Viruses are parasites that must infect the living cell of a host in order to grow and replicate. To do so, the virus attaches to the host’s cell and transfers its genetic material to the inside. The virus then hijacks the cell and forces it to build proteins and more genetic material that will assemble into new copies of the virus, and the completed virus particles are released to infect new cells.

The HIV-1 virus encodes a protein called Gag that coordinates the assembly of new copies of this virus. Thousands of Gag proteins accumulate into a lattice at the inside of the host cell membrane by an unknown mechanism, where they gather all essential components to form a new infectious virus. Once completed, the virus particle detaches itself from the host cell and Gag gives the virus its structure.

Previous research by several groups has shown that a lipid molecule found in cell membranes called PIP₂ can bind to Gag and helps the virus to assemble. However, until now, it was unclear if PIP₂ anchors Gag to the cell membrane of living cells and if it plays any other roles in the later stages of assembly.

Now, Mücksch et al. have developed a new approach to study this question by rapidly manipulating the levels of PIP₂ in living cells and monitoring Gag assembly through a fluorescent marker. The experiments showed that PIP₂ was needed to initiate the assembly process, but also to maintain the partially assembled Gag lattice at the membrane. When PIP₂ was removed, Gag proteins could not gather at the cell membrane and the already assembled Gag lattices broke down. When PIP₂ levels were increased again, the Gag proteins that had disappeared from the membrane formed new lattices. This suggests that PIP₂ has a much broader role in HIV-1 particle formation than previously assumed.

A next step will be to use this new experimental approach to study how particles of HIV are assembled and released. This knowledge may help scientists to develop antiviral drugs that interfere with the assembly step of the virus that could be used to prevent or treat HIV infections.

Human immunodeficiency virus type 1 (HIV-1) particles assemble and bud at the plasma membrane (PM) of infected cells. This requires trafficking of the main structural component Gag from its site of synthesis at cytosolic polysomes to the inner leaflet of the PM. Here, Gag assembles into a multimeric lattice comprising ~2500 Gag molecules. Gag also recruits the other constituents of the infectious virion as well as components of the cellular endosomal sorting complex required for transport (ESCRT) to nascent budding sites (reviewed in Sundquist and Kräusslich, 2012).

Gag consists of the individually folded MA (matrix), CA (capsid), NC (nucleocapsid) and p6 domains, which separate upon proteolytic maturation to the infectious virion. CA and NC mediate protein and RNA interactions during virion morphogenesis, while p6 recruits ESCRT components. PM targeting of Gag depends on the N-terminal MA domain. Mutational analyses revealed that the N-terminal myristoyl moiety and a surface exposed patch of basic residues in MA (the highly basic region, HBR) are important for Gag targeting, membrane association and virus formation (Lorizate and Kräusslich, 2011; Freed, 2015). However, these two features would not be sufficient to explain specific PM targeting. The phosphoinositide phosphatidylinositol 4,5-bisphosphate (PI(4,5)P₂) is known to act as a cue for specific recruitment of many peripheral PM proteins and was also shown to be important for Gag PM targeting. Depleting PI(4,5)P₂ abolished PM targeting of Gag, while increasing PI(4,5)P₂ at intracellular membranes redirected Gag to those sites (Ono et al., 2004). Consistently, several studies reported that presence of PI(4,5)P₂ enhances binding of Gag-derived proteins to synthetic liposomes in vitro (Chukkapalli et al., 2008, 2010; Dick et al., 2012; Olety and Ono, 2014; Mercredi et al., 2016), and PI(4,5)P₂ appeared to be enriched in the HIV-1 lipidome (Chan et al., 2008).

Gag seems to dominate in a monomeric form or in small oligomers in the cytoplasm (Kutluay and Bieniasz, 2010; Hendrix et al., 2015), and its N-terminal myristate group is proposed to be buried within the globular domain of MA at this stage (Spearman et al., 1997; Saad et al., 2006; Tang et al., 2004). Gag interaction with PI(4,5)P₂ has been suggested to trigger exposure of the myristate moiety followed by its insertion into the inner leaflet of the PM, thereby anchoring Gag. Recent models propose a regulatory role of RNA for this myristoyl switch (Chukkapalli et al., 2010; Alfadhli et al., 2011; Chukkapalli et al., 2013; Kutluay et al., 2014). In the cytosol, specific cellular tRNAs are bound to the basic region of MA in the absence of PI(4,5)P₂, thus preventing premature Gag association with intracellular membranes. This MA-tRNA interaction may be outcompeted by PI(4,5)P₂ at the PM, thereby triggering the myristoyl switch.

The NMR structure of HIV-1 MA bound to a PI(4,5)P₂ molecule carrying truncated acyl chains revealed the 2´ acyl chain to be buried in the myristoyl-binding pocket of MA (Saad et al., 2006). This observation suggested a model for Gag membrane anchoring, which was subsequently challenged by a recent report from the same group, however (Mercredi et al., 2016). Upon arrival of Gag at the PM, the 2´ acyl chain of PI(4,5)P₂ may be flipped outward and displace myristate from the acyl binding pocket of MA. Myristate in turn would insert into the PM. The consequent exchange of the unsaturated 2´ acyl chain of PI(4,5)P₂ by the saturated myristic acid would increase local lipid saturation and membrane order. If occurring in all or a majority of the ~2500 tightly packed Gag molecules, this mechanism could also explain the observed liquid-ordered state of the HIV envelope (Lorizate et al., 2009; Brügger et al., 2006; Chan et al., 2008). This model would further suggest that Gag clusters are stably anchored to the PM via their myristoyl moiety and PI(4,5)P₂ would then be expected to be dispensable once anchoring has occurred.

PI(4,5)P₂ is clearly important for Gag PM targeting, but the dynamics of PI(4,5)P₂ Gag interaction and the role of PI(4,5)P₂ in later stages of HIV assembly have not been examined so far. To do this, we need to monitor Gag localization in real time while manipulating PI(4,5)P₂ levels in living cells. Here, we made use of a recently developed reversible chemical dimerizer system (abbreviated rCDS) allowing rapid and controlled depletion and reconstitution of PM PI(4,5)P₂ levels in living, virus-producing cells by a small molecule (Feng et al., 2014; Schifferer et al., 2015). Using this system, we showed that the nascent HIV-1 Gag assembly site is highly dependent on PI(4,5)P₂ during the entire assembly process, and membrane association of apparently complete assembly sites remained PI(4,5)P₂ dependent. PI(4,5)P₂ removal from the PM not only abolished Gag PM targeting, but also caused dissociation of pre-assembled Gag clusters from the membrane. Reconstitution of bona fide Gag assembly sites at the PM was observed upon re-establishment of PM PI(4,5)P₂ levels. Our findings are inconsistent with stable anchoring of large Gag clusters at the PM through myristoyl moieties alone and suggest a highly dynamic mode of Gag PM binding. Furthermore, this approach allows synchronization of assembly and release of infectious HIV-1. These processes are usually highly asynchronous, both within a cell population and on the level of individual cells (Ivanchenko et al., 2009).

The key parts of the reversible chemical dimerizer system (rCDS; Figure 1A) used in this study are a PM anchor (LCK-ECFP-SNAP, here referred to as ‘Anchor’), and a cytosolic enzyme-bearing construct (FKBP-mRFP-5Ptase, here referred to as ‘Enzyme’), which catalyzes the conversion of PI(4,5)P₂ to PI(4)P (Figure 1A) (Schifferer et al., 2015). Turnover of PM PI(4,5)P₂ is induced by addition of a cell-permeant small chemical dimerizer (rCD1) (Feng et al., 2014), which binds to both the SNAP and FKBP moieties of the respective fusion proteins and thereby links Anchor and Enzyme. Accordingly, rCD1 addition leads to rapid PM recruitment of the Enzyme and consequent depletion of PI(4,5)P₂ from the PM, while PI(4,5)P₂ levels at the PM remain unchanged in the absence of this compound. Under our conditions, PM recruitment of the phosphatase was complete within 5 min after rCD1 addition. PI(4,5)P₂ depletion can be monitored through dissociation from the PM of an EGFP-tagged pleckstrin homology (PH) domain of PLCδ; its dissociation was observed as early as 2 min after rCD1 addition in HeLa cells (Figure 1—figure supplement 1A, Video 1). PM recruitment of the Enzyme can be reversed by adding the rCD1 competitor FK506 (Figure 1A). FK506 outcompetes the binding of rCD1 to the FKBP domain causing removal of the Enzyme from the PM within seconds and replenishment of normal PI(4,5)P₂ levels by endogenous PI(4)P₅ kinases. Full re-localization of the PH-domain reporter to the PM was detected 2 min after addition of FK506, indicating rapid and efficient recovery of PI(4,5)P₂ (Figure 1—figure supplement 1A, Video 1).

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To analyze the relevance of PI(4,5)P₂ for HIV-1 assembly site formation in transfected or infected cells, rCD1-mediated PI(4,5)P₂ depletion needs to be sustained for long periods (>12 hr), while maintaining the potential of rapid reversal and without unwanted toxic effects. These prerequisites were evaluated in HeLa cells co-transfected with plasmids expressing the rCDS components and the PI(4,5)P₂ indicator EGFP-PLCδ-PH. rCD1 was added 4 hr after transfection. Monitoring localization of the PI(4,5)P₂ reporter over time revealed complete loss of PM recruitment up to 22 hr post transfection without apparent cytotoxicity (Figure 1—figure supplement 1B, left). PM PI(4,5)P₂ levels were rapidly restored upon addition of FK506 at this time point (Figure 1—figure supplement 1B). We further observed that Gag assembly was not significantly affected by co-expression of Anchor and Enzyme (p=0.3714) (Figure 1—figure supplement 1C and D), thus validating that the rCDS is applicable to study Gag recruitment and assembly in living cells.

Synchronized induction of Gag assembly site formation upon PI(4,5)P₂ reconstitution

We then asked whether HIV-1 assembly site formation from cytosolic Gag molecules can be induced at the PM by restoring PI(4,5)P₂ levels. Similar to the previous experiments, PI(4,5)P₂ was depleted from the PM in rCDS expressing HeLa cells by rCD1 addition starting 4 hr after transfection. At 22 hr after transfection, Gag-EGFP was found to be mainly cytosolic with very few Gag assemblies at the PM (Figure 2A, left panel). Addition of FK506 and consequent restoration of PM PI(4,5)P₂ levels rapidly induced Gag cluster formation at the PM (Figure 2A and C and Video 2), whereas no major changes were observed upon addition of DMSO (Figure 2B and C, Video 3). After 90 min of FK506 treatment, the ventral PMs were covered with nascent HIV assembly sites and/or particles trapped between the cell and the coverslip (Figure 2A).

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Analogous experiments performed with the MA deletion variant Δ8–126 SR confirmed dependence of the PI(4,5)P₂ responsive phenotype on the MA domain of Gag. Assembly site formation at the PM was not impaired for this variant in PM PI(4,5)P₂ depleted cells, and no significant difference was observed between cells subsequently treated with FK506 or DMSO (Figure 2—figure supplement 1). Deletion of the globular MA domain thus makes Gag independent of and unresponsive to PI(4,5)P₂.

In order to determine whether the rate of Gag assembly differed between induced and native Gag assembly sites, we applied Total Internal Reflection Fluorescence (TIRF) microscopy, which had been used to track single nascent HIV assembly sites over time with low background and high time resolution (Jouvenet et al., 2008; Ivanchenko et al., 2009). Cells were co-transfected with the rCDS components and pCHIV derivatives and either left untreated (Video 4), or subjected to PI(4,5)P₂ depletion at 4 hr after transfection followed by rescue of PI(4,5)P₂ PM levels by FK506 addition at 22 hr after transfection (Video 5). Fluorescence intensity was recorded over time for individual Gag assembly sites. Figure 3A and B show normalized and averaged HIV-1 assembly traces for 175 native and 186 induced assembly sites from three different cells each. No obvious difference in assembly behavior was apparent from these data sets. For a quantitative comparison, data from each individual trace were fitted to a single exponential equation to extract the assembly rate constant k (Figure 3C). Mean assembly rate constants derived from averaging all individual k values did not differ significantly between induced and native assembly, with k = 4.5 ± 0.47*10⁻³ s⁻¹ for native and k = 3.49 ± 0.26*10⁻³ s⁻¹ for induced assembly sites (p=0.8563) and a similar distribution of rate constants (Figure 3—figure supplement 1A). This translates into half-times of approximately 150 s and 200 s for native and induced assembly, respectively, or 8.5 min and 11 min for 90% completion of native and induced assembly, respectively. These values are in agreement with previously published data (Ivanchenko et al., 2009).

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Visual inspection of the movies suggested that PM accumulation of assembly sites occurred more rapidly for induced compared to native assembly sites, while assembly rates at individual sites did not significantly differ between the two conditions. We therefore determined the number of native and induced PM assembly sites over time. Figure 3D shows that assembly sites accumulated faster and more synchronously when induced by PI(4,5)P₂ restoration compared to the native situation. Formation of 100 new assembly sites per 1000 µm² PM required only ~15 min for induced versus 45 min for native assembly.

Live cell TIRF imaging yielded assembly rates, but did not provide information on the morphology of induced assemblies, since the size of HIV-1 particles is below the diffraction limit of conventional light microscopy. Previous studies using PALM/dSTORM super resolution microscopy had revealed a distinct HIV-1 assembly site architecture, with compact Gag clusters of 100–120 nm diameter surrounded by larger patches of Env glycoprotein molecules recruited to the budding site in a Gag dependent manner (Lehmann et al., 2011; Muranyi et al., 2013). In order to obtain insight into the morphology of induced assembly sites, we applied Stimulated Emission Depletion (STED) super resolution microscopy to analyze the distribution of Gag and Env at the ventral membrane of virus expressing cells with or without expression and induction of the rCDS. To allow for detection by STED nanoscopy, we employed Gag tagged with the stainable protein tag CLIP (Gautier et al., 2008; Hanne et al., 2016) instead of the EGFP tag, while Env was detected by indirect immunofluorescence. Imaging at the ventral PM does not distinguish late virus budding structures from trapped extracellular particles, and we therefore refer to late HIV-1 assemblies here.

As can be seen in Figure 4, STED nanoscopy strongly increased resolution compared to confocal images and HIV-1 assembly subdomains became discernible (compare confocal and STED panels). STED images revealed that neither the overall appearance of the cell nor the spatial distribution of Gag and Env at late assemblies differed notably between native and induced assembly. Both structures were characterized by a densely-packed Gag cluster with a diameter of about 120 nm, surrounded by a clustered accumulation of Env (Figure 4), in line with previous observations (Muranyi et al., 2013; Roy et al., 2013). The morphology of induced late assemblies was indistinguishable from native sites in micrographs (compare Figure 4A and B) as well as in averaged line profiles of individual assemblies (compare Figure 4C and D).

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While the previous experiments established that induced Gag assemblies appeared indistinguishable from their native counterparts, the recovery of infectious virus would be ultimate proof for synchronized induction of functional HIV-1 assembly. To this end, we established a stable HeLa_rCDS cell line constitutively expressing Anchor and Enzyme tagged with autofluorescent proteins. The presence of both components of the rCDS in all cells prior to transfection with the HIV-1 expressing plasmid prevents constitutive production of infectious virus from cells only transfected with the HIV-1 plasmid, but lacking one or both components of the rCDS, while an uneven distribution of components between cells is expected to occur when the three plasmids are cotransfected. As shown in Figure 5A, HeLa_rCDS cells constitutively express both fusion proteins, with the Enzyme detected in the cytoplasm and the Anchor at the PM. Next, we assessed functionality of the rCDS in these cells by expressing EGFP-PLCδ-PH, whose PM localization serves as indicator for PI(4,5)P₂. In the absence of rCD1, the indicator was detected mainly at the PM (Figure 5B, left panel). Upon rCD1 addition (Figure 5B, middle panel), the Enzyme translocated to the PM (see insert), resulting in relocalization of the indicator protein to the cytoplasm as a consequence of PI(4,5)P₂ depletion from the PM. This effect was reversed upon addition of FK506 (Figure 5B, right panel).

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After validation of the stable cell line, constitutive or induced virus release and formation of infectious HIV-1 was analyzed following transfection of HeLa_rCDS cells with the complete proviral plasmid pNL4-3. rCD1 was added 4 hr after transfection. Comparing virus production from rCD1 and control cells at 22 hr after transfection showed a strong inhibition by PI(4,5)P₂ depletion (80–85% reduction compared to control cells) (Figure 5C; see also Figure 5—figure supplement 1A for non-normalized data). Virus assembly was then induced in rCD1-treated cells by addition of 1 µM FK506 at 22 hr after transfection. Extracellular virus was collected 2 hr after addition of FK506 since a short harvest time limits the analysis mainly to virions formed by induced assembly. Virus production was restored after FK506 addition (Figure 5—figure supplement 1A). Most importantly, relative infectivity of virions produced under inducing conditions was only mildly reduced (~65% compared to virions produced under native conditions from DMSO treated HeLa_rCDS cells) (Figure 5D; see also Figure 5—figure supplement 1B and C for non-normalized data). These results indicate that induced PM recruitment of Gag allows faithful assembly of infectious HIV-1 particles.

PI(4,5)P₂ is required to retain the partially assembled Gag lattice at the PM

Our results so far confirmed the essential role of PI(4,5)P₂ for Gag recruitment to the PM and established an inducible system for assembly of infectious HIV-1. To determine whether PI(4,5)P₂ is also required to retain multimeric Gag assemblies at the PM, we performed reversible depletion of PM PI(4,5)P₂ in cells displaying pre-formed HIV-1 assembly sites. Assuming insertion of the N-terminally attached myristic acid into the inner membrane leaflet for most or all Gag molecules in nascent assembly sites, PI(4,5)P₂ would not be expected to play a role at this stage. Rapid and reversible PM PI(4,5)P₂ depletion in living cells offers a unique opportunity to address this issue.

HIV-1 expressing HeLa cells carrying the rCDS were incubated without compound addition until a large number of Gag assemblies had accumulated at the PM. Gag assemblies at the ventral PM (towards the coverslip) comprise nascent assembly sites as well as extracellular particles trapped between the cell and the substrate. These extracellular particles can obviously not be affected by rCD1 mediated PI(4,5)P₂ PM depletion, and microscopy of the ventral PM is therefore not suitable for such an experiment. Instead, a central cell section was imaged in the subsequent experiments to focus on nascent assembly sites still connected to the cytosol.

PI(4,5)P₂ depletion was induced in cells with numerous Gag assembly sites by addition of rCD1. Unexpectedly, the vast majority of cells (52 of 59) lost most pre-formed Gag assemblies from the lateral PM upon PI(4,5)P₂ depletion (Figure 6A and Video 6). For quantitative analysis, we determined the relative number of Gag clusters over time following PI(4,5)P₂ depletion in the cell shown in Figure 6A (Figure 6C). This number decreased to 50% at ~30 min after rCD1 addition and reached a minimum level of ~30%. Averaging the relative amount of Gag clusters detected over time in n = 10 cells yielded a very similar result (Figure 6E). In contrast, DMSO treated cells retained or slowly increased the number of Gag assembly sites at the lateral PM over time (Figure 6C and E, Figure 6—figure supplement 1A, Video 7), excluding any unspecific effect of the carrier or the imaging procedure. PM PI(4,5)P₂ depletion also had no effect on preformed assembly sites for the Δ8–126 SR Gag variant (Figure 6—figure supplement 1B and C), and this was confirmed by quantitative analysis of Gag(Δ8–126 SR) cluster number over time (Figure 6—figure supplement 1D).

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The number of Gag assembly sites at the plasma membrane at a given time reflects the equilibrium between assembly site formation and loss of assembly sites, either by extracellular release of budded virions or by dissociation of pre-formed assembly structures from the PM into the cytosol. To assess the contribution of virus release to the observed reduction of PM assemblies following PI(4,5)P₂ depletion, we performed the depletion experiment with a Vpu-defective HIV-1 derivative. Vpu counteracts the host restriction factor tetherin (Figure 6—figure supplement 2A and B). In the absence of Vpu, tetherin retains fully budded HIV-1 particles at the PM and thereby inhibits virus release (Neil et al., 2008; Van Damme et al., 2008). High amounts of PM tetherin in cells producing Vpu(-) particles should therefore largely prevent virus release, while removal of assembly sites from the PM to the cytosol would be unaffected. Control experiments confirmed that neither the presence of the rCDS nor its activation affected the localization of tetherin (Figure 6—figure supplement 2C and D). Importantly, the loss of Gag clusters from the PM upon PI(4,5)P₂ depletion was unaffected by lack of Vpu (Figure 6—figure supplement 2E and F), supporting the conclusion that Gag from nascent assembly sites re-localizes to the cytosol upon PI(4,5)P₂ depletion.

The observed reduction of Gag clusters at the PM upon PI(4,5)P₂ depletion could also be caused by constitutive loss of Gag from the PM, which is normally overcome by PM binding of newly synthesized Gag molecules. In this case, one would expect a reduction of preformed assembly sites when blocking synthesis of new proteins by addition of cycloheximide (CHX). Adding CHX to cells with established Gag clusters did not affect the appearance or number of Gag assemblies over up to 90 min (Figure 6—figure supplement 2G and H), suggesting that loss of Gag assemblies from the cell membrane is caused by PI(4,5)P₂ dependence of their PM association.

HIV-1 Gag assembly sites can be reconstituted by rescue of PM PI(4,5)P₂

We next asked whether Gag molecules that had been dissociated from PM assembly sites by PI(4,5)P₂ depletion could form new assembly sites upon restoration of PM PI(4,5)P₂. For this experiment, we treated cells displaying pre-formed assembly sites with rCD1 for 90 min as described above, followed by addition of FK506 to allow PM PI(4,5)P₂ reconstitution. FK506 treatment yielded a rapid increase in the number of Gag assembly sites at the PM (Figure 6B, Video 6). The number of Gag clusters reached the initial level (prior to rCD1 mediated PI(4,5)P₂ depletion) at 30–40 min after addition of FK506 (Figure 6D and F). No effect on the number of Gag clusters was observed in control cells treated with DMSO (Figure 6—figure supplement 1A, Figure 6D and F, Video 7) or when the PI(4,5)P₂ independent MA variant Δ8–126 SR was used (Figure 6—figure supplement 1B, C and E). Visual inspection of movie sequences indicated that newly induced assembly sites generally did not appear at those positions where previously dissociated sites had been observed.

Quantitative analyses indicated that accumulation of reinduced HIV-1 Gag assemblies following a cycle of PI(4,5)P₂ depletion and restoration (Figure 6D and F) occurred twice as fast as initial accumulation of budding sites under induced conditions (compare Figure 2C). To analyze whether reinduced Gag assembly sites at the PM upon PI(4,5)P₂ reconstitution comprised only newly synthesized Gag molecules or whether ‘recycled’ molecules previously depleted from the membrane contribute, we blocked protein translation by cycloheximide (CHX) treatment 60 min prior to rCD1 addition. As shown in Figure 7A and C, PI(4,5)P₂ depletion resulted in loss of clusters from the PM that was fully reverted by restoring PI(4,5)P₂ levels in the presence of CHX (Figure 7B and D). These data indicate that Gag molecules synthesized before assembly site dissociation are sufficient for formation of new assembly sites following PI(4,5)P₂ reconstitution and new protein synthesis is not required.

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Based on these observations, we asked whether the more rapid accumulation of assembly sites upon dissociation and reinduction may be explained by multimeric Gag lattice remnants following assembly site dissociation. To address this issue, we employed a Gag variant lacking the N-terminal myristoylation site (Gag G2A). This variant cannot assemble at the PM (Bryant and Ratner, 1990; Göttlinger et al., 1989), but can be rescued by a low amount of co-expressed wild-type Gag (Park and Morrow, 1992; Morikawa et al., 1996). Accordingly, co-expression of Gag-EGFP(G2A) and wild-type Gag (in 2:1 molar ratio) led to formation of fluorescent assembly sites at the PM (Figure 6—figure supplement 3A) of cells expressing the Anchor and Enzyme, while Gag-EGFP(G2A) alone failed to assemble. As expected, addition of rCD1 resulted in a loss of Gag-EGFP(G2A) clusters from the PM (Figure 6—figure supplement 3A and C). However, only few Gag-EGFP(G2A) clusters reformed at the PM and the quantitative analysis revealed, that Gag-EGFP(G2A) was not efficiently re-recruited to the PM upon PI(4,5)P₂ rescue (Figure 6—figure supplement 3B and D), while wild-type Gag readily formed new assembly sites under these conditions (Figure 6). This result is consistent with complete dissociation of Gag clusters following PI(4,5)P₂ depletion. Quantitative analysis revealed that neither the size nor the fluorescence intensity of residual clusters increased following PI(4,5)P₂ restoration (Figure 6—figure supplement 3E–H), arguing against recruitment of Gag(G2A) molecules to residual assembly sites.

To compare the dynamics of assembly site formation for induced and reinduced assembly, we analyzed assembly rates following PI(4,5)P₂ depletion and reconstitution. Spinning disk confocal microscopy was performed at high time resolution in a 3D volume to identify and track single assembly sites at the lateral PM (Video 8). Figure 6—figure supplement 4A shows normalized and averaged traces of 96 individual reinduced assembly sites from three cells.

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For a quantitative comparison, data from each individual trace were fitted to a single exponential equation to extract the assembly rate constant k (Figure 6—figure supplement 4C). The mean assembly rate constant of k = 5.8 ± 0.6*10⁻³ s⁻¹ for reinduced assembly, which was derived from averaging all individual k values, was slightly, but significantly, increased compared to native assembly (native: k = 4.5 ± 0.5*10⁻³ s⁻¹, p<0.0001) (Figure 6—figure supplement 4C). The assembly constant for reinduced assembly translates into a half-time of ~120 s (native: t_1/2 ~150 s). Native and induced assembly kinetics were both determined at the ventral membrane, while reinduced assembly had to be measured at the lateral membrane, and this difference could influence the observed assembly rates. We therefore determined assembly kinetics of native Gag assembly site formation at the lateral membrane as well. Figure 6—figure supplement 4B shows normalized and averaged traces of 169 individual native assembly sites at the lateral PM of six cells. The average assembly constant derived from single exponential fits for all native assembly traces under these conditions of k = 5.9 ± 0.4*10⁻³ s⁻¹ (Figure 6—figure supplement 4C) was very similar to that observed for reinduced assembly (no significant change, p=0.1569) and was again significantly faster than observed for assembly at the ventral membrane (p<0.0001). The half-time of native assembly at the lateral PM was changed accordingly (t_1/2 ~115 s). The observed difference may reflect effects of the adjacent glass substrate on the kinetics of HIV-1 Gag assembly at the ventral membrane, but clearly shows that reinduced and native assembly site formation occur with very similar kinetics. The distribution of rate constants also did not reveal any major difference between reinduced and native assembly (Figure 6—figure supplement 4D).

Finally, we asked whether the reinduced HIV-1 Gag assemblies were morphologically normal using STED nanoscopy. Since depletion from the ventral PM was inefficient, we again focused on the lateral PM for these analyses. As shown in Figure 8A, late native HIV-1 assemblies at the lateral PM appeared very similar to those detected at the ventral PM (compare Figure 4). In both cases, condensed Gag clusters with a diameter of roughly 120 nm surrounded by larger Env clusters were detected in untreated cells. Reinduced assembly sites also displayed this distinct phenotype (Figure 8B) and were undistinguishable from native assembly sites at the lateral PM with respect to diameter and morphology in micrographs and line plot diagrams of Gag and Env (Figure 8C and D).

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PM association of HIV-1 Gag - and consequently assembly site formation and virus production - have previously been shown to depend on PM PI(4,5)P₂. Here, we made use of a recently established chemical biology tool allowing rapid and reversible manipulation of PI(4,5)P₂ levels at the PM in living cells (Feng et al., 2014; Schifferer et al., 2015) to study the role of PI(4,5)P₂ for HIV-1 assembly site formation and maintenance. This rCDS has several advantages compared to alternative strategies. First, PI(4,5)P₂ depletion is strictly dependent on rCD1 addition and can be rapidly achieved within minutes. This allows maintaining normal cell metabolism prior to compound addition despite the presence of all rCDS components in the cell, and should thus prevent induction of escape pathways. Second, the system is completely and rapidly reversible, allowing restoration of PM PI(4,5)P₂ levels within minutes of FK506 addition. Here, we show that long-term depletion of PM PI(4,5)P₂ is compatible with cell viability and viral protein expression, while completely preventing Gag membrane targeting and assembly site formation. Furthermore, PI(4,5)P₂ depletion also caused loss of pre-assembled Gag lattices from the PM and large clusters removed from the membrane appeared to dissociate into monomers or small oligomers in the cytosol. Restoration of PM PI(4,5)P₂ in these cells reinduced assembly site formation even in the absence of new protein synthesis indicating that the dissociated Gag molecules remained assembly competent. These results reveal an important role of PI(4,5)P₂ beyond Gag recruitment to the PM and suggest that Gag interactions with PM lipids may be more dynamic than previously thought.

PI(4,5)P₂ and MA-dependent PM targeting of Gag and assembly site formation is consistent with previous studies showing that continuous overexpression of 5Ptase causes loss of Gag targeting in Gag-producing cells, that Gag-derived proteins preferentially interact with PI(4,5)P₂ in vitro in an MA-dependent manner and that PI(4,5)P₂ was found to be enriched in virions (Ono et al., 2004; Saad et al., 2006; Chukkapalli et al., 2008; Chan et al., 2008). Applying an inducible system in the current study allowed us to investigate the kinetics of assembly site formation. Our results show that rate constants for native and induced assembly, and even for reinduced assembly following depletion and subsequent restoration of PI(4,5)P₂, are virtually identical. Given that a large pool of myristoylated and assembly competent Gag is present in the cells prior to induction (or reinduction), one may have expected assembly to occur faster than under native conditions, but this was not the case. This observation is consistent with the assumption that PI(4,5)P₂ is important to retain Gag once it reaches the PM, but does not affect its PM trafficking, which may be rate-limiting under these conditions. Interestingly, Gag assembly site formation was not faster, but more synchronous under induced or reinduced assembly conditions compared to native assembly. This finding most likely reflects the presence of a large pool of assembly competent Gag in PI(4,5)P₂ depleted cells, which can only be retained at the PM upon PI(4,5)P₂ restoration. Gag molecules may transiently associate with all cellular membranes including the PM in the absence of PI(4,5)P₂, but are stably retained only in the presence of this phosphoinositide. Assembly site formation therefore is synchronized by the rCDS, which provides a promising tool to study the role of host components during HIV-1 assembly and the induction of virus maturation. Analysis of these processes is so far limited by the inherently asynchronous nature of virus production – between cells and within an individual cell – and synchronizing this process via the described approach can overcome this limitation. The observation that virions released under those conditions are infectious and the opportunity to use stable HeLa_rCDS cells for bulk assays broadens the range of applications for this system.

The finding that PI(4,5)P₂ is also required to retain partially or completely assembled Gag lattices at the PM was unexpected, since a triggered myristoyl switch in conjunction with ionic interactions between the HBR and negatively charged lipids of the inner leaflet would be expected to retain Gag at the PM, especially considering the high avidity of the multimeric Gag lattice (ca. 2,500 Gag molecules per assembly site). Large PM Gag clusters resembling fully assembled HIV-1 budding structures were rapidly and completely lost from the PM upon PI(4,5)P₂ depletion, however. Thus, PI(4,5)P₂ is clearly required throughout assembly and budding of HIV-1 particles and not only for Gag membrane targeting. In vitro, MA binding to liposomes is significantly weaker for PI(4)P compared to PI(4,5)P₂ (Mercredi et al., 2016), suggesting that Gag dissociation from the PM may be directly caused by dephosphorylation of PI(4,5)P₂ and consequent loss of Gag interaction. It is not entirely clear, however, whether Gag-associated phosphoinositides are good substrates for the phosphatase since the enzyme would have to enter the tight protein lattice of the assembling virion. Accordingly, effects of general depletion of PM PI(4,5)P₂ with lipid exchange of Gag molecules must also be considered.

Our results are not easily reconciled with the suggested lipid flip, where the 2´acyl chain of PI(4,5)P₂ would be pulled from the membrane and exchanged for the myristic acid once the myristate switch is triggered. As discussed by Summers and colleagues (Mercredi et al., 2016), extrusion of the 20-carbon 2´-acyl chain of PI(4,5)P₂ would require ~24 kcal/mol (Tanford, 1979; Israelachvili et al., 1977), while inserting 10 carbons of myristate into the lipid bilayer would gain only 8 kcal/mol. More importantly, our results appear inconsistent with the continuous insertion of the myristate moiety of all or the majority of Gag molecules into the inner leaflet. While the binding energy of the myristate anchor is not sufficient to retain individual Gag molecules at the PM, the avidity effects of up to 2500 Gag molecules with all or the majority of their N-terminal myristate groups inserted into the inner leaflet should make membrane association of the Gag lattice almost irreversible, independent of PI(4,5)P₂. Given that the binding energy of myristate interaction with MA is the same as for extrusion of myristate from the PM (8 kcal/mol) (Charlier et al., 2014), we suggest a dynamic equilibrium between fully membrane-inserted and MA-bound PI(4,5)P₂ molecules at the assembly site. Accordingly, myristate may readily flip between the inner leaflet of the PM and the lipid binding pocket of MA. Once Gag reaches the PM, PI(4,5)P₂ association triggers the myristate switch, and this effect is enhanced by the presence of acidic phospholipids. The combination of specific PI(4,5)P₂ binding and myristate insertion into the PM then anchors the Gag molecule and makes it available for virus assembly. Subsequently, a dynamic exchange of lipid interactions may occur, where depletion of PM PI(4,5)P₂ causes loss of Gag molecules once their myristate moieties flip out from the membrane.

Finally, our results also shed some light on the question whether Gag assembly is directed to pre-existing assembly-prone membrane microdomains or whether Gag creates its own microdomain. Comparison of the membrane localization of Gag assembly sites prior to PI(4,5)P₂ depletion with assembly sites in the same region of the cell after PI(4,5)P₂ restoration indicated that they occur in different positions. The conclusion that there are no stable assembly-prone membrane microdomains is consistent with earlier observations showing that Gag initially binds to non-raft regions of the PM and subsequently either laterally associates with raft like domains or induces their formation (Ono and Freed, 2001; Mercredi et al., 2016).

Collectively, our study shows that PI(4,5)P₂ at the PM is crucial throughout the entire HIV-1 assembly process and suggests a dynamic equilibrium of Gag-lipid interactions. Furthermore, it establishes an experimental system that permits synchronized induction of HIV-1 assembly by targeted modulation of Gag PM targeting.

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Author details

1. Frauke Mücksch

   Department of Infectious Diseases, Virology, University Hospital Heidelberg, Heidelberg, Germany

   Contribution

   FM, Conceptualization, Formal analysis, Investigation, Visualization, Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared.

   "This ORCID iD identifies the author of this article:" 0000-0002-0132-5101

2. Vibor Laketa

   1. Department of Infectious Diseases, Virology, University Hospital Heidelberg, Heidelberg, Germany
   2. German Center for Infectious Disease Research, Partner site Heidelberg, Braunschweig, Germany

   Contribution

   VL, Conceptualization, Supervision, Validation, Investigation, Writing—review and editing

   Competing interests

   No competing interests declared.

3. Barbara Müller

   Department of Infectious Diseases, Virology, University Hospital Heidelberg, Heidelberg, Germany

   Contribution

   BM, Conceptualization, Resources, Supervision, Funding acquisition, Writing—review and editing

   Competing interests

   No competing interests declared.

   "This ORCID iD identifies the author of this article:" 0000-0001-5726-5585

4. Carsten Schultz

   1. Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
   2. Department of Physiology and Pharmacology, Oregon Health and Science University, Portland, United States

   Contribution

   CS, Resources, Funding acquisition, Writing—review and editing

   Competing interests

   CS: C. Schultz is a shareholder of the company SiChem, which distributes rCDS

5. Hans-Georg Kräusslich

   1. Department of Infectious Diseases, Virology, University Hospital Heidelberg, Heidelberg, Germany
   2. German Center for Infectious Disease Research, Partner site Heidelberg, Braunschweig, Germany

   Contribution

   H-GK, Conceptualization, Supervision, Funding acquisition, Writing—original draft, Project administration, Writing—review and editing

   For correspondence

   hans-georg.kraeusslich@med.uni-heidelberg.de

   Competing interests

   No competing interests declared.

   "This ORCID iD identifies the author of this article:" 0000-0002-8756-329X

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