specific infection. This information may also be important in the context of gene therapy. In the current electron microscopic study we describe the morphological findings in patients with BK allograft nephropathy. Our main goals are to: (a) describe the interactions between the BK virions and the renal tubular epithelial cells, particularly in relationship to viral entry, cytoplasmic trafficking and nuclear targeting; (b) characterize the overall cytopathic changes and the features of cell injury and death in the cells showing nuclear accumulation of viruses; and (c) identify and describe any other additional ultrastructural features directly or indirectly related to the viral infection.
Materials and Methods Renal transplant biopsies from eight patients and transplant nephrectomies from two patients with active BK virus nephropathy were studied ultrastruc-
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turally. The diagnosis was made following previously described criteria (21). By light microscopy, multiple tubular cells showed the typical cytopathic effects of polyoma virus. These have been well described previously (22) and mainly consist of nuclear enlargement with the presence of a basophilic, ‘gelatinous’ nuclear inclusion displacing the chromatin. The diagnosis of BK virus infection was confirmed with urine PCR studies.
PCR amplification Viral DNA was isolated from a low-speed centrifuged pellet of urine (10–15 mL) using the QIAamp Viral RNA/DNA isolation kit (Qiagen, Valencia, CA, #29304). The DNA was resuspended in 50 lL ultrapure H2O and stored at –20 ◦ C. Viral DNA were amplified by PCR using the TaKaRa Ex Taq Kit (Takara Bio Inc., Japan, #RR001A) in a reaction mixture as per manufacturer’s instructions. For detection of BKV DNA 215-bp fragment in VP1 gene of BKV was amplified using primers BLP-15 (5 -ACAGCACAGCAAGAATTCCCCTCCCand BLP-16 (5 -CAAGGGTTCTCCACCTACAGCAA-3 ). Two 3 ) sets of PCR primers were used for detection of SV40 DNA. The STP-1 (5 -CAGGTTCAGGGGGAGGTGTGGG-3 ) and STP-2 (5 GATGGTGGGGAGAAGAACATGG-3 ) amplify a 178-bp region of SV40 T antigen. SLP-1 (5 -TTGATGTGGGAAGCTGTTACTG-3 ) and SLP-4 (5 ATGAAAATTTGACCCTTGAATG-3 ) amplify a 129-bp region of SV40 VP-1. Each primer was used at a final concentration of 2.5 ng/lL. The PCR program consisted of 95 ◦ C, 1 min; 60 ◦ C, 1 min; and 72 ◦ C, 1 min for 40 cycles with an initial denaturation step of 95 ◦ C for 5 min and a final extension of 72 ◦ C for 10 min. The annealing temp was set to 64 ◦ C for the STP-1, -2 primer set. The PCR products were analyzed on a 2% agarose gel containing 0.5% lg/mL ethidium bromide. There was no evidence of concurrent JC or SV40 viral excretion by PCR in any of the patients. For electron microscopy the samples were fixed in a solution of 4% formaldehyde and 1% glutaraldehyde in mono-phosphate buffer, followed by 1% osmium tetroxide in mono-phosphate buffer. The tissue was then dehydrated in increasing concentrations of alcohol, cleared in propylene oxide and embedded in epoxy resin (Epon 812). Ultrathin sections from these blocks were stained with uranyl acetate and lead citrate. Transmission electron microscopy was performed with a JEOL 1200 EX1. A total of 64 grids were evaluated ultrastructurally (3 from each biopsy and 20 from each nephrectomy). An average of 150 tubular cross-sections were present per grid. Serial sections of 7 grids with abundant infected tubules were also evaluated.
Results Virions associated with the surface cytoplasmic membrane In infected tubules abundant virions were present over the cell surfaces of tubular cells (Figure 1a). The virions were roughly spherical with a diameter of 40–42 nm. The distribution of viruses over the cell surfaces was random and not restricted to the apical aspects. The viral load on the surface of individual cells varied from a few scattered isolated virions to thick clumps of innumerable virions. Some virions attached tightly to the cytoplasmic membrane. In viable cells a minority of virions was associated with flask-like invaginations of the membrane as they attached to the cell surface. Other virions were located deeper in the cytoplasm and apparently caused the cytoplasmic membrane to wrap around them. In other cases the virions were within a vesicle that appeared to have been separated from the surface membrane. The vast majority of the vesicular membranes American Journal of Transplantation 2003; 3: 1383–1392
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Figure 1: a: Renal tubular cell with BK virions attaching to the surface cellular membrane, including microvilli. b and c: Entry of BK virions. The virions attach to the surface membrane and induce formation of flask-like invaginations that surround the virus. d: The membrane encloses the virion and a vesicle is formed. e: The viral loaded vesicles separate from the inner aspect of the surface cellular membrane (arrows).
In the vicinity of the cytoplasmic membrane the vesicles carrying virions were more often found separate from each other (Figure 1e). In contrast, in deeper locations (400– 500 nm from the surface), the vesicles with virions were more often found in clusters of up to 100 vesicles (Figure 2). The clusters of vesicles, each vesicle loaded with a single virion, fused with irregularly shaped vesiculartubular structures (Figure 2). These latter structures were morphologically consistent with the descriptions of caveosomes (9,10,12). In rare instances there were recognizable changes in the cellular cytoskeleton with aggregation of intermediate and thin filaments in the adjacent cytosol.
Figure 2: Deeper in the cytoplasm the viral-loaded vesicles fuse with irregular tubulo-vesicular structures. Two virions are seen still on the surface of the cell (top right).
were smooth (i.e. lacking a clathrin-like cytoplasmic coat). Most vesicles were spherical and measured 60 nm in average diameter. The membranes surrounded the virus tightly, although there was a 10–12-nm narrow space between the virion and the membrane (Figures 1e and 2). Progressive stages of viral internalization are demonstrated on Figure 1(a–e). Rare vesicles had irregular shapes or a compound appearance, and rarely enclosed two or three virions. In some cases the latter findings appeared to result from tangential cutting of surface virions, partially enclosed by the cytoplasmic membrane. Extremely rare vesicles (less than 0.5%) appeared to show an external fuzzy layer of electron-dense material, in some cases suggesting a clathrin coat. In cells with evidence of advanced productive infection and accumulation of intranuclear virions, there were no significant numbers of virions in pinocytotic vesicles. American Journal of Transplantation 2003; 3: 1383–1392
Cytoplasmic virions In infected tubules, only a minority of tubular cells displayed cytoplasmic viral particles rather than the nuclear viral aggregates characteristic of the mature productive infection. In cells with predominant cytoplasmic virions, the latter were seen in monopinocytotic vesicles, sometimes near the nucleus, or were localized in complex membranebound viral aggregates. The aggregates were of two types: (I) The virions were localized in irregularly shaped vesicles or were aligned in long tubules with smooth and parallel walls (Figure 3). The vesicles and the tubules were often continuous. The tubulovesicular aggregates were randomly distributed in the cytoplasm, and when the sections demonstrated longitudinal images of the tubules, these were long and ran over significant portions of the cytoplasm (Figure 3). (II) A tubulo-reticular network of delicate undulating tubules that contained large numbers of virions mostly arranged in a lattice pattern (Figure 4). The second type of viral aggregates was more often noted in perinuclear locations (Figure 5). They were composed of a few or innumerable viruses that in some cases formed inclusions as large as the nucleus associated to them (Figure 6). The tubulo-vesicular 1385
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Figure 3: The virions accumulate in a complex system of smooth-walled tubulo-vesicular structures. The virions approximate the nucleus presumably through this tubular system. Accordingly, membrane-bound virions are seen by the nucleus (arrows).
Figure 5: An aggregate of virions within tubulo-vesicular structures is seen by the nucleus. Intact perinuclear envelope is seen to the left (arrowheads). Adjacent to the viral aggregate the membranes of the perinuclear envelope become distorted and fuse with the peripheral membranes of the viral aggregate. The contents of the latter appear to be in continuity with the perinuclear cisterna (arrow). Another cluster of tubules loaded with virions is seen in the top right area.
Figure 4: Large numbers of viruses accumulate in complex tubulo-reticular structures that tend to be close to the nucleus. These viral aggregates closely associate with the Golgi complex (G) and are continuous with the rough endoplasmic reticulum (asterisks).
aggregates were seen occasionally in continuity with the RER and in close proximity and/or continuity to the Golgi system (Figure 4). The cells with abundant virions attached on the external aspect of the cell membranes, only rarely contained abundant tubulovesicular or tubuloreticular cytoplasmic aggregates. The converse was also true, suggesting that these represented two stages (viral entry and trafficking, respectively) in the host cell. The membrane-bound viruses in the cytosol were morphologically identical to the extracellular ones, showing no morphological evidence of disassembly. 1386
Figure 6: Large, ‘inactive’-appearing paranuclear viral aggregates (asterisks). In contrast to the changes seen in Figure 5, in this case there is no clear evidence of interaction between the viral aggregate and the perinuclear envelope.
In viable cells demonstrating active endocytosis or trafficking of virions, all virions were membrane bound with no instances of virions lying free in the cytosol. This was in contrast to findings in dying cells (see below). Virions and the nucleus Viral aggregates or vesicles loaded with virions were commonly seen in the vicinity of the nucleus. In many cells American Journal of Transplantation 2003; 3: 1383–1392
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Figure 8: Several virions are seen in the perinuclear cisterna (arrows). The perinuclear cytoplasm contains an accumulation of excess membranes (arrowheads) and tubulo-reticular structures devoid of virions (asterisk). There is prominent rarefaction of the chromatin with clumping and formation of thick granules. Insert: Part of another nucleus contains a loose aggregate of partially membrane-bound virions associated with microtubules (asterisks). A nuclear pore is clearly seen (arrowhead) without any association to the virions.
Figure 7: (a,b,c) a: Membrane-bound virions are located close to the nucleus (arrow) and within the perinuclear cisterna (arrowhead). A non-membrane-bound virion is also seen in the nucleus (open arrow). In the vast majority of cases the virions found within the nucleus were membrane bound (b and c).
the viral aggregates were closely associated to the perinuclear cisternae, and in occasional nuclei the viral particles were seen within the perinuclear cisterna (Figure 7). In these cases, it was common to see apparent fusion of the membranes surrounding the virions and the perinuclear membranes (Figure 5). In association with the perinuclear viral aggregates, there were excess membranes and, rarely, aggregates of empty membranous tubulo-reticular aggregates (Figure 8). The viral particles in the perinuclear cisterna and inside the nucleus proper did not associate specifically with the nuclear pores, although in rare cases the virions were located in the vicinity of nuclear pore areas. In some nuclei, close to the perinuclear viral aggregates there were small aggregates of virions loosely surrounded by membranes, suggesting that this appearance could have resulted from tangential cutting of virions located in the perinuclear cisterna and herniating into the nucleus (Figure 7). Occasional nuclei contained fibrillar or microtubular arrays in association with the viral particles (Figure 8). Rarefaction of the chromatin with thick granules and aggregates was seen in cells with evidence of perinuclear viral activity. Occasional cells showed formation of small non-membrane-bound viral aggregates, presumably of daughter viruses. In numerous cases, despite the presence of very abundant paranuclear viral aggregates, there American Journal of Transplantation 2003; 3: 1383–1392
was no evidence of nuclear entry or interaction between the virions and the nuclear membrane (Figure 6). In those cells it appeared that the process of nuclear entry had been arrested, resulting in extensive accumulation of perinuclear virions in cells that appeared viable and otherwise healthy (Figure 6). Lysis of infected tubular cells In tubular cells with mature productive infection the nuclei were markedly enlarged due to the accumulation of daughter viral particles in loose groups or in dense crystalline arrays (Figure 9). In many cases the viral aggregates were separated from the nuclear membrane by a rim of chromatin (Figure 9). Numerous cells with abundant nuclear accumulation of virions showed various stages of cell injury and, in the most severe cases, necrosis. The latter was characterized by nuclear and cytoplasmic swelling, associated with generalized disruption of the cell membranes, including rupture of the nuclear envelope (Figure 9). In cells undergoing necrosis, there were large cytoplasmic aggregates of randomly distributed, non-membrane-bound virions. These often attached to fragments of membranes and could be observed occasionally in lysosomes (Figure 10). In occasional infected cells undergoing dissolution, the nuclear viral aggregates appeared very dense, with loss of morphological detail of the viral particles; these nuclei appeared mummified. Lysis of the infected cells resulted in massive shedding of virions into the tubular lumen associated with the formation of casts. Accumulation of viruses was also seen in the intercellular spaces. In heavily infected tubules, most cell 1387
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fection showed that virions reach the extracellular space only by cell lysis. Viral particles remain confined within the nuclear membrane until the latter ruptures. In the stage of cell lysis, there is generalized evidence of loss of integrity of the cellular membranes. The tubular cells appeared to become infected by direct contact with infected cells undergoing lysis. Therefore, in infected tubules the vast majority of cells contained viruses. In contrast, unaffected tubules usually did not contain any virions at all. Tubular cell necrosis and destruction of the tubular basement membranes occasionally resulted in large extracellular aggregates of virions. The latter were rarely seen within lysosomes in interstitial macrophages. Intact basement membranes appeared to preclude penetration of virions in adjacent (non-infected) tubules.
Figure 9: Infected cell undergoing lysis. A dense aggregate of progeny virions is seen in the nucleus (top left). The perinuclear membranes are disrupted. In the cytoplasm there are marked degenerative changes (extensive membrane vesiculation, mitochondrial swelling and disruption). Insert: Progeny virions in nucleus. The virions display a crystalline array and are surrounded by rim of chromatin that separates the bulk of virions from the nuclear membranes.
Figure 10: In necrotic areas the virions were randomly distributed in the cytoplasmic fragments, usually not membrane bound (arrows), although often attached to membranes. A crystallized cluster of virions is seen within a structure consistent with a lysosome (bottom center).
surfaces of viable and dying cells were covered by virions. Viral particles were also present between the infected cells and the underlying tubular basement membrane. Evaluation of more than a thousand tubular cells containing the typical nuclear features of a fully developed productive in1388
Vascular and interstitial changes In two of the ten cases, the endothelial cells and occasional epithelial cells showed abundant tubulo-reticular ‘viral-like’ inclusions of the type described in patients treated with interferon, HIV-associated nephropathy or lupus nephritis (23,24) (Figure 11). Virions, however, were not seen in endothelial cells in any instance. Prominent accumulation of inflammatory cells was seen in the interstitium. The inflammation mostly consisted of lymphocytes, macrophages and plasma cells (Figure 12).
Figure 11: Endothelial cell with a large tubulo-reticular inclusion. This does not contain virions and is identical to the viral-like aggregates seen in endothelium of HIV-infected patients. Numerous caveolae (a normal organelle of endothelial cells) are seen in all aspects of the endothelial plasma membrane (arrows). Insert: Surface cytoplasmic membrane of an epithelial cell covered by virions. In the cytoplasm a virion is located within a vesicle showing a fuzzy lining typical of a clathrin coat, an extremely rare occurrence (0.5%).
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stood infection, several recent studies have shed light on the clinical aspects of BK allograft nephritis (25,26). Specific functional and molecular studies on the biological behavior of BK virus at the cellular level are not available, and very little is known about the interactions between the virus and the host cells. In contrast, numerous studies have characterized the cellular interactions of the SV40 virus and murine polyoma virus (9–20). For the purpose of further understanding the pathogenicity of BK virus, it may be helpful to draw potential analogies with these better known polyoma viruses, particularly the SV40.
Figure 12: Tubular cross-section with obvious cellular damage (consistent with necrosis and apoptosis). There is extensive fragmentation of membranes. Condensed and swollen mitochondria are seen (asterisk). On both sides of the tubule the interstitium shows abundant inflammatory cells and cellular debris.
Admixed with the inflammation were cellular debris and deposition of immature collagen fibers. Tubular cell injury in non-infected tubules More than 50% of tubules in each biopsy lacked any virions. However, despite the absence of clear evidence of infection, there was evidence of generalized tubular cell injury. The tubular cells showed loss of nuclear polarity, cytoplasmic vacuolization or condensation, and loss of brush border. Numerous tubules were lined by irregularly shaped, overlapping epithelial cells rather than by a single layer of orderly columnar cells. Tubulitis (lymphocytic tubular inflammation) was seen in scattered infected and non-infected tubules (Figure 12). Scattered tubular cells showed necrosis, apoptosis or intermediate features between necrosis and apoptosis. Cells undergoing necrosis were swollen and showed extensive disruption of the cell membranes. Apoptotic cells showed condensation of the chromatin at the nuclear periphery in a crescentic fashion and cytoplasmic condensation. In most cells the subcellular organelles, in particular the mitochondria, showed a spectrum of changes from severe condensation to high amplitude swelling and terminal disruption. The endoplasmic reticulum and Golgi showed dissolution into small vesicles. In occasional cells the cytoskeleton collapsed, particularly at the cell base. Vesiculation and blebbing of the cell plasma membrane was more prominent at the luminal surface, but also occasionally towards the cell base.
Discussion BK virus nephritis is an important cause of renal allograft dysfunction and graft loss. Although still a poorly underAmerican Journal of Transplantation 2003; 3: 1383–1392
A large variety of compounds and organisms enter cells through the process of endocytosis (internalization of external particles or molecules). The type of vesicle involved in the process of cell entry and its coat, determines to a large extent the type of cargo that will be transported and its destination within the cell (27). First described in endothelial cells (see Figure 11), caveolae are small (50– 70 nm) flask-shaped vesicles present in the cell surface of many cell types (28). Caveolae are rich in lipids and contain caveolin-1. They are actively involved in cholesterol cell entry (10). It has recently been recognized that SV40 enters the host cell through caveolae (10–13). Ebola virus among other viruses also uses caveolae (29). In the case of SV40, the virions first attach to a receptor related to MHC class I molecules and then translocate to specialized membrane microdomains (lipid rafts), where the virions induce the formation of caveolae and therefore their own internalization (10–13). In this study we have found that BK virions appear to enter the renal tubular cell in smooth (non-coated) monopinocytotic vesicles that are morphologically consistent with caveolae. The morphological findings are remarkably similar to SV40 and murine polyoma virus host cell entry. This is in contrast to the manner of entry of JC virus, that has been found to enter cells by following the usual endocytic pathway using clathrin-coated vesicles (20). The different pathways of viral cell entry do not appear to directly correlate with the type of receptor, since BK, JC and murine polyoma viruses have sialic-acid type receptors, whereas SV40 has receptors related to the MHC system (30). It has been previously reported for SV40 that entry of virions in the host cell is restricted to the apical aspects of the cell (31). We have not, however, observed that the pinocytotic vesicles were restricted to the apical surface of the tubular cells. In the case of SV40, caveolae loaded with SV40 virions fuse with a tubulo-vesicular system of pre-existing organelles that have neutral luminal pH and also contain caveolin1 (10). These organelles, called caveosomes, start fusing with each other and undergo rapid shape changes, including formation of long tubular structures (10). These tubules have no attached ribosomes, but they are in continuity with the rough endoplasmic reticulum (32). The extensive network of tubules and vesicles that transport the virions from one part of the cell to another is also consistent with recently described cisternal compartments within 1389
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intracellular transport pathways that include branching tubular elements that can be several micrometers long (33). Specifically regarding SV40 virions, it has been demonstrated that, while avoiding lysosomal degradation, they take advantage of a retrograde endocytic pathway that leads to the RER and from there to the nucleus. This track retrieves, in usual circumstances, resident proteins that have escaped from the endoplasmic reticulum to the Golgi system (12). Based on our morphological findings, it is likely that BK virus uses similar mechanisms as SV40 to reach the endoplasmic reticulum, and later the nucleus. We observed that the monopinocytotic vesicles loaded with BK virions aggregated with each other and fused with polymorphous membranous organelles connected to long smooth tubular and tubuloreticular structures that appeared to carry the virions towards the paranuclear areas. We observed that the paranuclear viral aggregates were occasionally continuous with the perinuclear cisterna, the rough endoplasmic reticulum and the Golgi system. Similar findings have been described also for murine polyoma virus (15). The mechanism of nuclear entry is not clearly understood for any of the polyoma viruses. Some studies have suggested that nuclear entry was impaired by inhibitors of the nuclear pore complex system (10,34). Others believe, however, that nuclear pores are too small to allow the entry of intact virions (16). We did not see evidence of viral entry through nuclear pores, nor do we believe pore size could be a limiting factor for viral entry (see Figure 8 insert). Interactions between the perinuclear, membrane-bound, SV40 viral aggregates and the perinuclear cisterna have been described in several studies. It has been shown that fragments of labeled surface cytoplasmic membrane are carried with the SV40 virions and accumulate in the outer membrane of the nuclear cisterna (17,35). Also, several studies have confirmed that during the process of viral DNA nuclear entry, there is fusion of the membranes surrounding the virions with the outer nuclear membrane (36– 38). Similar to studies of SV40 and murine polyoma virus (17,19,34), we consistently saw BK virions in the perinuclear cisterna as well as evidence of membrane fusion between the perinuclear viral aggregates and the nuclear membranes. Interestingly, the numbers of virions in the perinuclear cisterna and within the nucleus were extremely sparse, in contrast to the large paranuclear aggregates of cytoplasmic virions. The viral minichromosome has to be uncoated before it can be replicated in the nucleus. It is, however, not known if uncoating occurs before or after entrance in the nucleus. Some authors believe that rapid virus uncoating occurs after nuclear entry (17,39). One study showed that intact viruses were not seen in the nucleus a few hours after infection, and the authors concluded that intranuclear uncoating is very efficient. However, if uncoating occurs within the nucleus, the question of how the progeny viruses resist disassembly is raised. It has been proposed 1390
that once replication starts there may be an inhibition of the nuclear uncoating enzymes, or that the progeny virus may have a protein coat different from the infecting particles (36). On the other hand, more recent studies have suggested that uncoating of polyoma viruses occurs in the vicinity of the nucleus or as the virions enter the nucleus rather than in the nucleus itself. Regarding murine polyoma virus, it has been demonstrated that VP1 protein which composes the bulk of the viral capside does not enter the nucleus (16). Another study has shown that VP2 and VP3 proteins, which link the capside to the minichromosome, accumulate in the perinuclear tubules carrying SV40. This finding supports the idea that disassembly starts within the tubular system, before the viral genetic material enters the nucleus (12). In the current study, the sparsity of intact intranuclear virions in association with the perinuclear viral aggregates appears to support the concept that viral uncoating occurs before nuclear entry of the DNA particle. In the case of SV40, it has been proposed that in cells with productive infection and accumulation of large number of nuclear viral particles, a fibrillary network associated with the nuclear pores prevents the virions from exiting towards the cytoplasm (40). It has been reported that the bulk of SV40 virions remains associated to the nuclear matrix until virus-induced cell lysis occurs (41). This is in concordance with the fact that non-enveloped virus, such as the polyoma viruses, are released from the infected cells by cell lysis (42). Some studies have shown that small amounts of SV40 virions reach the extracellular medium before cell lysis occurs (41,43). Evaluation of a very large number of infected cells in the current study indicated that the vast majority of intranuclear daughter virions remain within the nucleus until cell death by lysis occurs. Cells showing rupture of the nuclear membrane (lysis) often contained nonmembrane-bound virions in the cytoplasm, or occasionally in the perinuclear cisterna. Extensive cell necrosis characterizes infection with SV40 (44,45). Similarly, BK virus allograft nephropathy is characterized by extensive tubular necrosis (22). Surprisingly, in this study we observed that, in addition to necrosis of infected cells, there was generalized damage of apparently non-infected tubular cells. Widespread tubular cell injury and death appeared in the form of both necrosis and apoptosis. Several studies have shown that the SV40 virus has complex effects on the cell cycle of the host cell. The large T antigen has both apoptotic and anti-apoptotic effects, resulting from binding to the retinoblastoma tumor suppression proteins and to p53. In addition, the small t antigen inhibits the apoptosis-inducing effects of the large T antigen (46). The extensive damage in tubular cells that lack evidence of viral infection probably represents the background of tubular cell injury on which viral reactivation occurs. In clinical studies, tubular injury secondary to immunosuppressant American Journal of Transplantation 2003; 3: 1383–1392
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drug toxicity and immune-mediated damage (rejection) has been associated with BK virus nephropathy (21,26). 10.
It has recently been proposed that quantitative viral load in plasma correlates with the degree of BK virus nephritis (47,48). This notion is supported by the findings in the current study. We have observed that in tubules undergoing massive virally induced tubular cell necrosis, there is dissolution of basement membranes, with massive spillage of virions into the intertubular space and destruction of intertubular capillary walls. Through the latter, the virions would easily gain access to the systemic blood flow. Currently, there is no effective treatment for the BK virus infection. Understanding of the mechanisms of cell entry, cytoplasmic trafficking and nuclear targeting has important potential implications for the prevention and treatment of BK virus-associated nephropathy (49,50).
11.
12.
13.
14. 15.
16.
The conclusions derived from the current study are purely observational. From the morphological standpoint, however, it appears that BK and SV40 polyoma viruses follow a similar endocytic pathway for nuclear targeting (11,12). To a significant extent, similar findings have also been described for murine polyoma virus (15). Additional functional and molecular studies are necessary to further understand the interaction between BK and the permissive host cell.
17.
18.
19.
Acknowledgments We wish to thank Caroline Ryschkewitsch for her help in the performance of the molecular studies. We are also grateful to Perry Comegys for excellent photographic work. We wish to acknowledge the late Dr Gerald Stoner who was consulted on multiple occasions during the preparation of this work and who was a continuous source of inspiration for us.
References 1. Dorries ¨ K. New aspects in the pathogenesis of polyomavirusinduced disease. Adv Virus Res 1997; 48: 205–261. 2. Dorries ¨ K. Molecular biology and pathogenesis of human polyomavirus infections. Dev Biol Stand 1998; 94: 71–79. 3. Hirsch HH. Polyomavirus BK nephropathy: a (re-) emerging complication in renal transplantation. Am J Transplant 2002; 2: 25–30. 4. Shah KV. SV40 infections in simians and humans. Dev Biol Stand 1998; 94: 9–12. 5. Li RM, Mannon RB, Kleiner D et al. BK virus and SV40 co-infection in polyomavirus nephropathy. Transplantation 2002; 74: 1497– 1504. 6. Fanning E. Introduction to simian virus 40: getting by with more than a little help from its host cell. Dev Biol Stand 1998; 94: 3–8. 7. Howley PM. Molecular biology of SV40 and the human polyomaviruses BK and JC. In:Klein G, ed. Viral Oncology. NY: Raven Press; 1980. pp. 489–550. 8. Hershey EL. Simian virus 40–host cell interactions during lytic infection. J Virol 1979; 30: 76–83. 9. Pullmans L, Kartenbeck J, Helenius A. Caveolar endocytosis of
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20.
21.
22.
23. 24.
25.
26.
27.
28. 29.
30.
simian virus 40 reveals a new two-step vesicular-transport pathway to the ER. Nat Cell Biol 2001; 3: 473–483. Pullmans L, Puntener ¨ D, Helenius A. Local actin polymerization and dynamin recruitment in SV40-induced internalization of caveolae. Science 2002; 296: 535–539. Parton RG, Lindsay M. Exploitation of major histocompatibility complex class I molecules and caveolae by simian virus 40. Immunol Rev 1999; 168: 23–31. Norkin LC, Anderson HA, Wolfrom SA, Oppenheim A. Caveolar endocytosis of simian virus 40 is followed by Brefeldin A-sensitive transport to the endoplasmic reticulum, where the virus disassembles. J Virol 2002; 76: 5256–5266. Chen Y, Norkin LC. Extracellular simian virus 40 transmits a signal that promotes virus enclosure within caveolae. Exp Cell Res 1999; 246: 83–90. Upcroft P. Simian virus 40 infection is not mediated by lysosomal activation. J Gen Virol 1987; 68: 2477–2480. Richterova´ Z, Liebl D, Horak ´ M et al. Caveolae are involved in the trafficking of mouse polyomavirus virions and artificial VP1 pseudocapsids toward cell nuclei. J Virol 2001; 75: 10880–10891. Griffith GR, Marriott SJ, Rintoul DA, Consigli RA. Early events in polyomavirus infection: fusion of monopinocytotic vesicles containing virions with mouse kidney cell nuclei. Virus Res 1988; 10: 41–52. Griffith GR, Consigli RA. Isolation and characterization of monopinocytotic vesicles containing polyomavirus from the cytoplasm of infected mouse kidney cells. J Virol 1984; 50: 77–85. Mattern CF, Takemoto KK, Daniel WA. Replication of polyoma virus in mouse embryo cells: electron microscopic observations. Virology 1966; 30: 242–256. Gilbert J, Benjamin TL. Early steps of polyomavirus entry into cells. J Virol 2000; 74: 8582–8588. Pho MT, Ashok A, Atwood WJ. JC virus enters human glial cells by clathrin-dependent receptor-mediated endocytosis. J Virol 2000; 74: 2288–2292. Drachenberg RC, Drachenberg CB, Papadimitriou JC et al. Morphological spectrum of polyoma virus disease in renal allografts: diagnostic accuracy of urine cytology. Am J Transplant 2001; 1: 378–381. Nickeleit V, Hirsch HH, Binet IF et al. Polyoma virus infection of renal allograft recipients: from latent infection to the disease. J Am Soc Nephrol 1999; 10: 1080–1089. Rich SA. Human lupus inclusions and interferon. Science 1981; 213: 772–775. Kostianovsky M, Orenstein JM, Schaff Z, Grimley PM. Cytomembranous inclusions observed in acquired immunodeficiency syndrome. Arch Pathol Lab Med 1987; 111: 218–223. Ramos E, Drachenberg CB, Papadimitriou JC. Clinical course of polyoma virus nephropathy in 67 renal transplant patients. J Am Soc Nephrol 2002; 13: 2145–2151. Hirsch HH, Knowles W, Dickenmann M et al. Prospective study of polyomavirus type BK replication and nephropathy in renal transplant recipients. N Engl J Med 2002; 347: 488–496. Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P. Intracellular vesicular traffic. In: Molecular Biology of the Cell, 4th edn. New York, NY: Garland Science; 2003. p. 711. Bruns RR, Palade GE. Studies on blood capillaries. J Cell Biol 1968; 37: 244–276. Empig CJ, Goldsmith MA. Association of the caveola vesicular system with cellular entry by filoviruses. J Virol 2002; 76: 5266– 5270. Keppler OT, Stehling P, Herrman M et al. Biosynthetic modulation of sialic acid-dependent virus receptor interactions of
1391
Drachenberg et al.
31.
32.
33.
34. 35.
36.
37.
38.
39.
40.
two primate polyoma viruses. J Biol Chem 1995; 270: 1308– 1314. Clayson ET, Compans RW. Entry of SV40 is restricted to apical surfaces of polarized epithelial cells. Mol Cell Biol 1988; 8: 3391– 3396. Kartenbeck J, Stukenbrok H, Helenius A. Endocytosis of simian virus 40 into the endoplasmic reticulum. J Cell Biol 1989; 109: 2721–2729. Hopkins CR, Gibson A, Shipman M, Miller K. Movement of internalized ligand-receptor complexes along a continuous endosomal reticulum. Nature 1990; 346: 335–339. Yamada M, Kasamatsu H. Role of nuclear pore complex in Simian Virus 40 nuclear targeting. J Virol 1993; 61: 119–130. Nishimura T, Kawai N, Ichihara I. Interaction of endocytic vacuoles with the inner nuclear membrane in simian virus 40 into CV-1 cell nucleus. Cell Struct Funct 1991; 16: 441–445. Hummeler K, Tomassini N, Sokol F. Morphological aspects of the uptake of simian virus 40 by permissive cells. J Virol 1970; 6: 87–93. Maul GG. Fibrils attached to the nuclear pore prevent egress of SV40 particles from the infected nucleus. J Cell Biol 1976; 70: 714–719. Maul GG, Rovera G, Vorbrodt A, Abramczuk J. Membrane fusion as a mechanism of simian virus 40 entry into different cellular compartments. J Virol 1978; 28: 936–944. Mackay RL, Consigli RA. Early events in polyoma virus infection: attachment, penetration and nuclear entry. J Virol 1976; 19: 620– 636. Maul GG. Fibrils attached to the nuclear pore present egress of SV40 particles from the infected nucleus. J Cell Biol 1976; 70: 714–719.
1392
41. Schirmbeck R, von der Weth A, Deppert W. Structural requirements for simian virus 40 replication and virion maturation. J Virol 1993; 67: 894–901. 42. Clayson ET, Jones Brando LV, Compans RW. Release of simian virus 40 virions from epithelial cells is polarized and occurs without cell lysis. J Virol 1989; 63: 2278–2288. 43. Gwaltney JM, Jr. Rhinoviruses. Yale J Biol Med 1975; 48: 17– 45. 44. Eggleton KH, Norkin LC. Cell killing by simian virus 40: the sequence of ultrastructural alterations leading to cellular degeneration and death. Virology 1981; 110: 73–86. 45. Norkin LC. Cell killing by simian virus 40: impairment of membrane formation and function. J Virol 1977; 21: 872–879. 46. Hay S, Kannourakis G. A time to kill. viral manipulation of the cell death program. J Gen Virol 2002; 83: 1547–1564. 47. Hirsch HH, Knowles W, Dickermann M et al. Prospective study of polyomavirus type BK replication and nephropathy in renal transplant recipients. N Engl J Med 2002; 347: 527– 536. 48. Ramos E, Drachenberg CB, Portocarrero M et al. BK virus nephropathy diagnosis and treatment: experience at the University of Maryland Renal Transplant Program.Cecka and Terasaki, ed. Clinical Transplants 2002. Los Angeles, CA: UCLA Immunogenetics Center; 2003: 143–153. 49. Keppler OT, Stehling P, Herrmann M et al. Biosynthetic modulation of sialic acid-dependent virus–receptor interactions of two primate polyoma viruses. J Biol Chem 1995; 270: 1308– 1314. 50. Pietropaolo V, Degener AM, Di Taranto C, Sinibaldi L, Orsi N. The in vitro effect of monensin on BK polyomavirus replication. New Microbiol 1995; 18: 341–349.
American Journal of Transplantation 2003; 3: 1383–1392
