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Address correspondence to Thomas J. Hope, Dept. of Microbiology and Immunology, MSB E-704, M/C 790, 835 South Wolcott Ave., Chicago, IL 60612. Tel.: (312) 413-3424. Fax: (312) 996-6415. E-mail: thope{at}uic.edu
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Abstract |
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Key Words: HIV-1; reverse transcription complex; dynein; fluorescent microscopy; electron microscopy
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Introduction |
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HIV infects nondividing cells by delivering its genome into the nucleus through nuclear pores. To reach the pores in the nuclear envelope, the genome must travel from the plasma membrane through the cytoplasm. The integration competent particle, called the preintegration complex, has been estimated to be at least 50 nm in diameter (Miller et al., 1997). Because of the high viscosity of the cytoplasm, movement of these particles by diffusion is likely to be very limited (for review see Luby-Phelps, 2000). Some viruses overcome this obstacle by hijacking cytoplasmic motors to utilize the cellular cytoskeleton as a roadway. For instance, herpes simplex virus (HSV)-1 (Sodeik et al., 1997) and adenovirus (Ad; Suomalainen et al., 1999) are thought to use dynein motors to travel along the microtubule network for intracellular transport. To characterize the intracellular trafficking of HIV, we have developed several fluorescence-based methods that allow the detection and characterization of individual intracellular complexes of viral origin. Importantly, several of these labeling methods allow the visualization of individual virions in living cells. Observation of intracellular HIV in living cells revealed that HIV moves in the cytoplasm in curvilinear paths. Further analysis revealed that intracellular HIV is associated with microtubules and uses cytoplasmic dynein to move toward the nucleus of the cell. These studies suggest that the intracellular trafficking of the HIV genome is a highly ordered process using cellular motor pathways.
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Results |
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We generated labeled replication competent virus stocks by cotransfection of HIV proviral DNA with a plasmid that expresses GFP fused to the NH2 terminus of HIV-1 Vpr (GFPVpr). After target cells were exposed to the virus-containing supernatants, point sources of GFP fluorescence were found associated with the infected cells, presumably marking the presence of the GFPVpr-tagged intracellular particles (Fig. 1). Adherent human cells expressing CD4 were used in these studies because their flat morphology provides an extended cytoplasm, allowing greater resolution of cellular compartments. Early in infection, the GFP signal appeared as punctate signals spread throughout the cell (Fig. 1 A), and when the target cells were washed of free virus and allowed to incubate further, a significant proportion of the signal accumulated in the perinuclear region, often at the microtubule-organizing center (MTOC), in as little as 2 h (Fig. 1 B).
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HIV uses the cellular cytoskeleton to move inside of cells
HIV particle behavior in living cells was analyzed next. Target cells expressing CD4 and coreceptor were infected with GFPVpr-labeled HIV for 30 min, and the unbound virus was washed away (Fig. 3). In this experiment, the M-trophic HIV strain YU-2 and target cells expressing the appropriate CCR5 coreceptor were used. The red stain identifies mitochondria to visualize general cellular structure. Virus location was monitored through a z-series of images taken every 5 min. The paths of movement of several particles during the 95 min of observation is shown in Fig. 3 A. Individual frames for the movement of a single particle is shown in Fig. 3 B. The particle moves in a curvilinear path toward the region of the nucleus (videos 1 and 2 available at http://www.jcb.org/cgi/content/full/jcb.200203150/DC1). To quantify the migration of HIV within cells, we determined the distances of individual particles from the nearest edge of the cell and from the nuclear membrane. We reasoned that a fractional representation of the distances (in this case the distance to the cell edge over the sum of the distances) should give an accurate measure of relative nuclear migration, so that particles at the periphery have positional values near 0 and particles close to the nucleus have values near 1 (Fig. 3 B). This allows the movement toward the nucleus to be quantified even though the particles have different distances to travel because the virus can attach anywhere on the exposed surface of the cell and each cell has a distinct morphology, which can change during the course of the time-lapse experiment. To further define particle movement, Ghost cells were used. Ghost cells contain a GFP reporter for HIV infection and express CD4 and CXCR4 coreceptor. Background levels of GFP expression allow the target cells to be simultaneously observed with the GFP-labeled HIV. Ghost cells were infected for 20 min at a low multiplicity of infection (MOI) with GFPVpr-labeled HIV and then imaged as a z-series every 5 min after washing to determine particle motility over time. The graphical representations of the migration of five typical particles (Fig. 3 C) shows both inward and outward motion over time, but in four of the five examples shown, significant nuclear migration occurred over the course of the experiment, and the particles tended to remain near the nucleus once they arrived there. Of 19 particles from 13 different cells examined in this way, 14 (74%) particles were in the perinuclear region at the end of the 75 min time course (unpublished data).
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To address the possibility that HIV might be tracking along microtubules, we performed time-lapse experiments with GFPVpr-tagged HIV-infected cells containing microinjected, fluorescently labeled tubulin (Fig. 4; see videos 1 and 2). For this study, virus with DiD-labeled membranes was used to allow the identification of particles, which had lost their membranes as would be expected after functional entry into the cytoplasm. The labeled HIV was spinoculated onto the target cells to increase the number of bound virions (O'Doherty et al., 2000), and the cells were washed and observed by fluorescent microscopy. Images were captured every minute for a 14-min period. Several DiD-negative particles are observed to migrate during the time course. Although the microtubule network is dense because of the injected tubulin, the tracking of DiD-negative particles along microtubules is apparent. Two examples of movement along microtubules are shown in Fig. 4 B. The particle designated by the white arrow is likely the same throughout the panel; however, the particle moved out of the plane of imaging for a single frame so that it is possible that the movement of two different DiD-negative particles is shown (see videos 1 and 2). In addition, the movement of DiD-negative particles not associated with microtubules is also observed (Fig. 4 B, yellow arrow).
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Discussion |
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An important consideration in efforts to observe cytoplasmic complexes containing the HIV genome is the ability to distinguish productive from nonproductive binding events. HIV is considered to be relatively inefficient at infecting cells, with infectivity to particle ratios reported to be as low as 1 in 60,000 (Kimpton and Emerman, 1992; Piatak et al., 1993). However, more appropriate measures of infectivity, that is the fraction of particles which bind to the cell that cause an infectious event, suggest a ratio up to 1 in 10 for retroviruses (Andreadis et al., 2000). Even at this level of infectivity, it was necessary for us to distinguish between productive and nonproductive entry. For this reason, we focused on two types of GFP-tagged particles: those that had lost a membrane dye, suggesting they had fused with the cellular membrane, and those which had incorporated deoxynucleotides. In both cases, these particles represent a subset of HIV that had entered the cytoplasm of the target cells to form viral complexes which are competent to complete the infectious cycle.
Our time-lapse studies of internalized viral particles suggested both microtubule-dependent and microtubule-independent cytoplasmic movement. Since disruption of both actin and microtubule was necessary to completely halt motility the microtubule-independent movement may use actin. Previous studies have found that disrupting the actin network inhibits infection with HIV, and it was proposed that actin-based motility is important either for cytoplasmic entry (Iyengar et al., 1998) or for establishment of an infectious viral complex (Bukrinskaya et al., 1998). However, further study is required to determine if the microtubule-independent movement is directed through interaction with the actin cytoskeleton or by some other mechanism.
Role for cytoplasmic dynein in intracellular motility of HIV
Our studies suggest that HIV RTCs use the microtubule network for long range movement within cells. Most of the GFPVpr-labeled particles were found to be associated with microtubules. Over 80% (16 of 19) of the GFPVpr-labeled complexes that were also labeled with fluorescent nucleotides were associated with microtubules in Triton X-100extracted cells (unpublished data). We documented movement of GFPVpr-labeled HIV along fluorescently labeled microtubules using time-lapse microscopy. Further, preliminary analysis suggests that the DiD-negative, GFP-labeled particles can move at burst velocities on the order of 1 µ/s, a rate consistent with that known for microtubule based motors (Presley et al., 1998). We visually demonstrated that GFPVpr-labeled complexes extracted from infected cells with saponin could bind to fluorescently labeled microtubules generated in vitro (unpublished data). Finally, attachments between the RTC and microtubules were detected ultrastructurally using correlative EM. Association with microtubules was apparently not cell type specific because the interaction was detected in Hos, HeLa, and primary fibroblast cells. The likely function of the microtubule-based motility is to transport the HIV genome from the cell periphery to the nucleus.
Other viruses use dynein-mediated transport along microtubules to gain access to the nucleus. To determine the role of dynein in the movement of HIV particles along microtubules, we inhibited its motor function by microinjection of an inhibitory monoclonal antibody. After infection, GFPVpr-labeled particles defined as entering the cytoplasm either by the loss of membrane label or by the incorporation of fluorescent nucleotides remained in the periphery of injected cells relative to adjacent uninjected cells. In cells with inhibited dynein function, the particles were clustered at points most distant from the nucleus of the infected cell. Preliminary studies suggest that the mislocalization was due to a reversal of the dominant, microtubule minus enddirected movement seen in uninjected cells (unpublished data). Studies of Ad-2 mobility in living cells revealed a similar pattern when dynein function was inhibited by overexpression of p50 dynamitin (Suomalainen et al., 1999). The Ad-2 mislocalization is thought to be a consequence of plus end microtubule activity associated with the motor complex used by this virus. Additional evidence for a plus enddirected activity lies in the observation that Ad-2 oscillates on the microtubules between short range plus end and minus end movements, very similar to the type of motility we have observed with HIV. This oscillation is not seen with Ad-5, and disruption of dynein in that case results in cessation of movement, not with plus enddirected motion (Leopold et al., 2000). Our studies suggest that HIV uses a motor complex with similarities to the one used by Ad-2.
It has been reported previously that disruption of the microtubule network by nocodazole treatment inhibits HIV infection by approximately twofold (Bukrinskaya et al., 1998). We have obtained similar results in this type of study (unpublished data). Nocodazole treatment during HSV and Ad-2 infection has also been shown to decrease the efficiency but not prevent infection (Sodeik et al., 1997; Suomalainen et al., 1999). Several possibilities have been proposed to explain this incomplete inhibition, including entry proximal to the nucleus, transport along nocodazole-resistant microtubules, or movement by Brownian motion (Mabit et al., 2002).
Initiation of reverse transcription in cytoplasmic HIV containing p24CA
There has been much speculation about the cytoplasmic fate of the conical capsid that contains the HIV genome inside virions. In some models, the capsid dissolves immediately after membrane fusion. Others propose that the capsid remains intact until the genome reaches a nuclear pore. The conical core is a relatively unstable complex that is sensitive to all but the most mild detergent treatments. Recently, methods have been developed which allow the isolation of large amounts of HIV cores from virions (Kotov et al., 1999; Accola et al., 2000; Wilk et al., 2001), although intact cores have not been isolated from infected cells. Our in situ analyses identified two distinct species of GFPVpr-labeled complexes associated with reverse transcriptase activity, one lacking p24CA and the other containing significant amounts of p24CA, suggesting that the capsid is intact in the latter complexes. Structural studies of the HIV capsid have suggested the presence of pores large enough to allow entry of nucleotides into the capsid (Li et al., 2000). Further, the ability to facilitate reverse transcription within intact virions by exposure to high concentrations of deoxynucleotides along with polyanions also suggests that reverse transcription within intact capsids is possible (Zhang et al., 1996). Previous biochemical studies have not identified significant amounts of p24CA associated with the RTC. However the conditions typically used to generate lysates for RTC and PIC purification may confound this analysis because the HIV capsid easily disassembles in the presence of detergents or possibly in the low salt conditions of hypotonic lysis (Ganser et al., 1999; Kotov et al., 1999; Accola et al., 2000; Wilk et al., 2001). This study provides the first evidence of reverse transcription within an intact HIV capsid in the cytoplasm of an infected cell, suggesting that the capsid remains intact for at least a portion of the process from the initiation of reverse transcription to maturation of the PIC. Furthermore, the p24CA containing RTCs are associated with microtubules (Fig. 9), suggesting that this association with the microtubule network can occur before loss of the capsid. Alternatively, the p24CA-associated RTC may reflect dead end events due to a failure of capsid dissolution after fusion.
Ultrastructure of cytoplasmic RTCs
To gain insight into the morphology of the cytoplasmic RTC, we adapted a correlative electron microscopic technique used previously to study the cytoskeleton. This method uses the alignment of fluorescent and platinum replicas of detergent extracted cells. Unfortunately, this method only allows visualization a short distance into the cell, and therefore, only RTC in the periphery could be imaged. Using EM, we find structures overlapping with the fluorescently visualized RTC. They are typically cylindrical in shape with varying length and a very similar diameter. Previous biophysical characterization of cytoplasmic HIV RTC derived from acutely infected cells suggests that the composition of the RTC is dynamic over time (Fassati and Goff, 2001); while the density of these complexes remains constant, the sedimentation velocity of complexes containing HIV proviral DNA increased at later time points. This observation suggests that proteinDNA complexes with a range of sizes and/or shapes are found in the cytoplasm at different times during reverse transcription. The large size of the RTCs identified in the periphery by correlative EM is most consistent with the early, larger complexes detected in the previous studies. Alternatively, the detergent treatment of the infected cells required for our method may lead to an increase in the size of complexes detected. In either case, the structures we propose to be RTCs are consistent with previous biophysical descriptions. Further study, including immunogold staining for viral proteins and analysis of complexes generated after entry mediated by the HIV envelope will clarify if the observed structures are bona fide cytoplasmic RTCs. In most cases the RTC cylinder is overlapping a microtubule or connected to a microtubule by a stalk-like projection. The observed connections between the identified complexes and microtubules is consistent with our model that intracellular HIV complexes are using the microtubule network to move within the cytoplasm.
Our observations of the composition and trafficking of intracellular HIV complexes suggests the following scenario. After entering the cytoplasm, the HIV genome uses some aspect of the actin cytoskeleton to move within the peripheral regions of the cell. This is consistent with evidence that actin can be used to gain access to the microtubule network (Taunton, 2001). It is also supported by previous reports that an intact actin cytoskeleton is necessary for efficient infectivity (Bukrinskaya et al., 1998). The infectious viral particle must next make its way to the microtubule network, where it can initiate reverse transcription even before losing the majority of its capsid protein. At some point after interaction with the microtubule network, the conical capsid dissociates, yet the RTC maintains its interaction with microtubules. This interaction is likely mediated by tethering with a cellular motor complex which has both minus end and plus enddirected motor activities, as is proposed for Ad-2. Minus enddirected movement of the RTC along the microtubule network ultimately leads to the microtubule organizing center, very near the nuclear membrane, where the mature RTC can then enter through nuclear pore complexes in order to integrate into the host DNA.
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Materials and methods |
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Virus production
GFPVpr-labeled HIV is produced by CaPO4 transfection of 293T cells with the proviral constructs pLAI or pLAI-Yu (Yu2 env in the pLAI backbone) (Vodicka et al., 1998) and the plasmid peGFPC3 (CLONTECH Laboratories, Inc.) containing the entire Vpr-coding region fused to the COOH terminus of eGFP (GFPVpr). Cells are washed at 1620 h posttransfection, medium is replaced at 36 h, and supernatants containing labeled virus is collected at 24-h intervals for two or three harvests. For pseudotyped HIV, an env-deleted pLAI provirus, VSV-G expression plasmid, and the GFPVpr plasmid are cotransfected. Virus is collected as above and concentrated by ultracentrifugation for 2 h at 23,000 rpm through a 20% sucrose pad and resuspended in PBS at 50100-fold concentration (Bartz and Vodicka, 1997). Purification of virus by gradient centrifugation was performed using an Optiprep (Nycomed) density gradient as described (Dettenhofer and Yu, 1999). DiD (DiIC18; Molecular Probes) labeling was achieved by addition of DiD (10 µM) after the initial wash, rinsing away unincorporated dye the next morning, and collecting supernatants as above. All virus preparations were assayed for infectivity using MAGI indicator cells, and the GFPVpr incorporation was assessed by p24CA staining of virions bound to glass using mAb AG3.0.
Immunofluorescence
Cells are grown on glass coverslips, rinsed with PBS, and fixed with 3.7% formaldehyde (Polysciences) in 0.1 M Pipes, pH 6.8. Antibodies are added at predetermined dilutions in SB (PBS, 10% normal donkey serum [Jackson ImmunoResearch Laboratories], 0.1% Triton X-100) for 20 min at RT. Coverslips are rinsed extensively and stained with labeled (AMCA-, Cy3- or Cy5-) donkey antimouse antibodies (Jackson ImmunoResearch Laboratories) in SB. Coverslips are then mounted onto glass slides with Gel Mount (Biomedia) containing an antifade reagent. Dried slides are imaged on an Olympus IX70 epifluorescent microscope fitted with an automated stage (Applied Precision Inc.), and images are captured in z-series on a CCD digital camera. Out of focus light is digitally removed using the Softworks deconvolution software (Applied Precision Inc.). Live cell video microscopy is performed using a heated open or closed cell chamber along with an objective heater (Bioptechs Inc.), and cells are maintained on coverslips in medium supplemented with 50 mM Hepes, pH 7.5.
Quantification of nuclear migration
Particle location is determined by measuring the distances to the nearest edge of both the nucleus and the cell periphery using the Deltavision Softworks program and is expressed as the fraction of the distance to the edge divided by the total distance so that particles at the periphery have values approaching 0 and nuclear particles approaching 1. For Figs. 3 and 5, the image z-series were first projected as a single image using the three-dimensional reconstruction feature of the software. As a result, it is not possible to measure particles directly over the nucleus so these particles were excluded from the analysis. For Fig. 7, RTCs that are over the nucleus are generally obscured by the strong nuclear AlexadUTP signal.
Correlative EM
Cytoskeletal samples are prepared as described (Svitkina and Borisy, 1998) with modifications. Briefly, primary fibroblast cells are grown on glass coverslips patterned by evaporation of gold onto an EM locator grid. Cells are injected with Alexa-594dUTP (Molecular Probes), infected with GFPVpr-labeled, VSV-G pseudotyped HIV at high titer (MOI 50100), and incubated for 4 h at 37°C. Coverslips are then extracted with 1% Triton X-100, 4% PEG mol wt 40 kD (SERVA Biochemicals), and 10 mM taxol (Sigma-Aldrich) in PEM buffer (80 mM Pipes, pH 6.8, 1 mM EGTA, 1 mM MgCl2). Actin is depleted by incubation in G' buffer containing taxol and gelsolin (3 mg/ml; provided by S. Choe, The Salk Institute) at RT for 1 h. Cells are rinsed and fixed with 2% glutaraldehyde and immunostained with anti
-tubulin and Cy5 donkey antimouse secondary. Injected cells are imaged on an epifluorescent microscope, and the locations of candidate cells are noted with respect to the grid pattern. Samples are prepared for EM analysis by desiccation in ethanol and dried in a critical point dryer. They are then coated with platinum in a rotary vacuum evaporator followed by carbon coating to maintain structure. Candidate cells are mounted on formvar-coated EM grids and examined on a JEOL 1220 electron microscope. The unique microtubule structure is used to digitally align the immunofluorescent and EM images using Adobe Photoshop® software.
Online supplemental material
Figs. 3 and 4 time-lapse images are compiled into Quicktime videos available at http://www.jcb.org/cgi/content/full/jcb.200203150/DC1. Video 1 shows the movement of HIV particles in living cells depicted in Fig. 3. HeLa/CD4/CCR5 cells were infected with GFPVpr-labeled HIV (green), stained with MitoTracker (red), and mounted in a live cell chamber for observation at 37°C. 14 0.5-µm optical z-sections were acquired every 5 mn for 95 min (20 frames) and rendered in single three-dimensional volume views. The video is displayed at 1 s/frame. Individual HIV particles appear as several spots in some cases due to short rapid movement during acquisition of the z-series. Video 2 shows microtubule-dependent and -independent movement of HIV particles in living cells as depicted in Fig. 4. Hos/CD4 cells were microinjected with rhodamine-tubulin and incubated for 1 h at 37°C to label microtubules (blue), spinfected with GFPVpr (green), and DiD (red)-labeled HIV for 1 h at 1,200 g, 23°C. The cells were washed and placed in medium supplemented with 50 mM Hepes and 0.1 µM taxol. Images were collected every minute for 14 min (15 frames) in the three color spectra at 37°C. Video 1 shows the full frame images displayed at 1 s/frame, showing the movement of many particles near microtubules and in areas devoid of microtubules. Video 2 is a close-up of a region, showing microtubule-dependent (white arrow) and -independent (yellow arrow) movement.
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Footnotes |
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* Abbreviations used in this paper: Ad, adenovirus; GFPVpr, GFP fused to the NH2 terminus of HIV-1 Vpr; HIV, human immunodeficiency virus; HSV, herpes simplex virus; MOI, multiplicity of infection; MTOC, microtubule-organizing center; PIC, HIV-1 preintegration complex; RTC, reverse transcription complex; VSV-G, vesicular stomatitis virus envelope glycoprotein.
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Acknowledgments |
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This work was supported by National Institutes of Health R01 grant no. AI47770-04 to T. Hope and a grant from the University of Washington Center for AIDS Research to M. Vodicka.
Submitted: 29 March 2002
Revised: 27 September 2002
Accepted: 27 September 2002
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References |
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