HIV-1 usurps mixed-charge domain-dependent CPSF6 phase separation for higher-order capsid binding, nuclear entry and viral DNA integration

Sooin Jang; Gregory J Bedwell; Satya P Singh; Hyun Jae Yu; Bjarki Arnarson; Parmit K Singh; Rajalingam Radhakrishnan; AidanDarian W Douglas; Zachary M Ingram; Christian Freniere; Onno Akkermans; Stefan G Sarafianos; Zandrea Ambrose; Yong Xiong; Praju V Anekal; Paula Montero Llopis; Vineet N KewalRamani; Ashwanth C Francis; Alan N Engelman

doi:10.1093/nar/gkae769

. 2024 Sep 11;52(18):11060–11082. doi: 10.1093/nar/gkae769

HIV-1 usurps mixed-charge domain-dependent CPSF6 phase separation for higher-order capsid binding, nuclear entry and viral DNA integration

Sooin Jang ^1,², Gregory J Bedwell ^3,^4,², Satya P Singh ^5,², Hyun Jae Yu ^6,², Bjarki Arnarson ⁷, Parmit K Singh ^8,⁹, Rajalingam Radhakrishnan ¹⁰, AidanDarian W Douglas ¹¹, Zachary M Ingram ¹², Christian Freniere ¹³, Onno Akkermans ¹⁴, Stefan G Sarafianos ¹⁵, Zandrea Ambrose ¹⁶, Yong Xiong ¹⁷, Praju V Anekal ¹⁸, Paula Montero Llopis ¹⁹, Vineet N KewalRamani ²⁰, Ashwanth C Francis ²¹, Alan N Engelman ^22,^23,^✉

¹ Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA

² Department of Medicine, Harvard Medical School, Boston, MA 02115, USA

³ Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA

⁴ Department of Medicine, Harvard Medical School, Boston, MA 02115, USA

⁵ Institute of Molecular Biophysics, Department of Biological Sciences, Florida State University, Tallahassee, FL 32304, USA

⁶ Model Development Section, Cancer Innovation Laboratory, National Cancer Institute, Frederick, MD 21702, USA

⁷ Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA

⁸ Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA

⁹ Department of Medicine, Harvard Medical School, Boston, MA 02115, USA

¹⁰ Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA

¹¹ Institute of Molecular Biophysics, Department of Biological Sciences, Florida State University, Tallahassee, FL 32304, USA

¹² Department of Microbiology and Molecular Genetics, University of Pittsburgh School of Medicine, Pittsburgh, PA 15219, USA

¹³ Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT 06511, USA

¹⁴ Department of Biology, Massachusetts Institute of Technology, Cambridge, MA 02139, USA

¹⁵ Center for ViroScience and Cure, Laboratory of Biochemical Pharmacology, Department of Pediatrics, School of Medicine, Emory University, Atlanta, GA 30322, USA

¹⁶ Department of Microbiology and Molecular Genetics, University of Pittsburgh School of Medicine, Pittsburgh, PA 15219, USA

¹⁷ Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT 06511, USA

¹⁸ MicRoN Core, Harvard Medical School, Boston, MA 02215, USA

¹⁹ MicRoN Core, Harvard Medical School, Boston, MA 02215, USA

²⁰ Model Development Section, Cancer Innovation Laboratory, National Cancer Institute, Frederick, MD 21702, USA

²¹ Institute of Molecular Biophysics, Department of Biological Sciences, Florida State University, Tallahassee, FL 32304, USA

²² Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA

²³ Department of Medicine, Harvard Medical School, Boston, MA 02115, USA

^✉

To whom correspondence should be addressed. Tel: +1 6176324361; Fax: +1 6176324338; Email: alan_engelman@dfci.harvard.edu

The second, third and fourth authors should be regarded as Joint Second Authors.

PMCID: PMC11472059 PMID: 39258548

Abstract

HIV-1 integration favors nuclear speckle (NS)-proximal chromatin and viral infection induces the formation of capsid-dependent CPSF6 condensates that colocalize with nuclear speckles (NSs). Although CPSF6 displays liquid-liquid phase separation (LLPS) activity in vitro, the contributions of its different intrinsically disordered regions, which includes a central prion-like domain (PrLD) with capsid binding FG motif and C-terminal mixed-charge domain (MCD), to LLPS activity and to HIV-1 infection remain unclear. Herein, we determined that the PrLD and MCD both contribute to CPSF6 LLPS activity in vitro. Akin to FG mutant CPSF6, infection of cells expressing MCD-deleted CPSF6 uncharacteristically arrested at the nuclear rim. While heterologous MCDs effectively substituted for CPSF6 MCD function during HIV-1 infection, Arg-Ser domains from related SR proteins were largely ineffective. While MCD-deleted and wildtype CPSF6 proteins displayed similar capsid binding affinities, the MCD imparted LLPS-dependent higher-order binding and co-aggregation with capsids in vitro and in cellulo. NS depletion reduced CPSF6 puncta formation without significantly affecting integration into NS-proximal chromatin, and appending the MCD onto a heterologous capsid binding protein partially restored virus nuclear penetration and integration targeting in CPSF6 knockout cells. We conclude that MCD-dependent CPSF6 condensation with capsids underlies post-nuclear incursion for viral DNA integration and HIV-1 pathogenesis.

Graphical Abstract

Introduction

The retrovirus human immunodeficiency virus 1 (HIV-1; please refer to Supplementary Table S1 for a list of abbreviations used) remains a global health concern. While antiviral inhibitors can achieve high cure rates for some viral diseases (1), the integration step of the retroviral lifecycle underlies the formation of a replication-competent HIV reservoir that persists despite decades-long antiretroviral therapy (2). Integration is mediated by the viral integrase (IN) protein, which has emerged in recent years as a high-value drug target (3). Considered alongside the fact that lentiviral vectors based on HIV-1 are actively used to treat genetic disorders (4), there is intense interest to understand the mechanisms and consequences of HIV-1 integration.

HIV-1 integration favors active chromatin as marked by: genes, gene density (5), histone modifications (6), pre-mRNA splicing (7), Hi-C sub-compartment A1 (8), speckle-associated domains (SPADs) (9) and the SPIN (spatial position inference of the nuclear) state ‘Speckle’ (10,11). Conversely, integration disfavors heterochromatin-associated histone marks (6), lamina-associated domains (LADs) (12,13), sub-compartment B2 (8) and lamina-proximal SPIN states (10).

HIV-1 capsid engages a myriad of ‘FG cofactors’ (host proteins with minimally one Phe-Gly dipeptide) to facilitate cytoplasmic trafficking (14), transport through the nuclear pore complex (NPC) (15–17) and intranuclear incursion [reviewed in (18)]. One such protein, cleavage and polyadenylation specificity factor (CPSF) 6, plays a key role in HIV-1 nuclear entry and integration targeting (13,19,20). Viruses unable to interact with CPSF6 arrest at the nuclear rim (13,20–23) and uncharacteristically target LADs and lamina-proximal chromatin for integration (10,13). In this study, we have investigated the molecular mechanisms of CPSF6-capsid binding and HIV-1 nuclear penetration/integration targeting, focusing on the protein's liquid-liquid phase separation (LLPS) activity (24).

CPSF6 is a serine-arginine (SR)-related protein (25) that primarily regulates sites of pre-mRNA polyadenylation (26). Canonical SR proteins, including SR-rich splicing factor 1 (SRSF1; a.k.a. ASF/SF2) and SRSF2 (a.k.a. SC35), primarily regulate pre-mRNA splicing. SRSFs harbor RNA recognition motifs (RRMs) followed by C-terminal RS domains (RSDs) enriched in Arg-Ser dipeptides (27). CPSF6 likewise harbors an N-terminal RRM, which binds its alternative polyadenylation partner CPSF5 (28) to form the heterotetrameric cleavage factor I mammalian (CFIm) complex; CFIm can alternatively be composed of a heterotetramer of CPSF7 and CPSF5 (26,29). CPSF6 also harbors a proline-rich domain (PRD) with FG motif that binds HIV-1 capsid (30–32) and a C-terminal mixed-charge domain (MCD) enriched in Arg-(Asp/Glu) repeats (33,34) (Supplementary Figure S1A). The β-karyopherin transportin 3 (TNPO3) engages phosphorylated RSDs and the MCD to mediate SRSF1/2 and CPSF6 nuclear import, respectively (35–37).

CPSF6 at baseline displays some preference for paraspeckles (28) and nuclear speckles (NSs) (38,39). Upon HIV-1 infection, CPSF6 reorganizes into comparatively large capsid-dependent puncta (22) that co-localize with NSs (9,40–42). Recent cell-based fluorescence recovery after photobleaching (FRAP) and sensitivity to comparatively harsh treatments such as 1,6-hexanediol (a reagent commonly used to disperse condensates) and hypertonicity revealed that HIV-induced CPSF6 puncta display properties of biomolecular condensates (43,44). The mechanisms underlying the formation of these condensates, as well as their relevance to post-nuclear steps of HIV-1 replication, remain unclear.

Numerous subcellular structures, including paraspeckles (45) and NSs (46), are biomolecular condensates (a.k.a. membraneless organelles) that leverage LLPS activities of constitutive components for their assembly and maintenance [reviewed in refs (47,48)]. Intrinsically disordered regions (IDRs) of proteins can template multivalent interactions that drive LLPS activity (47,48) and CPSF6 contains two IDRs: the PRD, which harbors within it a prion-like domain (PrLD) (32), and the MCD (24,34,49) (Supplementary Figure S1A). Although isolated PRD and MCD domains have been shown to phase separate in vitro (24,34), their contributions to overall CPSF6 LLPS activity remain unclear (24,49). To address this as well as the role of LLPS activity in HIV-1 infection (capsid-binding, nuclear penetration, CPSF6 puncta formation, integration targeting), we have assessed CPSF6 mutant activities in vitro and in human cells. Our work establishes the MCD as an important driver of post-nuclear HIV-1 incursion for integration into speckle-proximal chromatin. Although located downstream from the known capsid-binding determinants (32), the MCD mediated LLPS-dependent, higher-order capsid interactions and was required to co-aggregate CPSF6 with capsid-like particles (CLPs) in vitro. At the same time, our work downplays a critical role for macroscale CPSF6 puncta formation in HIV-1 integration targeting.

Materials and methods

Reagents

This was a highly collaborative study, with experiments conducted across multiple laboratories. Similar reagents were accordingly oftentimes procured from different providers.

Mass standards for size exclusion chromatography (SEC) were from Bio-Rad (catalog# 1511901). The following reagents were procured from Thermo Fisher: mouse monoclonal antibody (MAb) anti-hemagglutinin (HA)-Alexa Fluor 488 (catalog# A-21287), rabbit polyclonal antibody (PAb) anti-SRRM2 (catalog# PA5-66827), goat anti-mouse-Alexa Fluor 488 (catalog# A11029), goat anti-mouse-Alexa Fluor 568 (catalog# A11031), goat anti-rabbit-Alexa Fluor 568 (catalog# A11036), goat anti-rabbit-Alexa Fluor 568 (catalog# A11011), goat anti-rabbit-Alexa Fluor 647 (catalog# A21245), λ protein phosphatase (λPP; catalog# 50-811-856), isopropyl β-d-1-thiogalactopyranoside (IPTG; catalog# 50-121-7426), Dulbecco's modified Eagle's media (DMEM; catalog# 11965084), fetal bovine serum (FBS; catalog# 10437028), trypsin-EDTA (catalog# 15400054), Tris(2-carboxyethyl)phosphine (TCEP; catalog# 20491), Hoechst 33342 (catalog# PI62249), Hoechst 34580 (catalog# H21486), TaqMan Gene Expression Master Mix (catalog# 4369016), Power SYBR Green PCR Mastermix kit (catalog# 4367659), 4,6-diamidino-2-phenylindole (DAPI; catalog# 62248), Zenon mouse IgG1 labeling kits (catalog# Z25008) and bovine serum albumin (BSA; catalog# 23209).

The following reagents were procured from Millipore Sigma: horseradish peroxidase (HRP)-mouse MAb anti-β-actin (catalog# 3854-200UL), rabbit MAb anti-HA (catalog# SAB5600116), mouse MAb anti-SC35 (catalog# 04-1550), rabbit anti-SON (catalog# HPA023535-100UL), polyethylene glycol (PEG)-3350 (catalog# P4338-500G), 1,6-hexanediol (catalog# 240117-50G), inositol hexakisphosphate (IP6; catalog# 407125–50MG), cOmplete EDTA-free protease inhibitor (catalog# 11836170001), Phosstop phosphatase inhibitor (catalog# 04906837001), dimethyl sulfoxide (catalog# D2650), PF74 (catalog# SML0835), poly-L-lysine (catalog# P4707), saponin (catalog# 558255), phosphate buffered saline–Tween 20 (PBST; catalog# P3563) and bovine thyroglobulin (catalog# 609310).

Abcam supplied the following antibodies: rabbit MAb anti-CPSF6 (catalog# ab175237), rabbit PAb anti-CPSF6 (catalog# ab99347), rabbit PAb anti-SON (catalog# ab121759), rabbit PAb anti-SRRM2 (catalog# ab122719), mouse MAb anti-SC35 (catalog# ab11826), rabbit PAb anti-lamin B1 (catalog# ab16048), goat anti-rabbit-Alexa Fluor 488 (catalog# ab150077), goat anti-mouse-Alexa Fluor 594 (catalog# ab150116) and donkey anti-mouse-Alexa Fluor 405 (catalog# ab175658). The following mouse MAbs were procured from Santa Cruz: anti-CPSF6 (catalog# sc-376228) and anti-SON (catalog# sc-398508). Remaining antibodies were procured from Dako (goat anti-rabbit-HRP; catalog# P0448), Cell Signaling Technology (mouse MAb anti-HA-HRP; catalog# 2999S), BD Biosciences (mouse MAb anti-SC35; catalog# 556363), Rockland (goat anti-rabbit-Cy5; catalog# 611-110-122), SouthernBiotech (goat anti-mouse-Cy5; catalog# 1031-15) and BEI Resources (mouse MAb anti-p24; catalog# ARP-4121). HIV-1 p24 ELISA kits were from Advance Bioscience Laboratories (catalog# 5447) or Xpress Bio (catalog# XB-1000).

The following reagents were procured from Life Technologies: human rhinovirus 3C (HRV3C) protease (catalog# 88947), lipofectamine RNAiMax (catalog# 13778150) and TURBO DNase (catalog# AM2239). The following chromatography columns were procured from Cytiva: HiTrap Q FF (1 ml, catalog # 17505301; 5 ml, catalog# 17515601), HiTrap Heparin (1 ml, catalog# 17040601; 5 ml, catalog# 17040703), HisTrap 5 ml (catalog# 17524802), HiTrap SP Sepharose FF 5 ml (catalog# 17515701), MBPTrap 1 ml (catalog# 28918778), Superose 6 10/300 (catalog# 29091596) and Superose 6 16/600 (catalog# 29323952).

The following products were procured from Corning: Terrific Broth (catalog# 46-055-CM), DMEM (catalog# 15-013-CV), DMEM lacking phenol red (catalog# 17-205-CV), FBS USDA approved Origin Heat Inactivated (catalog# 35-011-CV), penicillin streptomycin solution 100x (catalog# 30-002-CI), phosphate-buffered saline (PBS; catalog# 21-040-CV), Dulbecco's PBS (dPBS) with Ca and Mg (catalog# 21-030-CV) and dPBS without Ca and Mg (catalog# 21-031-CV).

Terrific Broth (catalog# DF0438-17) and Luria Broth (catalog# DF0446-07-05) were procured from Fisher Scientific. Paraformaldehyde (PFA) was procured from Electron Microscopy Sciences (catalog# 1570-S) or Boston Bioproducts (catalog# BM-155). Phos-tag gels were procured from Wako Diagnostics (catalog# 192-18001), Roche supplied phenylmethylsulfonyl fluoride (PMSF; catalog# 11359061001) and Refeyn supplied ready-to-use carrier slides (catalog# MP-CON-21009). Microscopy-based glassware included #1.5 coverslip (Warner Instruments catalog# 64-0712), high precision #1.5H coverslip (Electron Microscopy Services catalog# 71861-053), 8-well chambered slide (Cellvis catalog# C8-1.5H-N), 18 well chambered slide (Cellvis catalog# C18-1.5H) and μ-Slide angiogenesis (Ibidi catalog# 81506). Transfection reagents were procured from SignaGen Laboratories (PolyJet; catalog# SL100688), Polyplus (jetOPTIMUS; catalog# 101000051), Qiagen (Effectene; catalog# 301427) or Takara Bio (CalPhos; catalog# 631312). Efavirenz (EFV) was from BEI Resources (catalog# HRP-4624), Triton X-100 was from VWR (catalog# M143), 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS) was from G Biosciences (catalog# DG096), DAPI was from Cell Signaling Technology (catalog# 4083S) and BSA fraction V was from Research Products International (catalog# A30075-25).

All restriction endonucleases as well as T4 DNA ligase (catalog# M0202L) were procured from New England Biolabs. NEBuilder HIFI DNA assembly Master Mix was also from New England Biolabs (catalog# E2621L). Mycoplasma detection kit was from Lonza (catalog# LT07-218) and Quick-DNA microprep kit was from Zymo Research (catalog# D3021). The following short-interfering RNAs (siRNAs) were procured from Horizon Discovery: non-targeting (NT; UGGUUUACAUGUCGACUAA), CPSF6 (GAAUUGAGUCCAAGUCUU), SON #13 (GGAUAAGGCUCAAUUACUU), SON #14 (UCCUGAUCCCUAUAGGUUA), SRRM2 #15 (CUCACUAGUUCAAAGUUGC) and SRRM2 #18 (GGAUCACCUUUAGAAUUUA). Synthetic DNA fragments/oligonucleotides were procured from Integrated DNA Technologies, GenScript, or Twist Bioscience. Supplementary Table S2 lists all of the DNA oligonucleotides used in the study.

Biological resources

The following human cell lines were procured from ATCC: HEK293T (catalog# CRL-3216), HeLa (catalog# CCL-2) and U2OS (catalog# HTB-96). GHOST cells were from BEI Resources (catalog# ARP-3679) and HEK293T-derived CPSF6 knockout (CKO) cells (clone B8) were described previously (19).

This work used the following strains of Escherichia coli: BL21(DE3) (New England Biolabs catalog# C2527H), DH5α (Life Technologies catalog# 18265017) and JM109 (Zymo Research catalog# T3003). Previously described or procured plasmid DNAs included: pIRES2-EGFP, pCG-VSV-G (50), pNLX.Luc.R-.ΔAvrII (51), pHIVeGFPΔEnv, pVpr-INmNG (9), pNL4-3.Luc.R-E- (52), pMD2.G (Addgene catalog# 12259), pcDNA3.1 (Thermo Fisher catalog# V79020), pCDF-CLK1 (36), pETDUET-AS13 (17), pMAL-c5x-MMTV-IN (53), pEGFP-C1 (NovoPro catalog# V012024), pET-11a (Novagen catalog# 69436), pACYCDuet-1 (Novagen catalog# 71147) and pET-15b (Novagen catalog# 69661). See Supplementary Table S3 for a list of all plasmid DNAs used in this study.

Computer code resources

The study used the following computational software: IUPred2 (https://iupred2a.elte.hu/), PrDOS (https://prdos.hgc.jp/cgi-bin/top.cgi), PLAAC (http://plaac.wi.mit.edu/), ImageJ/Fiji (http://imagej.net/ij/), GraphPad Prism 10 (http://www.graphpad.com), LASX software (Leica Microsystems), ICY image analysis (http://icy.bioimageanalysis.org), R package mixdist (https://cran.r-project.org/web/packaes/mixdist/index.html), BWA-MEM (https://github.com/lh3/bwa), Refeyn DiscoverMP (https://www.refeyn.com) and Nikon Elements AR 5.2 Acquisition (https://www.microscope.healthcare.nikon.com).

Plasmid DNA constructions

CPSF6 constructs for expression in mammalian cells were fused with N-terminal HA tags, with or without the heterologous c-Myc nuclear localization sequence (NLS) (54). DNA fragments encoding the tags were PCR-amplified using primers AE8141/AE8142 (HA tag) or AE8582/AE8583 (HA with c-Myc NLS) and ligated with XhoI/PstI-digested pIRES2-EGFP. Previously described wildtype (WT) and mutant CPSF6 expression constructs ΔCPSF5b, F284A and ΔMCD (37) were amplified using primers AE8585/AE7106 (WT, ΔCPSF5b, F284A) or AE8585/AE8584 (ΔMCD), and the resulting amplicons were assembled with PstI/BamHI-digested pIRES2-EGFP. Synthetic DNAs for CPSF6 variants ΔRRM, SRSF1, SRSF2, SNRNP70, CPSF7, CPSF7+6, and RS/RD homopolymers were similarly assembled with PstI/BamHI-digested pIRES2-EGFP. N-MCD and N-4Glu PCR-amplified using AE8103/AE7480 primers from plasmids pIRES2-EGFP[551] and pIRES2-EGFP[4Glu] (37), respectively, were assembled with PstI-digested pIRES2-EGFP-HA-CPSF6[481] (37). DNA inserts for CypA-related constructs synthesized at GenScript were PCR-amplified using primers AE7042/AE8422 (^cNLSCypA) or AE7042/AE7106 (CypA-MCD and RRM-CypA-MCD) and assembled with NheI/BamHI-digested pIRES2-EGFP. For the pHA-CPSF6 plasmid series, open reading frames from corresponding pIRES2-EGFP constructs were PCR-amplified using primers AE7042/AE7043 and inserted into NheI/BamHI-digested pEGFP-C1, which removed the gene for EGFP.

Synthetic WT and A14C/E45C HIV-1 capsid protein (CA) coding sequences for bacterial expression derived from the HIV-1_NL4-3gag gene were assembled with NdeI/BamHI-digested pET11a. All constructs for bacterial CPSF6 expression were derived from pACYCDuet-1, which contains a p15a origin of replication and two multiple cloning sites (MCSs). We initially modified pACYCDuet-1 to harbor strong transcription terminators downstream from both MCS1 and MCS2. A synthetic fragment containing terminator sequences rrnB T1 and rrnB T2 (55) was assembled with EcoNI/AvrII-digested pACYCDuet-1. Next, rrnB T1 and rrnB T2 containing sequences PCR-amplified using primers GB101 and GB102 were assembled with AvrII-linearized pACYCDuet-1. The resulting plasmid was assembled with a PCR amplicon containing rrnB T2 and T7 terminators that was made using primers GB103 and GB104 following HindIII digestion. MCS2 was further modified by introducing the rhamnose-inducible rhaB promoter in place of the resident T7 promoter, and a modified MCS sequence lacking the lac operator. For this, AflII/PacI-digested plasmid was assembled with a synthetic fragment that was PCR amplified using primers GB169 and GB170.

DNA containing the maltose binding protein (MBP) coding sequence was PCR-amplified from pMAL-c5x-MMTV-IN using AE7713/AE7714 primers and assembled into NdeI/BamHI-digested pET15b. Following PCR with GB283/GB284 primers, DNA encoding His₆-MBP followed by the cleavage site for HRV3C protease was assembled with the pACYCDuet-1 derivative described above following digestion with NcoI and SacI. Finally, to create pGB12b, an NdeI site located between the His₆ and MBP coding regions was removed by primer-based (AE8278/AE8279) sequence modification and assembling the amplified fragment with NdeI-digested plasmid.

CDC like kinase 1 (CLK1) sequences PCR amplified from pCDF-CLK1 using primers AE8280/AE8281 were assembled with NdeI-linearized pGB12b to create pGB12b-CLK1. CPSF6 containing fragments were introduced into the SacI site of pGB12b-CLK1. Because SacI cut on either side of the HRV3C coding sequence, the HRV3C sequence was incorporated into PCR amplified fragments before HiFi DNA-mediated assembly. CPSF6 sequences codon optimized for expression in E. coli were initially procured from GenScript for the full-length 551-residue isoform. Fragments for the various WT and mutant CPSF6 forms were subsequently amplified by one or two sets of PCR reactions. WT was first amplified using primers AE8282/AE8283; plasmid-compatible sequences for assembly were then added via PCR using primers AE8270/AE8285. For ΔMCD, first round sequences were amplified using primers AE8282/AE8315, and plasmid assembly compatible ends were incorporated via primers AE8270/AE8316. Codon-optimized DNA fragments for ΔRRM and ΔPrLD were synthesized by Twist Biosciences. Following PCR amplification with primers AE8580/AE8581, these were assembled with SacI-digested pGB12b-CLK1.

The sequences of all PCR-amplified or de novo-synthesized regions of plasmid DNAs were sequence-verified by Sanger sequencing at GENEWIZ or whole-plasmid sequencing at Plasmidsaurus.

Expression and purification of recombinant proteins

WT and A14C/E45C CA proteins, which were expressed in E. coli strain BL21(DE3), were purified essentially as previously described (56). Briefly, transformed cells in Luria Broth–100 μg/ml ampicillin were induced in mid-log phase with 0.5 mM IPTG and propagated overnight at 20°C. Cells harvested by centrifugation were frozen at -80°C. Frozen pellets were resuspended in 50 mM Tris–HCl pH 8.0, 50 mM NaCl and 10 mM dithiothreitol (DTT) supplemented with cOmplete EDTA-free protease inhibitor tablets. Resuspended cells were homogenized with a Dounce homogenizer and sonicated on ice in 5 sec bursts interdigitated with 30 sec rests. The total duration of sonication was 10 min. Cellular debris was cleared by centrifugation at 50 000 × g for 30 min and ammonium sulfate was added to the supernatant to 25% saturation. Precipitate pelleted by centrifugation at 10 000 × g for 30 min was resuspended in 50 mM MOPS pH 6.8–10 mM DTT, dialyzed against the same buffer and applied to a 5 ml HiTrap SP Sepharose FF column. The column was developed by a linear gradient from 0 mM to 1 M NaCl in 50 mM MOPS pH 6.8–10 mM DTT. Peak fractions were pooled, dialyzed against 50 mM Tris–HCl pH 8.0, 50 mM NaCl, 10 mM DTT and applied to a 5 ml HiTrap Q FF column. CA collected in the flow-through was concentrated by ultrafiltration to 20-30 mg/ml, flash-frozen in liquid nitrogen and stored at −80°C. His₁₂-mCherry expressed in bacteria using pETDUET-AS13 was purified as previously described (17).

CPSF6 expression constructs propagated overnight at 37°C in BL21(DE3) in Luria Broth–30 μg/ml chloramphenicol were subcultured the following morning in Terrific Broth. The cultures were grown at 37°C until mid-log phase, when they were moved to orbital shakers at 15°C. The cultures were allowed to cool for ∼45 min at the reduced temperature and protein production proceeded overnight following induction with 0.5 mM IPTG. Cells harvested by centrifugation at 5000 rpm at 4°C in a Beckman JS-5.2 rotor were resuspended in PBS–10% (v/v) glycerol, recentrifuged, and frozen at −80°C. All lysis and purification steps were carried out on ice or at 4°C.

Cell pellets were thawed in 50 mM Tris–HCl pH 8.0, 150 mM NaCl, 5 mM CHAPS, 1 mM TCEP and 10% (v/v) glycerol supplemented to contain 1.5 cOmplete EDTA-free protease inhibitor cocktail tablets per liter of bacterial culture and 1 mM PMSF. The cell suspension was homogenized and sonicated as described above. The lysate was centrifuged at 50 000 × g for 30 min and the supernatant was applied to a 5 ml HisTrap column equilibrated with 50 mM Tris–HCl pH 8.0, 50 mM NaCl, 20 mM imidazole, 1 mM TCEP, 10% (v/v) glycerol. After loading, the column was washed with equilibration buffer until baseline stabilization. Proteins were eluted with a 10 column volume (CV) linear gradient of 20 mM to 300 mM imidazole in Tris–HCl pH 8.0, 50 mM NaCl, 1 mM TCEP, 10% (v/v) glycerol. Peak fractions identified by denaturing polyacrylamide gel electrophoresis were pooled and digested with HRV3C protease at ∼10 U/mg protein at 4°C for 2 days. The material was applied to two tandem 1 ml HiTrap Heparin columns or a 1 ml MBPTrap column connected in tandem to two 1 ml HiTrap Q FF columns equilibrated in 50 mM Tris–HCl pH 8.0, 50 mM NaCl, 1 mM TCEP, 10% (v/v) glycerol. The tandem columns were washed in the same buffer until baseline stabilization. In instances when an MBPTrap column was used, the MBPTrap column was removed after washing. Proteins were eluted using a 10 CV linear gradient of 50–500 mM NaCl in Tris–HCl pH 8.0, 1 mM TCEP, 10% glycerol (v/v). Peak fractions were pooled and injected onto a calibrated Superose 6 16/600 gel filtration column equilibrated in 50 mM Tris–HCl pH 8.0, 50 mM NaCl, 1 mM TCEP, 10% (v/v) glycerol. Monomeric CPSF6 species eluted at ∼60–65% of total CV. Pooled P2 column fractions were applied to an equilibrated 1 ml HiTrap Heparin or HiTrap Q FF column and concentrated via single step elution using 50 mM Tris–HCl pH 8.0, 500 mM NaCl, 1 mM TCEP, 10% (v/v) glycerol. Eluates were dialyzed against 50 mM Tris–HCl pH 8.0, 150 mM NaCl, 1 mM TCEP, 10% (v/v) glycerol, flash frozen in liquid nitrogen, and stored at −80°C. Protein concentrations were determined by A₂₈₀. Mutant protein concentrations were subsequently normalized to WT CPSF6 by immunoblotting 100 ng of each (based on A₂₈₀) using α-CPSF6 antibody (Abcam ab175237) followed by HRP-conjugated goat anti-rabbit secondary antibody (Dako P0448) (Supplementary Figure S1C). Relative band intensities were quantified using ImageJ software (57) and apparent protein concentrations were adjusted accordingly for downstream LLPS and capsid binding assays. The sequences of all recombinant proteins are listed in Supplementary Table S4.

Percent protein purity (2 μg WT or mutant CPSF6) was determined via quantifying Coomassie-stained images using the ImageJ Densitometry Tool. Protein bands were translated to peaks on a plot, from which peak areas were determined. The area of the peak corresponding to the protein of interest was divided by the sum areas of all detectable peaks along the entire gel lane. This value was multiplied by 100 to yield %-purity.

Recombinant CPSF6 mass assessment by mass photometry

Measurements were made on a Refeyn TwoMP mass photometer using ready-to-use carrier slides at the Center for Macromolecular Interactions at Harvard Medical School. Data was collected using the Refeyn AcquireMP software and analyzed using the Refeyn DiscoverMP software. For instrument calibration, a 10x solution of BSA + bovine thyroglobulin was prepared from serial dilutions of a 1000× stock in CPSF6 storage buffer [50 mM Tris–HCl pH 8.0, 150 mM NaCl, 1 mM TCEP, 10% (v/v) glycerol]. The instrument was auto-focused on a 17 μl droplet of CPSF6 storage buffer, and 3 μl of the 10× calibration mix was added to the droplet. Contrast data was collected for 1 min. A calibration curve was created by correlating the observed ratiometric contrast of BSA monomer, BSA dimer and thyroglobulin with their known molecular weights. CPSF6 samples were prepared by serially diluting stock solutions to 50 nM concentration in CPSF6 storage buffer. The TwoMP instrument was autofocused on a 15 μl droplet of CPSF6 storage buffer and 5 μl of 50 nM CPSF6 was mixed with the droplet. Data was collected for 1 min. Calibrated event data for all samples was fit to a two Gaussian mixture distribution using the R package mixdist.

LLPS assays with recombinant CPSF6 proteins

For kinetic measurements, proteins thawed on ice were clarified by centrifugation for 10 min at 20 000 × g and 4°C and transferred to a new tube. Reactions (50 mM Tris–HCl pH 8.0–150 mM NaCl at room temperature) were conducted in 96-well plates using a Bio-Rad xMark Microplate Spectrophotometer. Stock solution of 50% (w/v) PEG-3350 was prepared in reaction buffer and used at a working concentration of 10% (w/v). Reaction buffer, CPSF6 protein (10 μM final), and 1,6-hexanediol [2.5% (w/v), when used] were combined prior to the addition of PEG-3350 or the equivalent reaction buffer control volume. LLPS reactions were initiated by the addition of PEG-3350 (or mock) to the other pre-mixed reaction components. A multi-channel pipet was used to initiate all reactions in a given column of the 96-well plate simultaneously. A₃₅₀ measurements were taken every 30 s. During this 30 s delay, the tray compartment of the instrument was opened, and the next column of reactions was initiated. This process occupied nearly all of the 30 s dead time. The ∼30 s offset in initiation times was corrected in downstream data processing. Data was collected until all initiated columns had been followed for at least 30 min post-initiation. For all conditions, measurements were replicated at least 3 times. Statistical significance between PEG ± 1,6-hexanediol reactions was assessed using a two-tailed unpaired Student's t-test at 30 min post-initiation.

Micrographs of LLPS reactions were obtained as follows. A mixture of CPSF6 protein and 10% PEG-3350 in reaction buffer (50 mM Tris–HCl, pH 8.0–150 mM NaCl) was incubated for 30 min at room temperature. Following a split of the mixture and treatment with or without 10% (w/v) 1,6-hexanediol for 30 min at room temperature, A₃₅₀ was measured in a 384-well plate by Bio-Rad xMark Microplate Spectrophotometer. Following re-suspension by pipetting the entire solution, the mixture was diluted 20-fold into PBS or PEG-3350-containing reaction buffer and transferred to μ-Slide Angiogenesis. Micrographs were acquired by EVOS^TM M5000 microscope with 20X objective, phase contrast, 0.45NA/6.12WD (Thermo Fisher).

Propagation of human cells and viruses

HEK293T, HeLa, U2OS, GHOST and HEK293T-derived CKO cells (clone B8) (19) were cultured in DMEM supplemented to contain 10% FBS and 1% penicillin-streptomycin solution (100 U/ml). All cells were regularly monitored for mycoplasma and all experiments were conducted using mycoplasma-free cells.

This work used a series of single-round derivatives of HIV-1_NL4-3. For infectivity and integration site analyses, HIV-Luc was produced by co-transfecting HEK293T cells (∼4 × 10⁵ cells/ml plated the prior day in 15 cm dishes) with 30 μg pNLX.Luc.R-.ΔAvrII and 5 μg pCG-VSV-G using PolyJet. At 48 h post-transfection, virus supernatants were collected, passed through 0.45 μm filters, concentrated about 50-fold by ultracentrifugation at 53 000 × g for 2 h at 4°C and treated with TURBO DNase at 37°C for 1 h. Virus yield was assessed by p24 ELISA.

For single virus tracking [nuclear penetration; NS colocalization at 8 h post-infection (hpi)] and CPSF6 binding experiments, fluorescently-tagged HIV-1 particles pseudotyped with vesicular stomatitis virus glycoprotein G (VSV-G) were produced essentially as described previously (21). Briefly, HEK293T cells plated at 80% confluency in 6-well plates were co-transfected with envelope deleted pHIVeGFP (2 μg), VSV-G expressor pMD2.G (0.5 μg) and Vpr-IN fused to mNeonGreen (INmNG, 0.8 μg) using jetOPTIMUS reagent as recommended by the manufacturer (Polyplus). After 6 h, the medium was changed for phenol red minus DMEM complete with antibiotics and 10% FBS. Virus supernatants collected after an additional 36 h were clarified through 0.45 μm filters, aliquoted and stored at −80°C until use.

For CPSF6 puncta formation, HEK293T cells (∼1 × 10⁶ cells/ml in 10 cm dishes) were cotransfected with pNL4-3.Luc.R-E- and VSV-G expression vector at a 3:1 mass ratio. Culture supernatants were collected 48 h post-transfection, clarified by low-speed centrifugation (1000 × g, 10 min), and filtered through 0.45 μm filters. Virus yield was assessed by p24 ELISA and infectious units were determined using GHOST cells.

HIV-1 infection and integration site analyses

WT or CKO HEK293T cells were plated in 6-well plates (4-6 × 10⁵ cells/ml) and transfected with 2 μg of empty pIRES2-EGFP vector or CPSF6 expressing derivatives using Effectene or CalPhos reagents. The next day, green fluorescent protein (GFP)-positive cells were isolated by fluorescence-activated cell sorting at the Dana-Farber Cancer Institute Flow Cytometry Core in basic sorting buffer (1 mM EDTA, 25 mM HEPES, pH 8.0, 1% FBS, in PBS). Approximately 3-5 × 10⁵ sorted cells were in parallel lysed for immunoblotting or treated with HIV-Luc (0.25 pg of p24 per cell) for HIV-1 infection and integration site sequencing assays. At 48 hpi, cells were lysed, and luciferase activity was determined as previously described (58). U2OS and HeLa cells transfected with NT or SON/SRRM2 siRNAs were infected similarly, except the overall time of infection was capped at 24 h to limit side-effects associated with SON depletion.

For integration site analyses, genomic DNA isolated at 72 hpi (or 24 hpi for siRNA knockdowns) using Quick-DNA microprep kit was prepared by ligation-mediated (LM)-PCR essentially as previously described (59). Briefly, DNA was fragmented by MseI and BglII enzyme digestion, and sample-specific linkers were ligated using T4 DNA ligase. HIV-1 U5 DNA end-linker fragments amplified by nested PCR were subjected to 150 bp paired end Illumina sequencing using NovaSeq 6000 (GENEWIZ) or in-house NextSeq 2000. See Supplementary Table S2 for all LM-PCR primers.

Sequence reads trimmed to remove HIV-1 U5 and linker sequences were aligned with human genome build hg19 to identify unique integration sites. Percentiles of integration sites within genes and SPADs were determined as previously described (9,13). LAD annotations were obtained from the 4D Nucleome Data portal (Experiment Set 4DNESTAJJM3X). Genome annotation-matched random integration controls (RICs) were generated in silico to mimic wet-bench procedures of genomic DNA shearing, amplification and DNA sequencing (59). In brief, integration sites were randomly placed throughout hg19 and the genome was ‘cleaved’ at nearest upstream MseI or BglII restriction site. All fragments ≥14 bp and ≤1200 bp were kept and used to generate simulated 150 bp paired-end sequencing reads saved as R1 and R2 fasta files, which were aligned using BWA-MEM (60) to generate RIC BED files. Note that this procedure differed slightly from the one previously described in (61) that enforced a strict 100 bp limit between random integration and restriction enzyme sites and aligned these as single-end reads. The revised method used here defined random SPAD integration targeting (4.75%; see Supplementary Table S5) at a level that is comparable to the proportion of SPADs in the human genome.

Quantitative PCR for HIV-1 reverse transcription and bulk integration

To detect late reverse transcription (LRT) products, qPCR was carried out as described previously (62). Briefly, cells infected as above with HIV-Luc in the presence of 20 μM EFV or matched dimethyl sulfoxide vehicle control were washed at 2 hpi to remove virus. At indicated time points (8, 16, 24, 48 or 72 hpi), total DNA was extracted using Quick-DNA microprep kit and subjected to qPCR using the Bio-Rad C1000 Touch Thermo Cycler (CFX96 real-time system). Reactions included 25-50 ng of total DNA, primers AE2963/AE4422, probe AE2965, and TaqMan Gene expression master mix. Resulting HIV-1 DNA levels were normalized to sample-matched levels of human β-globin DNA, which was assessed by qPCR using primers AE1074/AE1075 and Power SYBR Green PCR Mastermix kit. Values obtained in the presence of EFV, which was included to control for potential plasmid carryover from HIV-Luc stocks made by DNA transfection, were subtracted from matched infections lacking EFV.

Nested HIV-Alu PCR was used to assess total levels of HIV-1 integration essentially as previously described (62). DNA isolated as above from cells at 16 hpi was amplified in the first PCR round using primers AE3014/AE1066. DNA from uninfected cells was included as a negative control. First round reaction products diluted 1:100 into second round qPCRs were amplified using primers AE3013/AE990 and TaqMan probe AE995. Values obtained from first round reactions that omitted Alu-specific AE1066 primer were subtracted from experimental C_t values, and these corrected values were normalized to the levels of human β-globin DNA as described above.

Western immunoblotting

Immunoblotting was performed as previously described (37). In brief, cells were lysed with buffer (50 mM Tris–HCl pH8.0, 150 mM NaCl, 1% IGEPAL CA-630, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate) supplemented with 1× cOmplete EDTA-free protease inhibitor and 1× Phosstop phosphatase inhibitor. Antibodies and working dilution factors were as follows: HRP-conjugated α-HA tag (Cell signaling technology 2999S; 1:5000), rabbit α-CPSF6 (Abcam ab175237; 1:10 000), HRP-conjugated α-actin (Millipore Sigma A3854-200UL; 1:50 000), rabbit α-SON (Millipore Sigma HPA023535-100UL; 1:1000 or Abcam ab121759; 1:1000), rabbit α-SRRM2 (Thermo Fisher PA5-66827; 1:1000 or Abcam ab122719; 1:1000) and HRP-conjugated α-rabbit IgG secondary antibody (Dako P0448; 1:10 000).

RNA interference-based depletion of targeted gene products

U2OS or HeLa cells at ∼50% confluency in 6-well plates were transfected with 20 nM total siRNA for NT control or single gene knockdowns, or 10 nM each of SON#13 and SRRM2#15 for double knockdown, using Lipofectamine RNAiMax. After 1–2 days, cells were trypsinized and subjected to downstream assays (western blotting, immunofluorescence, HIV-Luc infection, integration site sequencing).

Antigen detection by immunofluorescence-based assays

For verification of SON and SRRM2 knockdown, antibodies and working dilution factors were as follows: mouse α-SON (Santa Cruz sc-398508; 1:500), rabbit α-SRRM2 (Thermo Fisher PA5-66827; 1:500), rabbit α-CPSF6 (Abcam ab175237; 1:3000), secondary goat anti-rabbit-Alexa Fluor 488 (Abcam ab150077; 1:1000), secondary goat anti-mouse-Alexa Fluor 594 (Abcam ab150116; 1:1000) and mouse α-SC35 (Abcam ab11826; 1:3000). Nuclei were counter-stained with DAPI. Following siRNA transfection for 48 h, cells were trypsinized and attached onto glass coverslip for 18 h prior to fixation for immunostaining.

To assess nuclear penetration and NS colocalization at 8 hpi, CKO cells (∼800 000 cells/well) seeded in a 6-well plate were transfected with 2 μg empty pcDNA3.1 vector or indicated HA-tagged CPSF6 expression plasmid using jetOPTIMUS transfection reagent. After 6 h, cells were detached by trypsin and plated (∼50 000 cells/well) into a 8-well chamber slide in phenol red minus DMEM complete with antibiotics and 10% FBS. After overnight incubation, the cells were infected with INmNG-tagged HIV-1 at multiplicity of infection (MOI) 0.5-1 by spinoculation at 1450 × g for 30 min at 16°C. Cells were fixed at 8 hpi with 2% PFA for 5 min at room temperature, permeabilized with 0.1% Triton X-100 and blocked with 3% BSA in PBS for 1 h at room temperature. Fixed cells were incubated with 1:1000 dilution of rabbit MAb anti-HA (Millipore Sigma SAB5600116) and mouse MAb anti-SC35 (BD Biosciences 556363) overnight at 4°C. The following day, slides were washed twice with PBST and incubated with 1:2000 dilution of goat anti-rabbit-Alexa Fluor 568 (Thermo Fisher A11011) and goat anti-mouse-Cy5 (SouthernBiotech 1031-15) for 1 h at room temperature. Unbound antibodies were removed by three successive washes with PBST, and nuclei stained with Hoechst 33342 for 5 min at room temperature were washed twice with PBST and maintained in 200 μl dPBS per well until confocal imaging.

3D confocal imaging of HIV-1 nuclear penetration and NS colocalization, as well as 2D-imaging of CPSF6 binding to native HIV-1 cores in vitro (see below), was carried out on a Leica SP8 laser scanning confocal microscope using a C-Apo 63x/1.4NA oil-immersion objective. Adaptive focus control (Leica Microsystems) was utilized to maintain focus during Z-stack data collection and to correct for axial drift in 2D images. Tile-scanning was employed to image random neighboring (4 × 4) fields of view. Images were collected at 512 × 512 frame size, 120 nm pixel sizes (2x digital zoom) with 1.54 μs pixel dwell times, 2-line averaging by using 405, 488, 561 and 633 nm laser lines to excite the fluorophores with respective emissions collected between 420-480 nm (Hoechst or Alexa Fluor 405), 502-560 nm (INmNG), 572-630 nm (Alexa Fluor 568) and 645-700 nm (Cy5) using GaSP-HyD and PMT detectors. For cell-based assays, 3D images collected from minimally 30 nuclei per experiment were imaged and analyzed. 3D images were collected by acquiring 15 – 20 Z-stacks spaced by 0.5 μm apart. 2D and 3D-image series were later processed off-line using ICY image analysis software.

For image analysis, 3D z-stack images were analyzed using an in-house script in the ICY protocols module, essentially as previously described (9). Briefly, nuclear volume in 3D was detected using the lamin intensity by the HK-means and the Connected Components plugins in ICY (63). The obtained 3D region of interest (ROI) corresponding to the nuclear volume was shrunk by 0.5 μm in X–Y–Z using an ROI-erosion plugin. HIV-1 complexes detected within the eroded ROI were considered as nuclear spots. Similarly, Cy5-labeled SC35 NS compartments were segmented in 3D and HIV-1 INmNG puncta detected inside this segmented area were defined as colocalized with NS. The average number of HIV-1 INmNG puncta detected inside the nuclear volume and colocalized with NSs of individual cells were determined in each experiment by analyzing > 30 nuclei for individual samples. In most cases, ≥5 independent experiments were conducted for each CPSF6 construct (see Supplementary Table S5); 2 experiments were conducted for CPSF6 F284A and empty-pcDNA3.1 vector controls, and 4 independent experiments were conducted for CypA-related constructs. Average values of number of nuclear HIV-1 puncta, number colocalized with NSs and their respective average %-NS colocalization were plotted as dots.

CPSF6 puncta formation at 16 hpi was assessed by immunostaining essentially as previously described (40). HEK293T cells were infected with HIV-1_NLdE-luc at MOI 50 as defined on GHOST indicator cells. At 16 hpi, HA-CPSF6 puncta and SC35-earmarked NS co-localization was determined via examination of entire z-stack images of 200 total HA-CPSF6-positive CKO cells. For each condition, the percentages of the 200-examined cells that displayed NS-colocalized CPSF6 puncta versus lack of NS-colocalized puncta were determined.

To assess CPSF6 puncta formation in SON/SRRM2 knockdown cells, U2OS and HeLa cells transfected with NT, SON, SRRM2, or SON/SRRM2 siRNAs for 2 d were infected as above at MOI 50. At 16 hpi, cells were fixed with 4% PFA for 10 min and permeabilized with 0.5% Triton X-100 in PBS for 10 min. Nonspecific binding was blocked with 3% BSA for 15 min followed by incubation with anti-CPSF6 antibody at 1:200 (Santa Cruz sc-376228) and anti-SRRM2 antibody at 1:200 (Thermo Fisher PA5-66827) for 30 min. After five washes with PBS, goat anti-mouse-Alexa Fluor 488 at 1:1000 (Thermo Fisher A11029) and goat anti-rabbit-Alexa Fluor 568 at 1:1000 (Thermo Fisher A11036) were added, followed by incubation for 20 min. Cells were then stained for SON by direct staining with anti-SON at 1:200 (Santa Cruz sc-398508) and Zenon mouse IgG1 labeling kits according to the product manual. Cell nuclei were counterstained with Hoechst 34580 at 2 μg/ml. Images were captured in z-series with a digital CCD camera mounted on a Stellaris 5 widefield microscope (Leica Mycrosystems). Out-of-focus light was digitally removed using LASX deconvolution software (Leica Microsystems). CPSF6 intensities within 10 NSs per cell were quantified. For single knockdown conditions, NSs were identified via remaining SON or SRRM2 signal. NSs were identified in double SON/SRRM2 knockdown cells by exaggerating NS signal intensities via intensity scaling. Data were compiled for 10 cells (100 total NSs) per experimental condition (see Supplementary Table S5).

CA nanotube-CPSF6 binding assay

Nanotubes were assembled by diluting A14C/E45C CA to 3 mg/ml in 50 mM Tris–HCl, pH 8.0, 50 mM NaCl, 10 mM DTT and subsequent dialysis against 50 mM Tris–HCl, pH 8.0, 1 M NaCl, 10 mM DTT overnight at 4°C. A14C/E45C crosslinks were formed by dialysis against 50 mM Tris–HCl, pH 8.0, 1 M NaCl for 2-3 days followed by dialysis against 50 mM Tris–HCl, pH 8.0 with either 50 mM NaCl or 150 mM NaCl. Residual uncrosslinked CA was removed by centrifugation at 20 000 × g for 5 min followed by nanotube resuspension in 50 mM Tris–HCl, pH 8.0 with either 50 mM NaCl or 150 mM NaCl.

For CPSF6 binding assays, nanotubes based on 50 μM of unassembled A14C/E45C CA were mixed with indicated amounts of CPSF6 proteins in reaction buffer (50 mM Tris–HCl, pH 8.0–150 mM NaCl) and incubated for 30 min at room temperature. The mixtures were centrifuged at 21 000 × g for 10 min at 4°C. Total (before spin) as well as supernatant and pellet fractions after centrifugation were electrophoresed through denaturing polyacrylamide gels and visualized by Coomassie blue staining. Relative binding intensity of pelleted fractions was determined using ImageJ software (57).

Permeabilized virus binding assay

INmNG-labelled HIV-1 particles were immobilized onto a poly-l-lysine coated glass surface of a 18-well chambered slide for 30 min at 4°C. Slides were washed twice with dPBS to remove unbound viruses, and the immobilized particles were maintained in 50 μl dPBS. Purified WT or ΔMCD CPSF6 protein was diluted in 60 μl of 2× permeabilization buffer (dPBS containing 200 μg/ml saponin). Serial 2-fold dilutions, starting at 4 μM as the highest concentration, of each protein was then prepared. Each 2× protein solution (50 μl) was added to bound viral particles in 50 μl dPBS to achieve final CPSF6 concentrations. Reactions were incubated for 5 min at room temperature to allow saponin mediated lysis of virus membranes and protein binding to exposed HIV-1 cores. Following one wash with dPBS, the samples were immediately fixed in 2% PFA for 5 min at room temperature. Fixed samples were washed twice with dPBS, additionally permeabilized with 0.1% Triton X-100, and incubated with 3% BSA for 1 h at room temperature. Immunostaining for CPSF6 and CA/p24 was performed with primary anti-CPSF6 antibody (Abcam ab99347, at 1:1000 dilution) and anti-HIV-1 p24 antibody (BEI Resources ARP-4121, at 1:2000 dilution) overnight at 4°C. The following day, slides were washed twice with PBST and incubated with goat-anti-rabbit-Cy5 (Rockland 611-110-122, at 1:2000 dilution) and donkey-anti-mouse-Alexa Fluor 405 (Abcam ab175658 at 1:2000 dilution) for 1 h at room temperature. Unbound antibodies were removed by three successive washes with PBST and slides were maintained in 200 μl dPBS/well until confocal imaging. In parallel experiments, binding was assessed in the presence of 1.5% 1,6-hexanediol or 10 μM PF74.

In vitro binding capacity of CPSF6 to permeabilized viral capsids was determined from 2D confocal images by first detecting immunostained CA/p24 puncta using the spot detection plugin in ICY and then by determining the associated CPSF6 immunofluorescence signals. The background fluorescence signals residing within 1-pixel radius of the CA/p24 spot was quantified and subtracted from the CA/p24-associated CPSF6 fluorescence. Final plots of CA/p24-bound CPSF6 fluorescence was derived from minimally N = 3 independent experiments, whereby >1000 cores were imaged per experiment. Single exponential fits of the final datasets resulted in apparent K_D values. CPSF6 binding in the presence of 10 μM PF74 served as negative control.

Cherry-CLP assembly and interaction with CPSF6

Cherry-CLPs were assembled as previously described (17). Briefly, WT CA at ∼20 mg/ml and 3× Tris Assembly Buffer (25 mM Tris–HCl, pH 8.0, 150 mM NaCl, 12 mM IP6, pH 7.0, 650 μM mCherry, 1 mM TCEP) were pre-warmed in a 37°C water bath for 5 min. The two were mixed and the reaction was allowed to proceed for 1–2 h at 37°C. Reaction mixtures injected onto an equilibrated (20 mM Tris–HCl, pH 8.0, 150 mM NaCl, 0.2 mM IP6, pH 7.0) Superose 6 10/300 gel filtration column were fractionated (100 μl) just to the right of void volume peak apex as described (17) for downstream experiments.

Cherry-CLPs (∼1.3 μg/ml) were diluted 100-fold into 3x Tris Assembly Buffer and then mixed at 1:1 with protein dialysis buffer [50 mM Tris–HCl pH 8.0, 150 mM NaCl, 1 mM TCEP, 10% (v/v) glycerol] with or without 9 μM CPSF6. Following ∼10 min at room temperature, the mixture was thoroughly resuspended by pipetting and spotted onto a cleaned #1.5 H coverslip. A microscopy glass slide was placed over the coverslip and the slide sandwich was sealed using clear nail polish. Widefield and phase contrast imaging was performed on a fully automated Nikon Ti2 inverted microscope outfitted with a Nikon motorized stage, Lumencore Spectra III LED fluorescence illumination light engine and a Hamamatsu ORCA-Flash4.0 V3 Digital sCMOS camera. Imaging was performed with a Nikon Pla Apo 60x/1.4 Oil Ph3 objective. Cherry-CLP signal was obtained by exciting the fluorophore with 575/25 nm excitation, Semrock LED-CFP/YFP/mCherry-3X-A-00 multi-bandpass dichroic and Semrock FF01-641/75 nm emission filter. Sequential acquisition of mCherry fluorescence and phase contrast was performed using ND acquisition in Nikon Elements AR 5.2 Acquisition Software. Images were exported as .nd2 files to ensure all metadata information was preserved. Transmitted light optics were aligned for Koehler illumination prior image acquisition to ensure even and bright illumination across the field of view. For image acquisition, 11–15 different fields (N) per specimen were captured and approximately 3000 particles per image were analyzed and averaged (see Supplementary Table S5 for complete datasets). The images were imported into Fiji (57) and split into phase and fluorescence channels. The phase channel was inverted to convert phase dark regions into bright spots to mimic the fluorescence signals. General background subtraction using a 100 pixel kernel was used to reduce the impacts of uneven illumination. To isolate punctate structures on the fluorescence images, we used a TopHat filter with an aggressive kernel of 5 to ensure that regions close to full-width at half maximum were detected for each puncta. Resultant images were thresholded based on intensity with a minimum intensity of 50 and a maximum intensity of 70 000. These kernel size and threshold values prioritized smaller structures and were chosen with a few test images and verified with a human-in-the-loop to ensure that >90% of the puncta were correctly detected. Using the generated mask, we analyzed the intensities of the background-subtracted phase and fluorescence images for puncta area and intensities for both phase and fluorescence. Because the numbers of puncta detected were very large, we extracted summary statistics for each image.

Statistical analyses

Averages and standard deviations (SDs) were calculated and plotted using Microsoft excel or GraphPad Prism 10. N values generally represented independent experimental/biological replicates. For certain microscopy-based assays, N represented the number of cells (CPSF6 puncta formation), number of NSs (CPSF6 staining intensity), or the number of fields (CPSF6–Cherry-CLP co-aggregation) analyzed. P-values for pairwise comparisons were calculated using two-tailed unpaired Student's t test unless noted otherwise. When appropriate, P-values were corrected for multiple comparisons using the procedures noted in figure and table legends. Statistical significance indicators were defined as follows: *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.

Results

CPSF6 determinants of LLPS activity and condensate stability

Microscopy-based approaches, including FRAP, were previously used to characterize LLPS activities of fluorescent-CPSF6 fusion proteins isolated from bacteria (24,34,49). SR proteins are notoriously insoluble (64) and, in some cases, phase separation was initiated via buffer exchange from denaturing conditions (8 M urea) to isotonic salt (34). We herein strove to assess activities of comparatively well-behaved, tag-free proteins. The MCD is phosphorylated in cellulo (37,49) and phosphorylation countermanded self-association properties of glutathione S-transferase-MCD protein (36). After testing several conditions, His₆-MBP-CPSF6 co-expressed with CLK1 from a low-copy replicon was purified from soluble bacterial lysates to near homogeneity (92.4% ± 5.9%; N= 6); the His₆-MBP tag was removed following initial Ni²⁺-affinity chromatography. Size exclusion chromatography (SEC) of ion exchange-purified fractions revealed two populations (P1 and P2) that appeared oligomeric as gauged versus mass standards. Pooled and concentrated P2 fractions were however assessed by mass photometry to be >80% monomeric (Supplementary Figure S1B). CPSF6 migrated as a high molecular weight smear in denaturing gels containing alkoxide-bridged dinuclear Mn²⁺ (a.k.a. Phos-tag), which reacts with phosphate groups to impede gel mobility (65). Concordantly, pretreatment with λ protein phosphatase (λPP) collapsed the high molecular weight population to a single ∼70 kDa species that mimicked the migration pattern of untreated CPSF6 in Phos-tag-free gels (Supplementary Figure S1C).

LLPS activity was monitored using conventional approaches including polyethylene glycol (PEG)-induced changes in turbidity [absorbance at 350 nm (A₃₅₀)], droplet formation (phase contrast microscopy) and counteraction by 1,6-hexanediol (66). WT CPSF6 (10 μM) incubated at room temperature in isotonic buffer failed to detectably scatter light over a 30 min time course (Figure 1A, red data points). PEG-3350 (34), by contrast, induced a rapid turbidity change (Figure 1A, blue) that was largely counteracted by including 1,6-hexanediol in the reaction mixture (Figure 1A, green; P = 0.0004). Phase contrast microscopy confirmed the presence of largely spherical condensates (dia. = 3.2 ± 0.9 μm) (Figure 1B, C). Scalar protein levels revealed ∼1 μM as the minimal condensing concentration, and that subsequent treatment with 1,6-hexanediol reduced turbidity across CPSF6 concentrations (Supplementary Figure S1D). Considered alongside prior FRAP-based characterization of mCherry-CPSF6 dynamics (24), we conclude that monomeric CPSF6 supports LLPS activity in vitro.

Figure 1. — WT and mutant CPSF6 LLPS activities. (A) Turbidity-based activities. Each protein was reacted for 30 min under the following conditions: reaction buffer without PEG-3350 (red datapoints), with PEG-3350 (blue) or with PEG-3350 + 2.5% 1,6-hexanediol (HD; green). Results are means ± standard deviation (SD) for N= 3 experiments. ***P < 0.001; **P < 0.01; ns, P > 0.05 (PEG-3350 versus PEG-3350 + HD at 30 min; two-tailed unpaired Student's t test). (B) Phase contrast micrographs of LLPS reactions were acquired following 1:2 or 1:20 dilution under the indicated conditions. Scale bars, 10 μm. (C) Droplet numbers (sum of N = 2 experiments) and diameters (μm ± SD) were enumerated for the indicated WT or mutant CPSF6 reactions following 1:20 dilution in PEG-containing buffer. ****P < 0.0001 (vs. WT); ns, P > 0.05 (Kruskal–Wallis test with Dunn's post-hoc test). (D) Numbers of condensates were enumerated from reactions diluted 1:20 in PEG-containing reaction buffer (black bars) versus PBS (grey bars). Results are means ± SD for N= 3 experiments. ****P < 0.0001; ***P < 0.001; **P < 0.01; ns, P > 0.05 (black, mutant versus WT + PEG; grey, protein specific PBS versus PEG; two-way analysis of variance (ANOVA) with Tukey's post-hoc test).

Deletion mutant constructs were expressed and purified to test the roles of the RRM, PrLD and MCD in LLPS activity. Whereas purified ΔRRM and ΔPrLD P2 fractions were ∼91% and 71% monomeric, respectively, SEC had limited capacity to resolve the separate ΔMCD populations, resulting in an approximate 1.2:1 monomer-dimer mixture in purified P2 fractions (Supplementary Figure S1B). Consistent with prior observations that most phospho-acceptor residues reside within the MCD (37), λPP treatment marginally impacted ΔMCD mobility in Phos-tag gels, while ΔRRM and ΔPrLD behaved similarly as WT CPSF6 (Supplementary Figure S1C).

Similar to WT CPSF6, PEG-3350 induced a rapid ΔRRM A₃₅₀ change that was significantly counteracted by 1,6-hexanediol, though to a lesser extent than for the WT protein. For ΔPrLD and ΔMCD, magnitudes of PEG-3350-induced turbidity changes were less pronounced compared to WT and ΔRRM, and these responses were moreover unaffected by 1,6-hexanediol (Figure 1A). Phase contrast micrographs revealed significantly fewer ΔPrLD and ΔMCD condensates, which were also smaller in size than WT and ΔRRM condensates (Figure 1B, C). Residual ΔPrLD and ΔMCD activities are consistent with prior reports of isolated CPSF6 domain (PRD and MCD) activities (24,34). To assess condensate stability, droplets were enumerated following dilution in PEG-containing buffer versus PBS. Dilution in PBS had a comparatively minor though ostensibly significant effect on WT stability (Figure 1D, P < 0.01). By contrast, ΔRRM condensates dissolved in PBS (P < 0.0001). We conclude the PrLD and MCD each make important contributions to overall CPSF6 LLPS activity under these reaction conditions. While the RRM was dispensable for condensate formation, it nevertheless contributed to condensate stability.

The MCD is critical for CPSF6 puncta formation, viral nuclear incursion and speckle-proximal integration

Isogenic WT and CPSF6 knockout (CKO) HEK293T cells (19) were used to assess CPSF6 domain roles during HIV-1 infection (Figure 2A). CPSF6 mutant protein complementation of the CKO cell integration targeting defect was %-normalized to parallel WT CPSF6 expression vector transfections (see Supplementary Table S5 for raw and normalized data). CPSF6 was initially implicated in HIV-1 biology via relocalization of a ΔMCD variant of minor isoform 2 to the cell cytoplasm, which potently restricted viral nuclear import (52,67). Cytoplasmic mislocalization of major isoform 1, which is the isoform studied here, was subsequently shown to restrict HIV-1 reverse transcription (68,69). We accordingly used HIV-1 infection as an indirect readout of CPSF6 nuclear localization; nucleocytoplasmic localization was in parallel assessed by immunofluorescence (Supplementary Figures S2–S3). Integration targeting and HIV-1 infection were assessed for all cell-based CPSF6 constructs. Based on these findings, HIV-1 nuclear penetration and co-localization with NSs at 8 hpi (9,32) and CPSF6 puncta formation at 16 hpi (40) were determined for select mutants (Figure 2A). F284A was employed as a known capsid-nonbinding mutant control (30). Due to the contribution of additional PRD residues to capsid binding (32), PrLD deletion mutants were not studied in the context of HIV-1 infection.

In addition to ΔRRM, we analyzed a 7-residue internal deletion that is defective for CPSF5 binding (ΔCPSF5b, lacking residues 116–122) (70). As expected (71), ΔCPSF5b supported robust HIV-1 integration into genes [80.0% ± 2.0% (N= 3) of WT CPSF6 activity], which corresponded to 60.4% ± 14.9% of the level at which WT CPSF6 conferred SPAD integration targeting (Figure 2B). ΔRRM supported comparable levels of gene- and SPAD-specific HIV-1 integration [69.1% and 45.4% (N= 5), respectively] (Figure 2B). With their C-terminal NLSs intact, neither ΔCPSF5b nor ΔRRM restricted HIV-1 infection (Figure 2C).

CPSF6 puncta formation was assessed at high MOI to ensure all cells were infected and to enhance detection of the macroscale structures. Under these conditions, virtually all (98%) WT CPSF6-expressing cells supported puncta formation. Virus-infected F284A-expressing cells, by contrast, appeared indistinguishable from WT-expressing cells in the absence of added virus (Figure 2D, compare F284A to mock). Whereas ΔCPSF5b formed puncta at near-WT efficiency, ∼58% of ΔRRM-expressing cells supported puncta formation. Nuclear penetration was quantified as the number of fluorescently-labeled HIV-1 particles that had migrated beyond 0.5 μm from the nuclear rim. In cells expressing WT CPSF6, the vast majority (94.9% ± 6.8%; N= 8) of nuclear-penetrated particles colocalized with NSs at 8 hpi. While ΔRRM supported the WT efficiency of HIV-1 nuclear penetration, there was a marginal ∼20% reduction in the ability of these particles to colocalize with NSs (Figure 2E, P = 0.03).

Because the MCD is the functional CPSF6 NLS (37), the heterologous c-Myc NLS (54) (cNLS) was appended onto the CPSF6 N-terminus to assess post-nuclear MCD function. ^cNLSΔMCD appeared exclusively nuclear by immunofluorescence and cNLS effectively counteracted ΔMCD-mediated restriction of HIV-1 DNA synthesis (68,69) (Figure 2C and Supplementary Figures S2–S3); cNLS did not noticeably impact WT CPSF6 function (^cNLSWT; Figure 2B–E, Supplementary Figures S2–S3 and Supplementary Table S5). Strikingly, despite harboring the complete PRD, ^cNLSΔMCD failed to support CPSF6 puncta formation (Figure 2D) and, as was the case for cells transfected with the F284A-expression vector or empty vector (EV) control, HIV-1 capsids failed to penetrate into the nuclei of ^cNLSΔMCD-expressing cells (Figure 2E, P < 0.0001 versus WT-expressing cells). Of the few particles that managed to penetrate >0.5 μm from the rim, only a smattering (6.0% versus WT; P < 0.0001) colocalized with NSs. As a consequence, HIV-1 integration in ^cNLSΔMCD-expressing cells was extensively redirected to LAD-proximal chromatin (87.7% ± 8.1% compared to CKO cells; N= 4) with concomitant > 10-fold reductions in integration into genes and SPADs (Figure 2B and Supplementary Table S5). Based on these data, we concluded ^cNLSΔMCD functionally phenocopied loss-of-capsid binding CPSF6 in the context of HIV-1 infection.

Arginine mixed-charge sequences critically underlie MCD functionality in HIV-1 infection

Based on the above findings, we next tested the ability of heterologous SR-/SR-related RSDs/MCDs to complement CPSF6 MCD function in the absence of cNLS. Two representative RSDs (from SRSF1 and SRSF2) and two MCDs [from small nuclear ribonucleoprotein U5 subunit 70 (SNRNP70) (34,72) and CPSF7 (33)] (Figure 3A) were appended onto the ΔMCD C-terminus. The resulting chimeric fusion proteins were predominantly nuclear (Supplementary Figures S2–S3) and accordingly supported HIV-1 infection levels similar to WT CPSF6 (Figure 3B and Supplementary Table S5).

Figure 3. — C-terminal mixed charges underlie CPSF6 function during HIV-1 infection. (A) Primary RSD and MCD sequences with RS and R(D/E) repeats highlighted in blue and red, respectively. Numbers to the right tabulate RS and R(D/E) percentages of the indicated sequences. Identical and physiochemically similar residues in the CPSF6-CPSF7 MCD alignment are indicated by vertical lines and colons, respectively. The 11-residue sequence incorporated into the CPSF7+6 chimer is highlighted in yellow. (B) HIV-1 infection of cells expressing indicated CPSF6 constructs normalized to WT CPSF6-expressing cells (avg ± SD for N = 4 to 9 independent experiments) alongside representative immunoblots. ****P < 0.0001; ***P < 0.001; **P < 0.01; *P < 0.05; ns, P > 0.05 (versus WT CPSF6-expressing cells; one-way ANOVA with Tukey's post-hoc test). (C) HIV-1 integration into genes and SPADs in cells expressing the indicated CPSF6 proteins. Results (avg ± SD for N = 2–9 independent experiments) were normalized to WT CPSF6-expressing cells. Color-coded P-values indicate differences vs. WT CPSF6-expressing cells. ****P < 0.0001; **P < 0.01; *P < 0.05; ns, P > 0.05 (two-tailed unpaired Student's t test with Benjamini–Hochberg procedure). (D) HIV-1 nuclear penetration and colocalization with NSs. See Figure 2E for additional details. Black asterisks are versus WT CPSF6 while red asterisks compare to ^cNLSΔMCD-expressing cells. ****P < 0.0001; ***P < 0.001; **P < 0.01; ns, P > 0.05 (two-tailed unpaired Student's t test with Benjamini–Hochberg procedure). (E) CPSF6 puncta formation as enumerated in Figure 2D.

MCDs in this context significantly outshined RSDs to support early events of HIV-1 replication. While the RSD chimeric proteins conveyed integration into SPADs at ∼12–15% of the level of WT CPSF6, the CPSF7 and SNRNP70 proteins functioned at 44% and 68% of WT CPSF6, respectively (Figure 3C). HIV-1 penetrated SRSF1-expressing nuclei ∼24% as efficiently as compared to WT-expressing cells, and 41% of these particles colocalized with NSs. Although expression of the CPSF7 chimeric protein improved these metrics to ∼43% nuclear penetration and, of these particles, ∼57% colocalization with NSs, these levels still differed significantly from corresponding WT CPSF6 activities (Figure 3D), as did the ability for CPSF7 to mediate HIV-1 integration into SPADs (P < 10^–5 vs. WT). While neither RSD chimeric protein effectively formed CPSF6 puncta, ∼2% and 72% of CPSF7 and SNRNP70-expressing cells, respectively, supported CPSF6 puncta formation (Figure 3E). As CPSF6, SNRNP70 and CPSF7 MCDs are composed of 46%, 40% and 21% Arg-Asp/Glu sequences (Figure 3A), these data raised the possibility of a threshold mixed-charge density required for post-nuclear HIV-1 function. CPSF6/CPSF7 MCD alignment revealed ∼56% amino acid identity and 70% homology considering physiochemically similar side chains. The largest gap occurred near the C-termini, with a stretch of 10 mixed-charge residues absent from CPSF7 (Figure 3A). We accordingly added ‘missing’ RDRERDREREY residues at the analogous CPSF7 position to construct the CPSF7+6 MCD chimeric protein with 31% mixed-charge content. The added sequence improved the ability of the CPSF7 MCD to convey HIV-1 integration into SPADs ∼2.2-fold, a level that was indistinguishable from the WT (96.3%; P = 0.69) (Figure 3C). At the same time, CPSF6 + 7 only marginally improved puncta forming capacity of CPSF6-expressing cells, from about 2% (for CPSF7) to 8% (Figure 3E).

We also tested artificial RS and RD homopolymers as functional mimics of RSDs and MCDs, respectively. Previously, a 20-mer unit consisting of 10 RS repeats (RS₁₀) was shown to functionally substitute for the SRSF1 RSD NLS (73) and RD₅₀ fused to GFP colocalized with NSs as efficiently as CPSF6 and SNRNP70 MCD fluorescent fusion proteins (34). For completeness, RS₁₀, RD₁₀, RS₅₀ and RD₅₀ were tested at the ΔMCD C-terminus. Because all 4 chimeric proteins restricted HIV-1 infection >2-fold, a second set of constructs with N-terminally appended cNLS was generated, which supported HIV-1 infection levels indistinguishable from WT CPSF6-expressing cells (Figure 3B; Supplementary Table S5). Strikingly, ^cNLSRD₅₀ was the only construct amongst these 8 to effectively target HIV-1 to integrate into SPADs (78.8% of WT CPSF6 activity; P = 0.049) (Figure 3C). The majority of ^cNLSRD₅₀-expressing cells (96%) supported CPSF6 puncta formation and ^cNLSRD₅₀ additionally supported WT levels of HIV-1 nuclear penetration and NS colocalization (Figure 3D, E).

To assess functional domain organization, we appended the MCD onto the ΔMCD N-terminus (N-MCD). As expected (37), N-MCD was predominantly nuclear and HIV-1, accordingly, similarly infected N-MCD and WT CPSF6-expressing cells (Figure 3B and Supplementary Figures S2–S3). Introduction of four electronegative MCD substitutions (N-4Glu) that negated TNPO3 binding (37), moreover, disrupted N-MCD nuclear localization and potently restricted HIV-1 infection (Figure 3B and Supplementary Figures S2–S3). N-MCD supported ∼29% of WT CPSF6 SPAD integration targeting activity (Figure 3C) and ∼52% nuclear HIV-1 penetrance; ∼56% of these complexes, compared to the WT, colocalized with NSs (Figure 3D). Likely owing to suboptimal NS colocalization at 8 hpi, N-MCD in overnight infections failed to effectively form CPSF6 puncta (Figure 3E). Thus, although the N-terminal MCD conferred effective N-MCD nuclear import, its N-terminal location significantly influenced downstream aspects of CPSF6 function during HIV-1 infection.

The MCD is essential for higher-order capsid binding

Having determined a critical role for the MCD in post-nuclear HIV-1 incursion, we next assessed domain influences on CPSF6-capsid binding activity under a variety of assay conditions. Oligomerized nanotubes were assembled from A14C/E45C CA which, after disulfide-mediated crosslinking, stabilizes hexameric capsid lattices in isotonic salt (74). Following incubation with host factors and centrifugation, cellular proteins that bind CA co-pellet with the nanotubes (52,74). CPSF6 protein (1 μM) was incubated with nanotubes assembled with excess (50 μM) CA. Due to the central role of the PrLD in capsid binding (30,32), it was unsurprising ΔPrLD failed to appreciably pellet under conditions where the remaining proteins pelleted similarly as the WT (Supplementary Figure S4A). We next varied CPSF6 concentration while keeping CA concentration constant, yielding CA-to-CPSF6 molar ratios that spanned from ∼12.5 (4.0 μM CPSF6) to 200 (0.25 μM CPSF6). As observed at 1.0 μM CPSF6, 0.5 μM ΔPrLD failed to appreciably pellet while the remaining proteins bound similarly as the WT (Figure 4A). At higher CPSF6 concentrations, ΔMCD additionally revealed a capsid binding defect (Figure 4A, P < 0.0001).

Figure 4. — The MCD mediates higher-order interactions with capsids. (A) Coomassie-stained gel images of CPSF6 variants (0.25–4.0 μM input protein) co-pelleting with HIV-1 capsid nanotubes. The adjacent plot shows results (avg. ± SD) of N= 3 independent binding experiments. ****P < 0.0001 (mutant versus WT; two-way ANOVA with Tukey's post-hoc test). (B) Representative micrographs of WT and ΔMCD CPSF6 binding to permeabilized HIV-1 INmNG particles (CA, blue; CPSF6, red; IN, green) in the presence or absence of 1.5% 1,6-hexanediol (HD). Scale bar, 2 μm. The lower left graph plots WT and ΔMCD concentrations (x-axes) versus fluorescence arbitrary units (a.u.); opened symbols are reactions conducted in the presence of HD. The regions of both graphs encompassing 0–250 nM protein are expanded to the right with + HD curves dashed to help distinguish from corresponding minus-HD curves. Apparent K_D’s, calculated from the 0–250 nM data, are noted for the different conditions of CPSF6 binding.

Based on these observations, we next sought to investigate WT versus ΔMCD binding under more physiological conditions. Fluorescently-labelled HIV-1 particles affixed to glass were perforated to allow binding of exogenously added protein to exposed capsid structures (75). Following incubation and washing steps, capsid-bound WT and ΔMCD CPSF6 were detected via indirect immunofluorescence. Binding curves for both proteins revealed largely similar hyperbolic behavior at ≤250 nM concentrations, yielding ∼20–26 nM apparent K_D’s (Figure 4B, rightward zoomed-in graphs). Capsid binding increased an additional ∼40% as ΔMCD concentration increased to 2 μM. However, at these same conditions, WT binding increased practically 4-fold (P < 0.0001 versus ΔMCD; Supplementary Figure S4B), an increase that was largely counteracted by including 1,6-hexanediol in the binding reactions. Because ΔMCD in the 0.25–2 μM concentration range behaved similarly as WT + 1,6-hexanediol, we concluded that MCD-mediated LLPS activity underlies higher-order CPSF6-capsid interactions (Figure 4B). Reactions conducted in the presence of PF-3450074 (PF74), which competes with CPSF6 for binding to the capsid FG pocket (15), similarly inhibited WT and ΔMCD binding to permeabilized HIV-1 particles (Supplementary Figure S4B).

We next sought to visualize the effects of MCD-mediated, higher-order capsid interactions using purified reaction components. Capsid-like particles (CLPs) were fluorescently labeled via CA assembly in the presence of IP6 and excess soluble mCherry (Cherry-CLP) as described (17). Following purification by SEC, Cherry-CLPs were incubated with CPSF6 protein (WT or ΔMCD) or buffer control, and the mixtures were processed in parallel by phase contrast and fluorescence microscopy (Supplementary Figure S5). Compared to buffer-only control, ΔMCD reduced apparent particle size by ∼20%, with similar reductions in phase contrast and fluorescence intensity. By contrast, WT CPSF6 increased baseline particle size ∼1.5-fold, with concomitant 2- to 3-fold increases in fluorescence and phase contrast intensities, respectfully (Figure 5).

Figure 5. — The MCD is required for CPSF6-CLP co-aggregation *in vitro*. (A) Representative fluorescence (mCherry) and bright field (phase contrast) micrographs of Cherry-CLP–CPSF6 reactions. Fluorescence intensities were displayed as colors ranging from 45 to 1800 as shown on the top right. (B) Individual data points on the scatter plots represent an average of 3048 (± SD = 829.5) particles analyzed per field; see Supplementary Table S5 for complete datasets. Detailed pipeline for image processing and quantification is shown in Supplementary Figure S5. Mean ± SD values are tabulated at the bottom. N, number of different fields acquired per specimen; a.u., arbitrary units. ****P < 0.0001; *P < 0.05; ns, P > 0.05 (one-way ANOVA with Tukey's post-hoc test) as indicated or compared to buffer control (black asterisks).

NS depletion preferentially disrupts CPSF6 puncta formation

Our above findings with CPSF7+6 indicated that SPAD-proximal HIV-1 integration could occur in cells with reduced puncta forming capacity (Figure 3). Recent studies, moreover, have indicated that immortalized (but not transformed) epithelial cells (49), as well as primary T cells (76), harbor pre-existing CPSF6 puncta. To assess the role of puncta formation in HIV-1 integration, we depleted NSs by knocking down SON DNA and RNA binding protein (SON) and SR repetitive matrix 2 (SRRM2) using siRNAs as described (77). U2OS osteosarcoma cells were initially utilized for these experiments due to their sensitivity to siRNA-mediated knockdown (19). Preliminary experiments identified two SON (#13 and #14) and SRRM2 (#15 and #18) siRNAs that effectively depleted each or both proteins (SON#13/SRRM2#15) (Supplementary Figure S6A). Downstream experiments leveraged SON#13 and SRRM2#15 siRNAs. Results of confocal microscopy confirmed effective SON and SRRM2 depletion (Supplementary Figure S6B). While co-depletion as previously reported significantly reduced NS detection by anti-SC35 antibody (77), the CPSF6 staining pattern appeared largely unaffected by these knockdowns (Supplementary Figure S6C).

As expected (9,19), CPSF6 knockdown (Supplementary Figure S6A) countermanded HIV-1′s preference to integrate into genes and SPADs (Figure 6A). While SON depletion also reduced integration into genes and SPADs, these levels remained significantly enriched versus random calculated frequencies (Figure 6A, dashed lines). Moreover, HIV-1 in co-depleted SON + SRRM2 cells integrated into SPADs at a level that was statistically indistinguishable from the level observed in cells transfected with non-targeting (NT) control siRNA (P = 0.15; Figure 6A and Supplementary Table S5). CPSF6 puncta formation was quantified as NS-associated CPSF6 staining intensities. HIV-1 infection increased NS-associated CPSF6 in siNT-transfected cells ∼11.3-fold over the basal level observed in mock-infected cells. SON and SRRM2 depletion reduced this elevated level of CPSF6 staining by ∼44% and < 10%, respectively. Because CPSF6 staining intensity in double knockdown cells was also reduced ∼40% compared to siNT-transfected cells (Figure 6B, P < 0.01), we concluded HIV-1 integration into SPADs proceeded largely unabated in cells harboring ∼40% diminished puncta formation capacity. At the same time, the residual level of NS-associated CPSF6 staining intensity in SON/SRRM2-depleted cells remained ∼7-fold enriched compared to mock-infected cells. To assess the generalizability of these findings, we expanded the analysis to HeLa adenocarcinoma cells, which, like U2OS cells, are highly responsive to siRNA-mediated knockdowns (78). Under these conditions, HIV-1 infection resulted in an ∼10.1-fold increase in NS-associated CPSF6 staining intensity. While SRRM2 knockdown reduced CPSF6 staining by about 30%, SON knockdown yielded an approximate 73% reduction. CPSF6 staining intensity in double knockdown HeLa cells was moreover reduced to ∼19% of siNT-transfected cells (P < 10^–6), a level that was enriched < 2-fold from the basal level of NS-associated CPSF6 (Figure 6C, P > 0.05). Under these conditions, siNT- and SON#13/SRRM2#15-transfected cells supported similar profiles of HIV-1 integration targeting (Figure 6D and Supplementary Table S5; P > 0.05 for integration into genes and SPADs). Measures of bulk HIV-1 DNA metabolism (reverse transcription and integration via viral-cellular Alu sequence junctions) were similarly unaffected by SON, SRRM2, or SON/SRRM2 knockdown in U2OS or in HeLa cells (Supplementary Figure S6D, E).

Cyclophilin A-MCD-mediated HIV-1 nuclear penetration and integration targeting

To enable the penetration of HIV-1 cores beyond the nuclear rim, CPSF6 has been proposed through its FG motif to compete for capsid binding to the FG nucleoporin Nup153 (22). The FG pocket is one of several regions of host factor binding on the capsid. CypA, which regulates cytoplasmic trafficking and nuclear import during HIV-1 ingress (79,80), binds to a different region known as the CypA-binding loop [reviewed in ref. (18)]. To test if HIV-1 nuclear penetration strictly requires FG-mediated tethering of the MCD to the capsid, CPSF6 residues 441–551 containing the MCD were appended to the C-terminus of CypA (CypA-MCD). In a second iteration, CPSF6 residues 2–193 were added to the N-terminus (RRM-CypA-MCD). As a control, cNLS was incorporated at the CypA N-terminus (^cNLSCypA). All constructs were efficiently expressed, although to varying levels, and none impeded HIV-1 infection (Figure 7A).

Figure 7. — HIV-1 nuclear penetration and integration in CypA-CPSF6 fusion protein-expressing cells. (A) Levels of HIV-1 infection in cells expressing the indicated CypA-based construct, normalized to WT CPSF6-expressing cells (avg ± SD for N = 2 independent experiments) alongside representative immunoblots. The vertical dashed line indicates that the fourth lane of the immunoblot originated from a different location of the same polyacrylamide gel. *P < 0.05; ns, P > 0.05 (one-way ANOVA with Tukey's post-hoc test). (B) HIV-1 integration into genes and SPADs in cells expressing WT CPSF6 or the indicated CypA/CypA-CPSF6 protein. Results (avg ± SD for N = 2–9 independent experiments) were normalized to WT CPSF6-expressing cells. Color-coded P-values indicate differences vs. WT CPSF6-expressing cells. ****P < 0.0001; ***P < 0.001; *P < 0.05; ns, P > 0.05 (two-tailed unpaired Student's t test with Benjamini–Hochberg procedure; see Supplementary Table S5 for complete datasets. (C) HIV-1 nuclear penetration and colocalization with NSs. See Figure 2E for additional details. Black asterisks are differences versus WT CPSF6 while red asterisks compare to ^cNLSCypA-expressing cells. ****P < 0.0001; *P < 0.05; ns, P > 0.05 (two-tailed unpaired Student's t test with Benjamini–Hochberg procedure).

While cNLS localized CypA to CKO cell nuclei, ^cNLSCypA failed under these conditions to convey HIV-1 nuclear penetration and, as a consequence, NS association. Although ^cNLSCypA supported 23.6% of WT CPSF6’s ability to target HIV-1 integration into genes, it failed to target integration into SPADs (Figure 7B, C). The MCD, by contrast, conveyed comparatively impressive HIV-1 nuclear penetration, NS colocalization, and integration targeting activities to CypA. As compared to WT CPSF6, CypA-MCD supported about 81% and 88% HIV-1 nuclear penetration and NS colocalization, respectively. Under these conditions, HIV-1 integrated into genes and SPADs at 83.4% and 24.4% of the levels observed in WT CPSF6-expressing cells, respectively. RRM-CypA-MCD supported overall similar levels of HIV-1 nuclear penetration, NS colocalization and integration targeting profiles as CypA-MCD (Figure 7B, C). Likely due to the homing capacity of the isolated MCD for NSs (34), we were unable to assess puncta formation in CypA-MCD and RRM-CypA-MCD-expressing cells due to baseline predispositions for NS co-localization (Supplementary Figure S2).

Discussion

CPSF6 LLPS activity

SR-/SR-related proteins are comparatively insoluble, requiring high concentrations of RS peptides or arginine to obtain protein concentrations suitable for biochemical and biophysical measurements (64,81). Using a low-copy replicon that co-expresses CLK1, we have established a system that yields monomeric CPSF6 in isotonic buffer without the need for these added solubility cofactors. Our approach may very well be suitable to obtain tactile quantities of other SR protein superfamily members.

PRD-deleted mCherry-CPSF6 protein failed to form condensates, which initially missed a role for the MCD in CPSF6 LLPS activity (24). More recently, Liu et al. (49) concluded that the MCD was the sole driver of CPSF6 LLPS activity. Through careful analysis of purified WT and domain-deleted proteins, our results clarify that the PrLD and MCD both contribute to CPSF6 LLPS activity in vitro (Figure 1 and Supplementary Figure S1).

CPSF6 LLPS activity in capsid binding and HIV-1 infection

The replication cycles of several viruses leverage LLPS activities of macromolecules to drive biomolecular condensation. Viral proteins can template the formation of so-called replication factories to enhance nucleic acid replication, gene expression and virion assembly (82–90). The lynchpin event of retroviral replication is integration of a single linear viral DNA molecule into a cell chromosome, which would seemingly obviate the need for multiscale replication factories. Along these lines, LLPS activities of HIV-1 proteins have been reported to orchestrate internal capsid component functionality during HIV-1 ingress (91).

Recent work has indicated that HIV-induced CPSF6 puncta display properties of biomolecular condensates (43,44). Although this has led to speculation that CPSF6 LLPS activity underlies CPSF6 puncta formation during HIV-1 infection (92), such puncta colocalize with NSs, which themselves harbor numerous phase separating entities (93,94). Herein, we combined results of in vitro biochemistry and cell biology/virology to determine the role of CPSF6 LLPS activity in the early events of HIV-1 replication.

Previous work established that the affinity of the CPSF6-capsid interaction is highly context dependent. Thus, the PrLD-derived FG peptide 276-PVLFPGQPFGQPPLG bound CA hexamers ∼10-fold more effectively than it bound the isolated CA N-terminal domain (31,95,96). The mature capsid lattice further templates intermolecular PrLD interactions, increasing binding affinity an additional ∼50–100-fold from that seen with capsid hexamers (32). We determined here that WT and ΔMCD CPSF6 proteins bound permeabilized virus particles at roughly similar 20–26 nM K_Ds (Figure 4B). Moreover, at >2 μM CPSF6 concentrations, MCD-mediated LLPS activity drove additional higher-order interactions (Figures 4 and 5). Future work will focus on the structural basis of these higher order interactions.

Although our data establish a key role for MCD-mediated LLPS activity in capsid binding and HIV-1 nuclear incursion, they at the same time indicate the PrLD-capsid interaction may not leverage the inherent LLPS activity of the PRD (Figure 4B). This interpretation seems consistent with the appearance of PrLD-mediated ‘ladders’ that interdigitate between adjoining hexamers along the surface of the capsid lattice (32). Although significantly enhancing the avidity of CPSF6-capsid interactions (32), it seems possible the templating capsid honeycomb might restrict additional higher-order PrLD interactions, such as those observed for expanded β-sheets within pathological amyloid assembles [reviewed in (97)]. By contrast, we envision the MCD in the context of capsid-bound CPSF6 is unrestrained from forming higher-order, LLPS-mediated interactions. As the MCD was reportedly the sole driver of CPSF6 LLPS activity in cellulo (49), additional work is required to ascertain precise contributions of different CPSF6 domains to phase separation under different biological conditions.

Although dispensable for LLPS activity, the RRM played an important role in condensate stability in vitro and ΔRRM supported partial NS colocalization, CPSF6 puncta formation and HIV-1 integration targeting activities. These data are consistent with a prior report for enhanced RD₅₀ NS co-localization via the SRSF2 RRM (34). As RSD-RRM interactions are moreover implicated in SRSF1 phase separation (64), we would find it unsurprising if RRM-MCD interactions similarly contribute to CPSF6 condensate formation and/or stability in cellulo. The partial activities observed for the rearranged N-MCD construct indicate that RRM/MCD domain order is important for WT CPSF6 function. We suspect the globular nature of the central CypA ‘domain’ (as compared to the disordered CPSF6 PRD) restricted the ability of the RRM to enhance MCD activity in the context of the RRM-CypA-MCD construct (Figure 7).

HIV-1 DNA synthesis terminates in the nucleus (9,40,42,98) where HIV-induced CPSF6 condensates have been suggested to serve as reverse transcription hubs (42,44). However, several observations indicate that condensate formation is not critical for reverse transcription. HIV-1 DNA synthesis proceeds normally in CKO cells (19) and in primary macrophages infected with A77V CA mutant virus that is defective for CPSF6 binding (22). Moreover, ^cNLSΔMCD-expressing CKO cells supported the WT level of reverse transcription in the absence of detectable nuclear puncta (Figure 2), and DNA synthesis and integration proceeded normally in U2OS and in HeLa cells depleted for NSs and HIV-induced CPSF6 puncta (Supplementary Figure S6). Because SPAD-proximal integration was retained in cells with minimal puncta forming capacity, either through CPSF7+6 expression (Figure 3) or SON/SRRM2 depletion (Figure 6), we conclude that CPSF6 LLPS activity—as compared to macroscale condensate formation—is the more important driver of post-nuclear early replication events.

Our work fails to address the mechanism of capsid transport from the nuclear periphery for SPAD-proximal integration. Biomolecular condensates can generate capillary forces (99) and dynamic NSs display directional movement and can fuse to form larger structures (46). Possibly, CPSF6 ties into this biosynthetic pathway proximal to the NPC to leverage directional inward movement. The MCD on its own co-localizes with NSs (34) and conferred partial HIV-1 nuclear penetration and integration targeting activities to CypA (Figure 7). Recent data indicates that HIV-1 capsids can permeate the interior hydrogel of the NPC (16,17) and our results are consistent with the notion that the capsid continues to favor a phase separated environment for its downstream journey to chromatin for integration. CPSF6 regulates immunity to VSV and herpes simplex virus infection via its alternative polyadenylation activity (100,101) and innate immunity to HIV-1 via direct capsid binding (69), both of which leverage the protein's LLPS activity (Figure 4) (49). Investigating the precise role of CPSF6 LLPS activity in regulating HIV-1 evasion of cellular innate immune sensors is subject for future work.

Study limitations

Although dispensable for infection of cell lines (19,52,102), CPSF6 depletion reduced HIV-1 infection of primary resting T cells by ∼7 to 10-fold (103) and abrogated multi-cycle replication in primary macrophages (69). Despite extensive efforts, we have been unable to isolate CKO T cells (61), necessitating the use of HEK293T cells as a model system. Importantly, HIV-1 targets SPAD regions of human DNA for integration across cell types, including primary T cells and macrophages (9,10,61). Because the MCD mediated higher-order interactions with capsids in vitro (Figures 4 and 5), we would expect its role in HIV-1 nuclear incursion established here using non-lymphoid cells to readily apply to physiological target cells of HIV-1 infection.

Supplementary Material

gkae769_Supplemental_Files

gkae769_supplemental_files.zip^{(22MB, zip)}

Acknowledgements

We thank the Center for Macromolecular Interactions at Harvard Medical School for training and use of the Refeyn TwoMP mass photometer and T. U. Schwartz for critical feedback.

Contributor Information

Sooin Jang, Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA; Department of Medicine, Harvard Medical School, Boston, MA 02115, USA.

Gregory J Bedwell, Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA; Department of Medicine, Harvard Medical School, Boston, MA 02115, USA.

Satya P Singh, Institute of Molecular Biophysics, Department of Biological Sciences, Florida State University, Tallahassee, FL 32304, USA.

Hyun Jae Yu, Model Development Section, Cancer Innovation Laboratory, National Cancer Institute, Frederick, MD 21702, USA.

Bjarki Arnarson, Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA.

Parmit K Singh, Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA; Department of Medicine, Harvard Medical School, Boston, MA 02115, USA.

Rajalingam Radhakrishnan, Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA.

AidanDarian W Douglas, Institute of Molecular Biophysics, Department of Biological Sciences, Florida State University, Tallahassee, FL 32304, USA.

Zachary M Ingram, Department of Microbiology and Molecular Genetics, University of Pittsburgh School of Medicine, Pittsburgh, PA 15219, USA.

Christian Freniere, Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT 06511, USA.

Onno Akkermans, Department of Biology, Massachusetts Institute of Technology, Cambridge, MA 02139, USA.

Stefan G Sarafianos, Center for ViroScience and Cure, Laboratory of Biochemical Pharmacology, Department of Pediatrics, School of Medicine, Emory University, Atlanta, GA 30322, USA.

Zandrea Ambrose, Department of Microbiology and Molecular Genetics, University of Pittsburgh School of Medicine, Pittsburgh, PA 15219, USA.

Yong Xiong, Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT 06511, USA.

Praju V Anekal, MicRoN Core, Harvard Medical School, Boston, MA 02215, USA.

Paula Montero Llopis, MicRoN Core, Harvard Medical School, Boston, MA 02215, USA.

Vineet N KewalRamani, Model Development Section, Cancer Innovation Laboratory, National Cancer Institute, Frederick, MD 21702, USA.

Ashwanth C Francis, Institute of Molecular Biophysics, Department of Biological Sciences, Florida State University, Tallahassee, FL 32304, USA.

Alan N Engelman, Department of Cancer Immunology and Virology, Dana-Farber Cancer Institute, Boston, MA 02215, USA; Department of Medicine, Harvard Medical School, Boston, MA 02115, USA.

Data availability

Illumina FASTQ files are available from the National Center for Biotechnology Information Sequence Read Archive using accession codes PRJNA1095530 (SON/SRRM2 depletion datasets) and PRJNA1095531 (all remaining integration sites). The authors confirm that all other data supporting the findings of this study are available within the article and Supplementary Data

Supplementary data

Supplementary Data are available at NAR Online.

Funding

US National Institutes of Health [R01AI052014, U54AI170791 to A.N.E., U54AI170855 to S.G.S., A.C.F.]; Gilead Research Scholars award (to A.C.F.). The open access publication charge for this paper has been waived by Oxford University Press – NAR Editorial Board members are entitled to one free paper per year in recognition of their work on behalf of the journal.

Conflict of interest statement. None declared.

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PERMALINK

HIV-1 usurps mixed-charge domain-dependent CPSF6 phase separation for higher-order capsid binding, nuclear entry and viral DNA integration

Sooin Jang

Gregory J Bedwell

Satya P Singh

Hyun Jae Yu

Bjarki Arnarson

Parmit K Singh

Rajalingam Radhakrishnan

AidanDarian W Douglas

Zachary M Ingram

Christian Freniere

Onno Akkermans

Stefan G Sarafianos

Zandrea Ambrose

Yong Xiong

Praju V Anekal

Paula Montero Llopis

Vineet N KewalRamani

Ashwanth C Francis

Alan N Engelman

Abstract

Graphical Abstract

Graphical Abstract.

Introduction

Materials and methods

Reagents

Biological resources

Computer code resources

Plasmid DNA constructions

Expression and purification of recombinant proteins

Recombinant CPSF6 mass assessment by mass photometry

LLPS assays with recombinant CPSF6 proteins

Propagation of human cells and viruses

HIV-1 infection and integration site analyses

Quantitative PCR for HIV-1 reverse transcription and bulk integration

Western immunoblotting

RNA interference-based depletion of targeted gene products

Antigen detection by immunofluorescence-based assays

CA nanotube-CPSF6 binding assay

Permeabilized virus binding assay

Cherry-CLP assembly and interaction with CPSF6

Statistical analyses

Results

CPSF6 determinants of LLPS activity and condensate stability

Figure 1.

The MCD is critical for CPSF6 puncta formation, viral nuclear incursion and speckle-proximal integration

Figure 2.

Arginine mixed-charge sequences critically underlie MCD functionality in HIV-1 infection

Figure 3.

The MCD is essential for higher-order capsid binding

Figure 4.

Figure 5.

NS depletion preferentially disrupts CPSF6 puncta formation

Figure 6.

Cyclophilin A-MCD-mediated HIV-1 nuclear penetration and integration targeting

Figure 7.

Discussion

CPSF6 LLPS activity

CPSF6 LLPS activity in capsid binding and HIV-1 infection

Study limitations

Supplementary Material

Acknowledgements

Contributor Information

Data availability

Supplementary data

Funding

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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