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. Author manuscript; available in PMC: 2017 Nov 9.
Published in final edited form as: Cell Rep. 2017 Sep 19;20(12):2792–2799. doi: 10.1016/j.celrep.2017.08.076

Localized phosphorylation of a kinesin-1 adaptor by a capsid-associated kinase regulates HIV-1 motility and uncoating

Viacheslav Malikov 1, Mojgan H Naghavi 1,*
PMCID: PMC5679262  NIHMSID: NIHMS903155  PMID: 28930676

Summary

Although microtubule motors mediate intracellular virus transport, the underlying interactions and control mechanisms remain poorly defined. This is particularly true for HIV-1 cores, which undergo complex, interconnected processes of cytosolic transport, reverse transcription and uncoating of the capsid shell. Although kinesins have been implicated in regulating these events, curiously there are no direct kinesin-core interactions. We recently showed that the capsid-associated kinesin-1 adaptor protein, fasciculation and elongation protein zeta-1 (FEZ1) regulates HIV-1 trafficking. Here, we show that FEZ1 and kinesin-1 heavy, but not light chains regulate not only HIV-1 transport, but also uncoating. This required FEZ1 phosphorylation, which controls its interaction with kinesin-1. HIV-1 did not stimulate widespread FEZ1 phosphorylation, but instead bound MT affinity-regulating kinase 2 (MARK2) to stimulate FEZ1 phosphorylation on viral cores. Our findings reveal that HIV-1 binds a regulatory kinase to locally control kinesin-1 adaptor function on viral cores, thereby regulating both particle motility and uncoating.

eTOC Blurb

Many viruses exploit microtubule networks to facilitate their movement within the cell, yet the underlying mechanisms remain poorly understood. In this report, Malikov and Naghavi demonstrate how HIV capsids interact with host microtubule regulatory factors to control their transport and disassembly during early post-entry stages of infection.

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Introduction

Microtubule (MT) networks and their associated motors are important for directed long-range intracellular transport. MTs are polarized filaments whose plus-ends direct toward the cell periphery, while their minus-ends are usually anchored at microtubule organizing centers (MTOCs). Generally, dynein motor complexes bind cargoes and direct their motion towards the minus-end, while plus-end directed motion is driven by kinesin motor complexes (Welte, 2004). As cargoes themselves, many viruses including human immunodeficiency virus type 1 (HIV-1), have evolved various strategies to exploit MTs for their intracellular movement throughout their lifecycle (Dodding and Way, 2011; Naghavi and Walsh, 2017).

Although the early post-entry steps of HIV-1 infection are still poorly understood, it is known that after cell entry the HIV-1 RNA genome is reverse transcribed into double-stranded (ds) DNA that enters the nucleus and integrates into the host genome. Given the size of the viral core, the capsid (CA) shell has to disassemble or “uncoat” to allow the genome to pass through nuclear pores (Campbell and Hope, 2015). Although some CA can be detected in the nucleus and functions in processes after nuclear entry (Chen et al., 2016; Hulme et al., 2015; Koh et al., 2013; Peng et al., 2014; Schaller et al., 2011; Stultz et al., 2017), whether reverse transcription and uncoating occur during transport along MTs in the cytoplasm or at nuclear pores remains a matter of debate (Arhel et al., 2007; Fassati and Goff, 2001; Le Sage et al., 2014; McDonald et al., 2002; Miller et al., 1997; Rasaiyaah et al., 2013; Zhou et al., 2011). However, recent live cell imaging supports the notion that uncoating begins in the cytoplasm (Francis et al., 2016). Notably, uncoating and reverse transcription are intricately coupled processes (Cosnefroy et al., 2016; Hulme et al., 2011; Rankovic et al., 2017; Stultz et al., 2017; Yang et al., 2013) that are further intertwined with the bi-directional movement of HIV-1 particles. Interfering with MT or dynein function suppresses HIV-1 trafficking and delays uncoating during early infection (Arhel et al., 2006; Delaney et al., 2017; Lukic et al., 2014; Malikov et al., 2015; McDonald et al., 2002; Pawlica and Berthoux, 2014; Sabo et al., 2013). Although dynein heavy or light chain subunits, DYNC1H1 or DYNL1, promote or delay uncoating, respectively, DYNL1 interacts with integrase and may act independently of the dynein motor (Jayappa et al., 2015). Kinesin-1 (formerly conventional kinesin or Kif5) heavy chain, Kif5B is also required for efficient uncoating (Lukic et al., 2014) but the underlying mechanism is unclear, particularly since interactions between motors and viral cores have eluded detection. We recently showed that the Kif5 adaptor, FEZ1 associates with in vitro assembled HIV-1 capsid-nucleocapsid (CA-NC) complexes and regulates early HIV-1 transport (Malikov et al., 2015). Here, we show that FEZ1 and Kif5B, but not kinesin-1 light chain (KLC1) not only regulate HIV-1 trafficking but also control uncoating. In addition, we find that early HIV-1 infection is promoted by MARK2/PAR-1 (Drewes et al., 1997), a kinase that phosphorylates FEZ1 to control its interaction with Kif5 (Butkevich et al., 2016). HIV-1 did not globally activate MARK2 or FEZ1 phosphorylation, but HIV-1 cores bound MARK2 to phosphorylate FEZ1 on viral particles. These findings reveal how HIV-1 binds and usurps a host kinase to locally activate a motor adaptor to control core motility and uncoating.

Results and Discussion

While many cargos exploit KLCs to bridge heavy chain motors for movement, some use adaptors such as FEZ1 (Blasius et al., 2007; Chua et al., 2012). Notably, depletion of either FEZ1, Kif5A or Kif5B blocks early HIV-1 infection (Lukic et al., 2014; Malikov et al., 2015). Testing whether light chains contribute to infection, we found that while siRNA-mediated depletion of Kif5B inhibited early infection with HIV-1 carrying either vesicular stomatitis virus G (VSV-G) or wild-type (WT) envelope in both primary normal human dermal fibroblasts (NHDF) or human microglia CHME3 cells, KLC1 depletion had no effect (Figure S1A–F). Live or fixed imaging using HIV-1 particles labeled with GFP-tagged Vpr (HIV-1-GFP-Vpr) further revealed that the movement and perinuclear accumulation of viruses that enter using either VSV-G or WT envelopes was suppressed in Kif5B-depleted NHDFs or CHME3, while control siRNAs or KLC1 depletion had no effect (Figure S1G–J, S2A–B and Supplemental Movies 1–2). This contrasts with reports that Kif5B depletion does not block the accumulation of HIV-1 at the nucleus in HeLa cells (Dharan et al., 2016). However, Kif5B and Kif5C are functionally redundant, with Kif5C normally expressed at high levels in neurons (Hirokawa and Takemura, 2005). We find that Kif5B is universally expressed in NHDFs, HeLa and cell lines originating from distinct natural target cell types for HIV-1 infection (CHME3 and Jurkat cells), but only HeLa cells express detectable Kif5C (Figure 1A). As such, the role of Kifs in HIV-1 transport in HeLa cells is complicated by functional redundancy not observed in primary fibroblasts or in microglia or T cell lines, explaining this apparent discrepancy. Finally, noting the cell-specific role of Kif5B in HIV-1 transport, we recently found that expression of a FEZ1 Ser58-Ala mutant (S58A) that fails to bind kinesin-1 suppresses HIV-1 infection in NHDFs (Malikov et al., 2015). Live or fixed imaging using HIV-1-GFP-Vpr particles carrying WT envelope confirmed that FEZ1-S58A expression also blocked HIV-1 transport in CHME3 (Figure S3, and Supplemental Movies 3). Overall, these findings identified HIV-1 as a KLC-independent cargo that utilizes FEZ1 to bridge Kif5B for transport in a variety of cell types including microglia.

Figure 1. FEZ1 regulates HIV-1 uncoating as measured by two independent assays.

Figure 1

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(A) Endogenous levels of Kif5B and Kif5C in NHDF, CHME3, HeLa and Jurkat cell lines. (B–E) RNAi-mediated depletion of FEZ1 (B) or Kif5B (C) does not affect HIV-1 fusion as detected by the loss of S15 from virus particles (D) while it delays the loss of p24 (uncoating) as measured by the amounts of p24-associated fluorescence with S15 negative viral particles (E–F) at the indicated h.p.i in CHME3 cells infected with GFP-Vpr/S15-Tomato labeled HIV-1. (G) Measurements of soluble and particulate p24 CA in HIV-1-VSV-luc infected CHME3 cells depleted of FEZ1 or Kif5B. (H) Quantification of % pelletable capsid from samples in G. Data shown as mean ± SD, n= 3.

See also Figure S1, S4 and Table S1.

Given that Kif5B has been implicated in HIV-1 uncoating in HeLa cells, we next tested whether FEZ1 might regulate uncoating in microglia using in situ fluorescence microscopy (Delaney et al., 2017; Lukic et al., 2014). To do this, siRNA-treated CHME3 cells were infected with VSV-G pseudotyped virus carrying GFP-Vpr (labeling cores) and S15-Tomato (labeling viral membranes). Importantly, virus preparations used were ~95% double-labeled. Cells were fixed at various time and stained for p24. Fusion was assessed by the loss of S15 (membrane) from virus particles, while uncoating was assessed by the amounts of p24 associated with fused (S15-negative) viral particles. While depletion of either FEZ1 or Kif5B did not affect virus fusion (Figure 1B–D and Table S1), loss of p24 uncoating was delayed in either FEZ1- or Kif5B-depleted CHME3 cells (Figure 1E–F). To independently confirm these findings, fate-of-capsid uncoating assays based on the biochemical separation of soluble capsid protein from particulate (pelletable) capsid cores were used (Delaney et al., 2017). To do this, siRNA-treated CHME3 cells were infected for 3h. Cells treated with PF74, a small molecule that destabilizes capsids and inhibits reverse transcription at high concentrations (Saito et al., 2016; Shi et al., 2011), were included to control for detection of uncoating. In line with in situ fluorescence microscopy, depletion of FEZ1 or Kif5B increased the recovery of intact pelletable HIV-1 cores compared to control siRNA-treated cultures, while PF74 destabilized cores (Figure 1G–H and S4A). This demonstrated that both FEZ1 and Kif5B were required for efficient uncoating.

HIV-1 cores have not been reported to bind kinesins directly, but they efficiently interact with FEZ1 (Figure S4B and (Malikov et al., 2015)). This suggested that FEZ1 might mediate Kif5B’s effects on uncoating. To test this, we examined the recovery of pelletable HIV-1 cores from CHME3 lines stably expressing Flag-tagged FEZ1 or FEZ1-S58A (Figure 2A). Notably, unlike NHDFs, CHME3 express high levels of endogenous FEZ1 and FEZ1 overexpression in these lines does not enhance infection. In line with this, the levels of pelletable capsid were found to be similar in control and FEZ1 CHME3, while PF74 again destabilized capsids (Figure 2B–C and S4C). In stark contrast, expression of FEZ1-S58A resulted in a large increase in pelletable capsids and a decrease in infection by HIV-1 carrying WT envelope (Figure 2D). Expression of FEZ1-S58A also suppressed infection by HIV-1 carrying WT envelope in Jurkat cells (Figure 2E). This demonstrated that binding to Kif5B is required for FEZ1 to promote HIV-1 uncoating and infection in both microglia and T cells. As HIV-1 cores have not been found to directly interact with Kif5B, and Kif5B is expressed in FEZ1-S58A-expressing cells that exhibit defects in transport and uncoating, our data suggests that FEZ1 is likely to be the bridging factor or mediator that enables kinesins to control HIV-1 motility and capsid disassembly.

Figure 2. Binding to kinesin-1 is required for FEZ1 to regulate HIV-1 uncoating and early infection.

Figure 2

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(A–B) Stable expression of S58A-Flag, but not Flag control or FEZ1-Flag, increases the recovery of pelletable capsid as determined by measurements of soluble and particulate p24 CA levels in CHME3 lines infected with HIV-1-VSV-luc.. (C) Quantification of % pelletable capsid from samples in B. (D–E) HIV-1-WT-luc infection is suppressed in CHME3 (D) or Jurkat cells (E) expressing S58A-Flag, but not control Flag or FEZ1-Flag. C–E are shown as mean ± SD, n= 3.

See also Figure S3–S4.

Different models have been proposed for how HIV-1 transport and uncoating might be coupled (Campbell and Hope, 2015). One proposes that bi-directional motility itself pulls the viral core apart due to “tug-of-war” forces between opposing motors on the same particle (Ally et al., 2009; Welte, 2004). However, HIV-1 exhibits fast bi-directional motility in FEZ1-depleted or FEZ1-S58A-expressing cells but with net outward rather than inward directionality, as seen in microglia here and previously in NHDFs (Malikov et al., 2015). This suggests that FEZ1 regulates kinesin-1 activity to tilt the balance of bi-directional motility toward a net retrograde movement. This also implies that “tug of war” forces alone may be insufficient to drive uncoating, although full force and directionality of movement may be important. Another suggestion posits that motors influence the localization of particles to regions of the cell where uncoating occurs. While dynein perturbation or Kif5B depletion cause viral particles to remain near the cell periphery (Arhel et al., 2006; Malikov et al., 2015; McDonald et al., 2002), particles are distributed throughout the cytoplasm in FEZ1-depleted cells. Although particles localize to different subcellular regions under each condition, in each case particles notably fail to reach the nucleus. Kif5B binds RanBP2/Nup358 (Cho et al., 2009), a HIV-1 nuclear entry factor (Schaller et al., 2011), while Kif5B and Nup358 were recently shown to cooperatively mediate nuclear import of HIV-1 (Dharan et al., 2016). As such, while FEZ1 may regulate uncoating directly, FEZ1 could potentially facilitate nuclear docking both by enabling virus particles to reach the nucleus in the first place, and by bridging pore-associated Kif5B and HIV-1 cores. Until the location of uncoating is unequivocally defined and better spatio-temporal assays are developed this remains difficult to test, but our findings identify FEZ1 as a capsid-associated motor adaptor whose binding to Kif5 is required for efficient HIV-1 uncoating.

To understand how HIV-1 regulates FEZ1 activity we noted that MARKs were recently shown to phosphorylate FEZ1 at S58 in C. elegans (Butkevich et al., 2016). We confirmed this for GFP-tagged human MARK2 (Lin et al., 2009) when expressed in 293T cells along with Flag-tagged FEZ1 using a phospho-specific antibody against S58 (Butkevich et al., 2016) (Figure 3A). The specificity of this antibody signal was confirmed by failure to detect phosphorylation on the FEZ1-S58A mutant. Notably, an infection time course revealed that HIV-1 did not induce phosphorylation of FEZ1 globally (Figure 3B). However, MARK2 was found to bind in vitro assembled CA-NC complexes similarly to its substrate FEZ1 (Figure 3C). To test if MARK2 binding phosphorylated FEZ1 on capsids, 293T cells were transfected with GFP control or GFP-MARK2 in combination with Flag or FEZ1-Flag. Input samples confirmed overexpressed MARK2 increased FEZ1 S58-phosphorylation, while MARK2 and FEZ1 independently bound CA-NC complexes (Figure 3D). Notably, in MARK2 and FEZ1 co-expressing samples, the amounts of CA-NC-bound phosphorylated FEZ1 were greatly increased. Seemingly paradoxically, MARK2 levels were decreased compared to levels observed when MARK2 was expressed alone. There are a number of potential explanations for this phenomenon. First, both proteins potentially share a common binding site and FEZ1 outcompetes MARK2 when both are overexpressed. Alternatively, interactions between FEZ1 and MARK2 may alter their relative affinities for viral cores, favoring FEZ1. Finally, MARK2 may exchange more quickly on the capsid when it has more FEZ1 present to phosphorylate. Indeed, kinases often rapidly release from their substrates upon phosphorylation, possibly suggesting a dynamic association of the FEZ1 kinase with its capsid-bound substrate. In line with this, although HIV-1 did not globally activate FEZ1 phosphorylation, an increase in early infection with HIV-1 carrying either VSV-G or WT envelope was observed in either 293T or Jurkat cells expressing MARK2-GFP compared to control GFP-expressing cells (Figure 4A and B, respectively). Moreover, FEZ1 depletion abrogated the ability of MARK2 expression to enhance infection (Figure 4C), suggesting that MARK2 exerted its effects through its substrate, FEZ1. The modest yet statistically significant increases in HIV-1 infection reflected the fact that high levels of MARK2 expression could not be achieved, as this leads to MT breakdown and cell death (Timm et al., 2003). This also offers a potential reason for why HIV-1 might avoid widespread activation of MARK2. Instead, HIV-1 binds MARK2 to regulate FEZ1 phosphorylation on HIV-1 cores, locally controlling FEZ1 activity while avoiding the detrimental effects of widespread MARK2 activation on the structure of MT networks needed for virus transport. Given the lack of direct interactions with motor proteins, our findings provide mechanistic insights into how HIV-1 usurps a host kinase and motor adaptor to bridge to kinesins, locally controlling their activity on the capsid surface during early post-entry stages of infection.

Figure 3. MARK2 binds in vitro assembled HIV-1 CA-NC complexes.

Figure 3

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(A) GFP-tagged MARK2 (GFP-MARK2) expression in 293T cells results in phosphorylation of FEZ1-Flag at S58, which is not detected with a FEZ1-S58A-Flag mutant. (B) Time course infection demonstrating HIV-1 does not induce phosphorylation of FEZ1 in HIV-1-VSV infected 293T cells at indicated h.p.i. (C) Similar to FEZ1-Flag, GFP-MARK2, but not control GAPDH-HA, binds to in vitro assembled HIV-1 CA-NC complexes. (D) GFP-MARK2, but not GFP control, increases FEZ1 phosphorylation at S58 in 293T cells co-expressing GFP-MARK2 and FEZ1-Flag. Results in A–D are representative of three or more independent experiments.

See also Figure S4.

Figure 4. Overexpression of MARK2 enhances HIV-1 infection.

Figure 4

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(A–B) Transient or stable expression of MARK2-GFP, but not control GFP, increases infection by HIV-1-VSV-Luc in 293T cells (A) or HIV-1-WT-Luc in Jurkat cells (B) at different virus dilutions. (C) RNAi-mediated FEZ1 depletion abrogates the ability of MARK2-GFP to enhance HIV-1-VSV-Luc infection. Data in A–C, upper panels, are shown as mean ± SD, n= 3.

Experimental Procedures

Cells, viruses and infections

293T, CHME3 and NHDF cells were described previously (Malikov et al., 2015). Jurkat cells were a kind gift from Thomas Hope. Cells were infected using HIV-1 pseudotyped with VSV-G or WT envelope carrying either a luciferase reporter (HIV-1-VSV-Luc or HIV-1-WT-Luc, respectively), GFP-Vpr or GFP-Vpr/S15-Tomato (Delaney et al., 2017). Virus titers/multiplicity of infections (m.o.i) were as follow: HIV-1-VSV-Luc 3×106/0.01 to 0.04 for luciferase assay and 3×106/1.5 for fate-of-capsid assay; HIV-1-WT-Luc 2.7×106/0.3; HIV-1-VSV-GFP-Vpr 2.7×106/2.7; HIV-1-WT-GFP-Vpr 2.5×106/2.5; double-labeled HIV-1-VSV (GFP-Vpr/S15-Tomato) 2.8×106/1.4; and HIV-1 no Env control 3.2×106/1.6.

Antibodies and Western Blotting

Antibodies used for WB were anti-FEZ1 (HPA038490), anti-HA (H6098) and anti-Flag (F7425) from Sigma; anti-Kif5B (ab15705), anti-Kif5A (ab25715), anti-KLC1 (ab174273), anti-GFP (ab13970), anti-HIV-1 p55/24/17 (ab63917), anti-HIV1 p24 (ab9071), and anti-Kif5C (ab5630) from Abcam; anti-GAPDH (sc-25778) from Santa Cruz; anti-elF4E (610269) from BD Biosciences; anti-GFP (2555) and anti-PARP (9542) from Cell Signaling; anti-MARK (MBS8221493) from MyBioSource; anti-HIV-1 p24 (AG3.0) from NIH AIDS Reagent Program; anti-phospho-S58-FEZ1 was a kind gift from Dr. John Jia En Chua. For WB analysis all primary antibodies were used at 1:1000 dilution with exception of anti-MARK and anti-phospho-S58-Fez1, which were diluted 1:500. The primary antibodies were detected using the appropriate HRP-conjugated secondary.

RNA interference (RNAi) and transient expression

Transient knockdowns were performed using the following siRNAs: ID# 45101 (FEZ1-C), ID# s731 (Kif5B), ID# s7900 (KLC1), and ID# AM4635 (control non-targeting siRNA) from Ambion. For transient expression of control GFP or GFP-tagged MARK2, 293T cells were transfected with 1.2 μg of pEGFP-C1 or a C′ terminally GFP-tagged MARK2 (Addgene plasmid # 66706, deposited by Gary Banker) or a N′ terminally GFP-tagged MARK2 (GFP-MARK2) expressing construct (pEGFP-EMK C1, kindly provided by Andrew S. Shaw)(Lin et al., 2009) in a 12-well plate or with 5–10 μg of each of the constructs in a 100 mm dish using polyethylenimine. Transfected cells were then infected with HIV-1-VSV-Luc or HIV-1-WT-Luc followed by cell lysis and measurement of luciferase activity (Malikov et al., 2015).

Live imaging, particle tracking and analysis

Infections with GFP-Vpr labeled HIV-1 were performed for 2h for VSV-G pseudotyped virus or 30 min for WT envelope via spinoculation followed by live cell video microscopy at 1 frame per 20 sec or at 1 frame per sec for 2h or 5–10 minutes, respectively (Malikov et al., 2015). Particle tracking was analyzed in 300 sec image sequences with Fiji software (Schindelin et al., 2012) using Mosaicsuite plugin (Sbalzarini and Koumoutsakos, 2005). At least two independent experiments were analyzed for each condition, and each experiment consisted of 3–8 cells. Statistical significance was determined using a one-way ANOVA for correlated samples, with a confidence interval of 95% or greater.

Generation of stable pools

Retroviruses encoding either control Flag (Flag), full-length C′ terminally flag-tagged human FEZ1 (FEZ1-Flag) or the FEZ1-S58A (S58A-Flag) mutant were used to infect CHME3 or Jurkat cells, followed by selection to generate stably expressing pools (Malikov et al., 2015).

In vitro capsid binding assay

293T cells were transfected with HA-tagged GAPDH control, C′ terminally Flag-tagged FEZ1, C′ terminally Flag-tagged S58A, or N′ terminally GFP-tagged MARK2 (pEGFP-EMK C1). Cell lysates were incubated with purified in vitro assembled CA-NC (See Supplemental Procedures). Input samples were taken before lysates were ultra-centrifuged through a 70% sucrose cushion followed by re-suspension of the pellet in SDS-PAGE loading buffer (Bound)(Delaney et al., 2017). WB analysis was used to determine the levels of proteins in input and bound samples, with anit-p24 antibody confirming HIV-1 CA-NC pelleting.

In situ uncoating assay

Equal numbers of siRNA-treated CHME3 cells plated on coverslips were infected with GFP-Vpr/S15-Tomato labeled virus via spinoculation. Cells were then fixed and stained for p24 followed by confocal microscopy (see Supplemental Experimental Procedures). Individual virus particles were then analyzed to measure the maximum p24 fluorescence intensity associated with each virus particle that had fused and entered the cytoplasm. Virus particles that did not productively fuse and become endocytosed were S15-Tomato positive and GFP-Vpr positive whereas virus particles that had productively fused into the cytoplasm were S15-Tomato negative and GFP-Vpr positive. The percentage of fusion was determined by dividing the number of fused virus particles by the number of virus particles in each of the fields. Changes in p24 intensity during infection were determined (Delaney et al., 2017). The mean of the maximum p24 fluorescence intensity of each virus particle in control siRNA-treated cells was calculated for each time point. Data was then normalized within each time point by dividing the maximum p24 fluorescence intensity of each virus particle by the mean of the control siRNA-treated cells.

Fate-of-capsid assay

Equal numbers of CHME3 cells treated with siRNAs or stably expressing Flag, FEZ1-Flag or S58A-Flag were infected with HIV-1-VSV-luc or HIV-1 carrying no envelope glycoprotein via spinoculation followed by incubation at 37 °C for 3h (Delaney et al., 2017). Cells were lysed, centrifuged and the supernatant fractions were collected as “Input”. Upon fractionation of the rest of supernatant, the top fractions were collected as “soluble” capsid and pellets were re-suspended in Laemmli buffer (See Supplemental Proceduers).

Statistical Analyses

Statistical significance for two groups was performed using the Student’s t-test. Data are represented as means ± standard errors of the standard deviations (SD). One-way analysis of variance (ANOVA) with post-test was used when more than two groups were compared. Statistical significance are shown by *, ** and ***, which represents P ≤ 0.05, 0.01 and 0.001, respectively.

Supplementary Material

1. Movie 1. Representative movies of HIV-1-GFP-Vpr movement in NHDF cells treated with control siRNA, or siRNAs specific to Kif5B or KLC1, Related to Figure S1.

NHDFs were treated with indicated siRNAs and infected with HIV-1-VSV-GFP-Vpr. Single confocal sections were acquired at 20 sec intervals for 2h on a spinning-disc confocal microscope with heated stage. In the control and KLC1 siRNA treated cultures, the majority of viral particles exhibited bi-directional motility and disappeared from the cytoplasm over the course of acquisition. While in Kif5B knockdown cultures, the majority of viral particles exhibited slow, short-ranged movements at the cell periphery throughout the course of acquisition. Movies are representative of 3 independent experiments. Scale bar = 10 μm.

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2. Movie 2. Representative movies of HIV-1-GFP-Vpr movement in CHME3 cells treated with control siRNA, or siRNAs specific to Kif5B or KLC1, Related to Figure S1.

CHME3 cells were treated with indicated siRNAs and infected with HIV-1-WT-GFP-Vpr. Single confocal sections were acquired at 20 sec intervals for 2h on a spinning-disc confocal microscope with heated stage. The majority of viral particles exhibited bi-directional motility and disappeared from the cytoplasm over the course of acquisition in the control and KLC1 siRNA treated cultures. While in Kif5B knockdown cultures, the majority of viral particles exhibited slow, short-ranged movements at the cell periphery throughout the course of acquisition. Movies are representative of 3 independent experiments. Scale bar = 10 μm.

Download video file (42.2MB, avi)
3. Movie 3. Representative movies of HIV-1-GFP-Vpr movement in CHME3 cells expressing either control Flag alone, FEZ1-Flag, or S58A-Flag, Related to Figure S3.

CHME3 cells stably expressing Flag alone (Flag), full-length C′ terminally flag-tagged FEZ1 (FEZ1-Flag) or a C′ terminally flag-tagged FEZ1 mutant unable to bind kinesin-1 (S58A-Flag) were infected with HIV-1-WT-GFP-Vpr. Single confocal sections were acquired at 20 sec intervals for 2h on a spinning-disc confocal microscope with heated stage. In the Flag or FEZ1-Flag cultures, the majority of viral particles exhibited bi-directional motility and gradually disappeared from the field of view due to long-range transport, nuclear entry and/or uncoating. While in S58A-Flag culture, viral particles exhibited bi-directional movement in the cytoplasm but the majority of particles remained in the field of view through acquisition, failing to either move long distances or disappear with nuclear entry and/or uncoating. Movies are representative of 3 independent experiments. Scale bar = 10 μm.

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4

Highlights.

  • FEZ1 and Kinesin-1 heavy, but not light chains, control HIV-1 motility and uncoating

  • FEZ1’s effects on early infection require phosphorylation at serine 58

  • FEZ1 phosphorylation and early infection are regulated by the kinase MARK2

  • HIV-1 binds MARK2 to locally control motor adaptor function on viral cores

Acknowledgments

We thank Scott Brady, Andrew S. Shaw, and John Jia En Chua for reagents and Vladimir Gelfand and Derek Walsh for fruitful discussions. AG3.0 antibody was obtained from Dr. Jonathan Allan through the NIH AIDS Research and Reference Reagent Program. This study was supported by National Institute of Health (NIH) grant GM101975 and GM105536 and NS099064 to M.H.N.

Footnotes

Author Contributions

V.M., performed the research; V.M., and M.H.N., designed the research and analyzed the data; M.H.N. wrote the manuscript.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

1. Movie 1. Representative movies of HIV-1-GFP-Vpr movement in NHDF cells treated with control siRNA, or siRNAs specific to Kif5B or KLC1, Related to Figure S1.

NHDFs were treated with indicated siRNAs and infected with HIV-1-VSV-GFP-Vpr. Single confocal sections were acquired at 20 sec intervals for 2h on a spinning-disc confocal microscope with heated stage. In the control and KLC1 siRNA treated cultures, the majority of viral particles exhibited bi-directional motility and disappeared from the cytoplasm over the course of acquisition. While in Kif5B knockdown cultures, the majority of viral particles exhibited slow, short-ranged movements at the cell periphery throughout the course of acquisition. Movies are representative of 3 independent experiments. Scale bar = 10 μm.

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2. Movie 2. Representative movies of HIV-1-GFP-Vpr movement in CHME3 cells treated with control siRNA, or siRNAs specific to Kif5B or KLC1, Related to Figure S1.

CHME3 cells were treated with indicated siRNAs and infected with HIV-1-WT-GFP-Vpr. Single confocal sections were acquired at 20 sec intervals for 2h on a spinning-disc confocal microscope with heated stage. The majority of viral particles exhibited bi-directional motility and disappeared from the cytoplasm over the course of acquisition in the control and KLC1 siRNA treated cultures. While in Kif5B knockdown cultures, the majority of viral particles exhibited slow, short-ranged movements at the cell periphery throughout the course of acquisition. Movies are representative of 3 independent experiments. Scale bar = 10 μm.

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3. Movie 3. Representative movies of HIV-1-GFP-Vpr movement in CHME3 cells expressing either control Flag alone, FEZ1-Flag, or S58A-Flag, Related to Figure S3.

CHME3 cells stably expressing Flag alone (Flag), full-length C′ terminally flag-tagged FEZ1 (FEZ1-Flag) or a C′ terminally flag-tagged FEZ1 mutant unable to bind kinesin-1 (S58A-Flag) were infected with HIV-1-WT-GFP-Vpr. Single confocal sections were acquired at 20 sec intervals for 2h on a spinning-disc confocal microscope with heated stage. In the Flag or FEZ1-Flag cultures, the majority of viral particles exhibited bi-directional motility and gradually disappeared from the field of view due to long-range transport, nuclear entry and/or uncoating. While in S58A-Flag culture, viral particles exhibited bi-directional movement in the cytoplasm but the majority of particles remained in the field of view through acquisition, failing to either move long distances or disappear with nuclear entry and/or uncoating. Movies are representative of 3 independent experiments. Scale bar = 10 μm.

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4
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