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PMC11075828 | Direct and indirect effects of CYTOR lncRNA regulate HIV gene expression | The implementation of antiretroviral therapy (ART) has effectively restricted the transmission of Human Immunodeficiency Virus (HIV) and improved overall clinical outcomes. However, a complete cure for HIV remains out of reach, as the virus persists in a stable pool of infected cell reservoir that is resistant to therapy and thus a main barrier towards complete elimination of viral infection. While the mechanisms by which host proteins govern viral gene expression and latency are well-studied, the emerging regulatory functions of non-coding RNAs (ncRNA) in the context of T cell activation, HIV gene expression and viral latency have not yet been thoroughly explored. Here, we report the identification of the Cytoskeleton Regulator (CYTOR) long non-coding RNA (lncRNA) as an activator of HIV gene expression that is upregulated following T cell stimulation. Functional studies show that CYTOR suppresses viral latency by directly binding to the HIV promoter and associating with the cellular positive transcription elongation factor (P-TEFb) to activate viral gene expression. CYTOR also plays a global role in regulating cellular gene expression, including those involved in controlling actin dynamics. Depletion of CYTOR expression reduces cytoplasmic actin polymerization in response to T cell activation. In addition, treating HIV-infected cells with pharmacological inhibitors of actin polymerization reduces HIV gene expression. We conclude that both direct and indirect effects of CYTOR regulate HIV gene expression.The introduction of antiretroviral therapy (ART) has successfully limited the spread of Human Immunodeficiency Virus (HIV) and improved patient clinical outcomes. However, a complete cure for HIV infection remains out of reach, as the transcriptionally silent but replication-competent provirus that is integrated into the host genome persists in long-lived cellular reservoirs, which are comprised of memory-resting CD4 T cells, as well as cells of myeloid lineages . These reservoirs are highly stable and are resistant to both ART and the effects of the host immune surveillance, thus posing a significant obstacle to eradicating the HIV reservoirs. Consequently, in most people living with HIV, interrupting ART leads to rapid viral load rebound, usually within weeks after treatment cessation [3–6]. As T cell stimulation triggers activation of proviral transcription, one strategy that has been proposed to eliminate the HIV reservoirs is a “Shock-and-Kill” approach, which utilizes latency-reversing agents (LRAs) to first activate dormant HIV-infected T cells and facilitate cell death by viral cytopathic effects or immune-mediated killing. This step is done in the presence of ART, so there are no further rounds of HIV replication. [7–9]. Alternatively, a “Block and Lock” approach frees infected individuals from ART by silencing HIV transcription and inducing a deep state of latency. Nevertheless, despite promising therapeutic options, these strategies and others have regretfully failed to achieve significant clinical efficacy. These failures highlight our lack of knowledge of the molecular mechanisms that govern latency establishment and reversal and the need for alternative therapies capable of eliminating the viral reservoirs [10–15]. Epigenetic constraints that suppress proviral gene transcription are essential for establishing HIV latency . Low levels of basal and elongating transcription factors in the infected T cell, together with the absence of the viral trans-activator of transcription (Tat), ensure that proviral transcription remains below detectable thresholds . Within the infected T cells, gene transcription of the integrated provirus and the host genome are synchronized . Both display key steps of gene transcription, which include initiation, promoter arrest, and elongation. HIV-Tat orchestrates transcription elongation of the provirus by binding to TAR RNA and recruiting P-TEFb and Super Elongation Complex (SEC) to the viral promoter [22–26]. However, despite extensive efforts to elucidate the mechanisms of metazoan transcriptional control and its role in the regulation of HIV gene transcription, the knowledge of how HIV latency is established is still incomplete . Long non-coding RNAs (lncRNAs) are transcripts with longer than 200 nucleotides that lack protein-coding capacity. To date, over 200,000 cell type-specific lncRNAs have been identified and display critical regulatory functions of many processes within cells [28–31]. However, the functions of most of these transcripts remain poorly understood. In the context of HIV, roles for several cellular lncRNAs have been documented [32–40]. Moreover, significant gaps still remain in our knowledge about the mechanistic roles that lncRNAs play in CD4 T cell activation and HIV latency. In this study, we monitored changes in gene expression in an HIV-infected Jurkat-derived T cell line (J-Lat 6.3) upon response to T cell stimulation with Phorbol 12-myristate 13-acetate—PMA/Ionomycin (P/I). We documented RNA expression in stimulated J-Lat 6.3 cells that carry either active or cells latent HIV, and among identified ncRNA, Cytoskeleton Regulator RNA (CYTOR) exhibited a profound change in expression in cells that expressed active HIV following T cell stimulation. CYTOR directly binds the HIV promoter and activates viral gene transcription and latency reversal by recruiting P-TEFb to the viral promoter. CYTOR also exerts its effects indirectly by controlling global gene expression along with actin dynamic pathways, thereby affecting T cell activation and HIV infection. In search for novel host regulators of HIV gene expression and viral latency, we employed RNA-Sequencing analysis to monitor changes in the transcriptome of Jurkat-derived J-Lat 6.3. These cells carry a transcriptionally repressed intact copy of HIV-1 proviral DNA with a GFP reporter under the control of the HIV promoter that is inserted in the nef gene. As expected, in response to T-cell stimulation, HIV gene expression in J-Lat 6.3 cells was enhanced, as exhibited by elevated expression levels of GFP . J-Lat 6.3 cells were stimulated with the PKC activator PMA and Ionomycin (P/I), which potently activate CD4+ T lymphocytes. Stimulated J-Lat 6.3 cells were then sorted by FACS based on their HIV-GFP expression and divided into two distinct populations: Stimulated cells that expressed HIV genes (GFP+; P1) and stimulated cells that carried latent provirus (GFP-) (Fig 1A). RNA from both cell groups was isolated, and libraries were generated for transcriptome analysis by next generation sequencing (RNA-Seq). As expected, a pronounced change in cellular gene expression, including mRNAs, miRNAs, snoRNAs, snRNAs and lncRNAs was observed in stimulated cells that expressed active HIV or latent HIV (Fig 1B). Subsequent RNA-Seq from HIV expressing cells that carry active (GFP+) or latent (GFP-), indicated different transcriptional profiles in cells where HIV is activated versus cells where the virus remained latent. A total of 3490 annotated transcripts were identified whose expression was changed in cells that carried transcriptionally active HIV relative to latent HIV. Of these, 2400 transcripts corresponded to protein-coding genes, while 843 were lncRNAs. Upon T-cell stimulation, 468 lncRNA transcripts were upregulated (enriched in cells expressing active HIV; GFP+), and 375 were downregulated (enriched in cells carrying latent HIV provirus; GFP-) (Fig 1C). We further assessed the relative expression levels of the highly ranked lncRNAs in CD4+ T primary T cells by RT-qPCR. Analysis was performed under resting or stimulating conditions of primary CD4+ T cells. For most tested lncRNAs, a shift in expression levels was confirmed when comparing primary CD4+ T cells under resting or stimulated conditions, where HIV was latent or active, respectively (Fig 1D). Notably, ncRNAs with reported effects on HIV replication and latency, including HEAL and NRON were identified via our RNA-Seq analysis, demonstrating the potential of this screening approach (Fig 1D). Also indicated are ncRNAs that are currently under investigation like IL21R-AS; PCBP3-AS; APOBEC3B-AS; IER3-AS (Fig 1D). Similarly, mRNAs genes, including HSP90 , ESR-1 , and IFI16 , that were previously reported to control HIV replication were also identified by our screening approach (S1 Table; GSE254771). Finally, we confirmed surface expression of CD25 and CD69 activation markers in stimulated primary CD4+ T cells that were infected with HIVGKO and carry active or latent HIV (S1 Fig). (A) FACS histogram analysis of PMA/Ionomycin (P/I)-stimulated J-Lat 6.3 cells. GFP(+) cells carrying active HIV (P1 region) were sorted from GFP (-) cells carrying latent HIV (GFP-). Cells were sorted and sent to RNA-Seq (n = 4). (B) Heatmap of differential transcript expression pattern (FC≥ a 2-fold change and above between cells carrying active versus latent HIV with an adjusted p value of ≤0.05. (C) Pie chart corresponding to the numbers of differentially expressed mRNAs and lncRNAs, up and downregulated in cells where HIV was reactivated. (D) RNA levels of selected ncRNA in primary CD4+ T cells. Analysis of expression levels of selected ncRNA based on the RNA-Seq analysis in primary CD4+ T cells that were either under resting conditions (-) or stimulated with P/I (+). RNA levels were analyzed by RT-qPCR. Data were normalized to gapdh levels. Data are from 2 healthy donors. Among the lncRNAs that were strongly induced upon T cell stimulation, we focused our work on Cytoskeleton Regulator RNA (CYTOR)—also known as lincRNA00152. Elevated RNA levels of CYTOR upon T cell stimulation were confirmed in J-Lat 6.3 and in primary CD4+ T cells (Fig 1D). To monitor the effects of HIV infection on CYTOR RNA levels, Jurkat T cells were infected with HIV and levels of CYTOR RNA were determined by RT qPCR relative to non-infected cells. Our analysis confirmed that CYTOR RNA levels were not affected by HIV infection (S2 Fig). CYTOR is an intergenic 828 nucleotide lncRNA located on chromosome 2p11.2. It is highly conserved in primates and rodents but less so in lower organisms. CYTOR is mainly present in the cytoplasm. However, previous reports show that it is also localized to the nucleus. Within the nucleus, CYTOR functions as an oncogene and is upregulated in multiple human malignancies . CYTOR also acts as an “endogenous sponge” for several micro-RNAs by binding to them, inhibiting their activity, and promoting malignancy. Interestingly, CYTOR reportedly regulates cellular actin dynamics and cytoskeletal reorganization in fibroblasts by controlling the expression of genes of the actin polymerization machinery . However, the functional importance of CYTOR in CD4 T cells and in the context of HIV infection has not been studied. We next conducted gain and loss-of-function studies in J-Lat 6.3 T cells to determine the role of CYTOR in regulating HIV gene expression. To achieve CYTOR over-expression, cells were transduced with a lentivirus that drives the expression of CYTOR—exons 1, 4, and 5, the most abundantly expressed form in humans . Following antibiotic selection, resistant J-Lat 6.3 T stable cells were subjected to RT-qPCR and exhibited a significant increase in CYTOR RNA levels relative to control cells (Fig 2A; blue bar versus grey bar). Reducing CYTOR expression (knockdown; KD) was also achieved by transducing J-Lat 6.3 T cells with a lentivirus encoding a CYTOR-targeting small-hairpin RNA (shRNA), resulting in a significant decrease of CYTOR RNA levels relative to control cells, expressing a scrambled shRNA as measured by RT-qPCR (Fig 2A; red bar versus grey bar). Parallel FACS-based analysis of GFP expression in HIV-infected J-Lat 6.3 cells, as a measure of viral gene transcription, revealed that in the absence of T-cell stimulation, no effects on HIV gene expression were observed upon modulation of CYTOR expression. However, following T cell stimulation with P/I, CYTOR over-expression led to a relatively small 2-3-fold increase of HIV GFP expression over control cells (Fig 2B; compare blue to grey bars). In contrast, depletion of CYTOR led to a 5-fold decrease in HIV gene expression over control cells (Fig 2B; compare red to grey bars). HIV GFP expression in control cells expressing scrambled shRNA was unaffected (Fig 2B; grey bar). (A). Modulation of CYTOR RNA levels in J-Lat 6.3 cells. RT-qPCR analysis measuring CYTOR RNA levels in J-Lat 6.3 T cells, where CYTOR expression is knockdown (KD; red bar) or overexpressed (light blue bar). RNA levels were normalized to gapdh and presented relative to control cells expressing scrambled shRNA (grey bar). Statistical significance is based on calculating ±SD of data points from four independent experiments using two-way ANOVA. ***p≤0.05. (B) Effects of CYTOR on HIV gene expression. FACS quantification analysis of the percentage of cells that express HIV-GFP in P/I stimulated J-Lat 6.3 cells expressing control scramble shRNA (grey bar), or in which CYTOR was overexpressed (blue bar) or knockdown (KD; red bar). Statistical significance is based on calculating mean ± SD from three independent experiments using two-way ANOVA. ***p≤0.05. (C) Kinetics of latency establishment in the context of CYTOR expression. 2D10 latency model Jurkat T cells that carry a mini-Tat-Rev GFP under the regulation of the HIV LTR promoter and express either scrambled shRNA (grey), CYTOR KD (red), or cells over-expressing CYTOR (blue) were reactivated and sorted to obtain a pure cell population that expresses GFP. GFP expression was then followed over time as a measurement of entry into latency. Statistical significance is based on calculating mean ± SD from three independent experiments using two-way ANOVA. ***p≤0.05 and ns: not significant. We also followed the establishment of HIV latency post-T cell activation, documenting HIV-GFP expression in control 2D10 cells that expressed scramble shRNA or in cells where CYTOR expression levels were modulated. Like J-Lat 6.3 cells, 2D10 cells serve as a Jurkat-based latency cell model that carries a minimal Tat-Rev cassette in the context of a GFP reporter under the regulation of the HIV promoter. Upon T-cell stimulation of control 2D10 Jurkat cells, we confirmed that the expression of HIV-GFP was significantly induced. Control and CYTOR-modulated stimulated 2D10 Jurkat cells were sorted based on their HIV-GFP expression, obtaining a relatively pure cell population with 100% HIV-GFP expression levels. We then monitored latency establishment by following GFP expression in the context of control or CYTOR-modulated cells (Fig 2C). Our FACS analysis revealed that lower CYTOR levels were associated with a rapid establishment of latency relative to control cells (Fig 2C; grey versus red lines). Conversely, CYTOR-over-expression enhanced latency reversal, as determined by the elevated levels of HIV-GFP expression that remained relatively high for an extended period following T cell stimulation (Fig 2C; blue line). These results suggest that CYTOR expression activates HIV gene expression, significantly reversing latency in 2D10 cells. Next, we shifted our analysis to CD4+ primary T cells isolated from healthy donors and the natural target cells for HIV infection. Depletion of CYTOR in primary human CD4+ T cells was achieved by first stimulating purified cells with anti CD3/CD28 beads and IL2 and then transducing them with a lentivirus encoding a CYTOR-specific shRNA. Lentivirus driving the expression of scrambled shRNA was used as a control (Fig 3A; n = 3). RT qPCR confirmed a significant decrease in CYTOR expression RNA levels relative to control cells that expressed the scramble shRNA (Fig 3B). The next day, CYTOR-depleted CD4+ primary stimulated cells (KD) or control cells were transduced with HIVGKO, which codes for a codon-optimized GFP reporter under the control of the HIV-1 promoter and in the context of expression of all viral proteins, and a mKO2 reporter under the control of the constitutive promoter EF10 α . HIVGKO transduction can be analyzed by FACS two days later, monitoring parallel transduction efficiency (mKO2+), as well as HIV gene expression (GFP+). Upon transduction of stimulated primary CD4+ T cells, KD of CYTOR led to decreased HIV gene expression, as monitored by reduced levels of HIV-GFP-expressing cells. Conversely, the proportion of cells that expressed EF10 α -mKO2 was slightly elevated in control or CYTOR KD-expressing cells, implying that transduction efficiencies were not affected due to CYTOR depletion but rather specifically drove HIV into a latency state (Fig 3C and 3D). (A). Experimental workflow overview for isolating primary CD4+ T cells. See methods for a detailed description. The figure was generated by Biorender. (B). Depletion of CYTOR in stimulated primary CD4+ T cells using lentivirus encoding CYTOR shRNA. Data were measured by RT-qPCR, normalized to GAPDH, and presented relative to cells expressing scrambled shRNA—set to 1. Statistical significance is based on calculating mean ± SD from three independent experiments using two-way ANOVA. ***p≤0.05; n = 3. (C). FACS analysis presenting effects of CYTOR knockdown (KD) on HIVGKO infection in primary CD4+ T cells. Cells were stimulated and then transduced with HIVGKO before being analyzed by FACS for mKO2 and GFP expression. (D). Quantification of quadrate percentage from three independent experiments of FACS analysis for HIVGKO transduction in CD4+ primary T cells, where CYTOR is KD. Statistical significance is based on the calculation of mean ±SD from three independent experiments (n = 3) using Two-way ANOVA. ***p≤0.05. Towards direct effects of CYTOR on HIV gene transcription, this would require its localization within the cell nucleus. We therefore monitored the subcellular distribution of CYTOR lncRNA between the nucleus and cytoplasm in resting or activated conditions of primary CD4+ T cells by cell fractionation and subsequent RT-qPCR analysis (Fig 4A). Levels of CYTOR RNA were compared to those of the abundant 7SK lncRNA, which is known to interact with inactive P-TEFb. RNA levels were normalized to the 7SL RNA, which does not bind to the HIV promotor and is commonly used as a specificity control for these experiments . Our analysis showed that CYTOR is localized to the cytoplasm and the nucleus. Notably, upon T cell stimulation, levels of nuclear CYTOR were elevated relative to the levels of nuclear 7SK, which were decreased (Fig 4A). To further extend our understanding of the mechanism by which CYTOR activates HIV gene expression, we tested whether CYTOR binds to the HIV promoter, thereby regulating HIV gene transcription. We monitored CYTOR occupancy on the HIV promoter by employing Chromatin Isolation by RNA Purification (ChIRP) analysis in J-Lat 6.3 cells. In vitro-transcribed biotinylated CYTOR RNA was synthesized, purified, and then incubated with ChIP material isolated from unstimulated or P/I stimulated HIV J-Lat 6.3 cells. RNA-protein complexes were then specifically pulled down with streptavidin beads, and pulled down levels of CYTOR on the HIV promoter were monitored by RT-qPCR with specific primers that target the viral promoter (Fig 4B). Our analysis showed that CYTOR binds to the HIV promoter even in unstimulated conditions. Significantly, CYTOR occupancy on the viral promoter was further elevated following T-cell stimulation (Fig 4B). CYTOR occupancy on gene promoters was also demonstrated for cellular genes that are known to be regulated by P-TEFb, such as NF-κB, IL21Ra, myc. Of note, the binding of CYTOR to HIV downstream reverse transcriptase sequences was not observed, suggesting that the specificity of CYTOR within the HIV genome lies within the HIV promoter (S3 Fig). (A). CYTOR is localized to the nucleus and its levels are elevated upon T-cell stimulation. Resting or stimulated primary CD4+ T cells were subjected to cell fractionation, separating the samples into a nuclear fraction (grey bar) or cytoplasmic fraction (light blue bar). Samples were then subjected to RT-qPCR and monitored for CYTOR or 7SK ncRNA levels. Data were normalized to 7SL RNA in each of the cellular fractions and conditions. Data are presented relative to cytoplasmic fraction in each condition—set to 1. (B). ChIRP-qPCR analysis for CYTOR binding to the HIV promoter. CYTOR-specific (black bar) or control lacZ (red bar) antisense biotinylated probes were incubated with lysates isolated from unstimulated or P/I stimulated J-Lat 6.3 cells. Biotinylated RNA was pulled down with streptavidin beads, and following washing, associated DNA was eluted and analyzed by qPCR with primers for the HIV promoter. Statistical significance was calculated between the two probes and between unstimulated and stimulated states. IgG served as a non-specific antibody for IP control (grey bar). The analysis is based on calculating mean ± SD from three independent experiments using two-way ANOVA. ***p≤0.05. **0.05≤p≤ 0.1; n.s—not significant. (C, D) CYTOR affects the phosphorylation state of RNAPII CTD and histone landscape. ChIP qPCR analysis in control or CYTOR KD J-Lat 6.3 cells. ChIP material from cells was immune-precipitated (IP) with antibodies targeting RNAPII-Ser2P or RNAPII Ser5P (C); or for H3K4me3 and H3K27Ac histone activation marks (D). IP fraction was analyzed for enrichment of the indicated modifications on the HIV promoter by qPCR with specific primers. Non-specific IgG served as a control (grey bar). Percentage of input are means ±SD; n = 3; *** p≤0.05 calculated between scrambled and KD cells for each antibody. n.s—not significant. To further obtain insights into the mechanisms of action of CYTOR, we performed Chromatin immunoprecipitation (ChIP) qPCR from J-Lat 6.3 cells, where CYTOR expression was manipulated. We monitored the levels of phosphorylated C-terminal domain (CTD) of RNA Polymerase II (RNAPII) at Ser2 (Ser2P) or Ser5 (Ser5P) residues on control or CYTOR KD expressing cells, using specific antibodies that target the CTD phosphorylation states of RNAPII (Fig 4C). CDK9/P-TEFb phosphorylates Ser2 and marks RNAPII pause-release and elongation of transcription [50–52]. CDK7/TFIIH phosphorylates Ser5P on the CTD and catalyzes transcription initiation and promoter clearance. Our analysis demonstrated that in the context of CYTOR depletion, levels of Ser2P on the HIV promoter were decreased without affecting those of Ser5P, implying the involvement of P-TEFb in CYTOR-mediated HIV gene activation (Fig 4C). In addition, ChIP-qPCR was employed using antibodies that target the histone activation markers, H3K27Ac or H3K4me3. Our analysis confirmed that CYTOR mediates its activation properties by modifying the histone landscape around the HIV. Upon CYTOR depletion, levels of these histone activation markers were reduced (Fig 4D). These results further imply that CYTOR activates HIV gene expression via P-TEFb, affecting transcription elongation. To expand the above results on the mechanism by which CYTOR enhances HIV gene expression, we employed RNA-precipitation (RNA-IP; RIP) followed by RT-qPCR in J-Lat 6.3 cells under resting or stimulated conditions (Fig 5). As our above results indicate that CYTOR promotes the Ser2 phosphorylation on the CTD, which is mediated by CDK9, we monitored CYTOR association with P-TEFb. Lysates isolated from nuclei from resting or stimulated J-Lat 6.3 cells were incubated with antibodies that target CDK9 or CYCLIN T1, and samples were IP followed by RT-qPCR to detect CYTOR RNA levels by using specific primers. We show that upon T cell stimulation, the levels of CYCLIN T1 and CDK9 that were associated with CYTOR RNA increased. As expected, levels of 7SK that are associated with P-TEFb were reduced upon T cell stimulation. We also followed the association of P-TEFb with 7SL, which served as control. As expected, P-TEFb was not associated with 7SL in each of the tested conditions. These results indicate that CYTOR associates with P-TEFb in cells (Fig 5A). (A) RIP analysis demonstrates the association of CYTOR with P-TEFb. Isolated ChIP material from resting or stimulated J-Lat 6.3 CD4+ T cells was subjected to immune precipitation with antibodies targeting CDK9 or CYCLIN T1 of P-TEFb, followed by RT-qPCR with primers for the relevant lncRNA (7SK or CYTOR). Non-specific IgG served as a control for the IP step. 7SL ncRNA served as a control for an RNA that does not associate with P-TEFb and, therefore, not precipitated with CDK9 or CYCLIN T1 antibodies. Statistical significance is based on the calculation of mean ±SD from three independent experiments using two-way ANOVA. ***p≤0.05. ** 0.05≤p≤0.1; ns: not significant. (B) CYTOR associates with P-TEFb in cells. RNA pull-down followed by western blotting where lysates from J-Lat 6.3 cells were incubated with an in-vitro transcribed biotinylated CYTOR anti-sense probe and reactions were pulled down with streptavidin beads. Eluted RNP complexes were subjected to western blotting with indicated antibodies. Non-specific IgG served as a control for non-relevant IgG. Scramble RNA served as RNA that does not associate with P-TEFb. 7SK probe confirmed association with P-TEFb. Input is 5% of the total cell lysate . Next, we performed RNA pull-down experiments combined with western blotting to detect P-TEFb subunits (CYCLIN T1/CDK9) that are associated with CYTOR. Lysates from J-Lat 6.3 cells were incubated with an in-vitro transcribed biotinylated anti-sense CYTOR probe, and reactions were pulled down with streptavidin beads. Eluted RNP complexes were then subjected to western blotting with antibodies that target CYCLIN T1 or CDK9, demonstrating the association of CYTOR lncRNA with P-TEFb within cells. Non-specific IgG was used as a specificity control for the IP step, while a non-specific scramble RNA probe served as a control for RNA-protein association. In addition, a 7SK RNA probe confirmed the association with P-TEFb (Fig 5B). These results establish that CYTOR binds to the HIV promoter and suggest that its activation effects are mediated by association with P-TEFb. Since CYTOR has been previously recognized as a regulator of cytoskeleton-regulating genes in fibroblasts , we assessed if it could also affect HIV gene expression by indirect mechanisms through regulation of its downstream targets. For this, we performed RNA-Seq analysis in stimulated primary CD4+ T cells, where CYTOR expression was depleted or over-expressed (n = 3). CYTOR modulation of expression did not affect the activation state of cells as monitored by staining with T cell activation markers (S4 Fig). Analyzing changes in the cellular transcriptome of stimulated CD4+ T cells upon depletion of CYTOR revealed a modest change in the cell gene expression program (S2 Table). Additional gene GO analysis identified significant enrichment scores in various cellular pathways, including those of gene expression, signal transduction as well as actin dynamics and T-cell activation (Fig 6A and 6B). (A). Volcano plot of the expression pattern of genes from an RNA-Seq analysis upon CYTOR depletion following T cell stimulation. -Log10 P is shown on the y-axis, and Log2FC is on the x-axis. RNA was isolated from 3 biological replicates (n = 3). The fold of change cutoff is defined as FC ≥2. FDR of p≤0.05 was used as a cutoff for significance. (B). Gene Ontology analysis for enriched CYTOR gene targets. For enrichment analysis, the DAVID program was employed to identify enriched pathways and terms associated with the selected genes. (C). Experimental flow for microscopy-base analysis of cell morphology and formation of F-actin rich structures. The figure was generated by Biorender. (D). Representative confocal images of F-actin organization for control and CYTOR KD Jurkat cells after contact with anti-CD3/28 coated surfaces. Cells were stained with fluorescent phalloidin and DAPI to visualize F-actin and cell nuclei. Shown are merged images of both channels, scale bar = 10 μM). (E). Relative frequency of cells with circumferential F actin ring (AR) in control or CYTOR KD cells with proper cell spreading and circumferential F-actin relative to control cells (mean± SD, 100 cells per experiment/condition, n = 3). (F). Relative CYTOR RNA levels in CYTOR KD Jurkat cells relative to control cells of the cells analyzed in (E). (G). Representative images of the different morphotypes observed for Jurkat cells after anti-CD3/28 surface stimulation (analyzed as in D), (H). Quantifying the frequency of the morphotypes defined in (G) for control and CYTOR KD Jurkat cells (mean± SD, 100 cells per experiment/condition, n = 3). ** 0.05≤p≤0.1. (I). Inhibition of actin remodeling disrupts HIV gene expression upon T cell activation. 2D10 cells carrying an integrated HIV-GFP provirus where CYTOR expression was either depleted or over-expressed were treated with an actin polymerization inhibitor, Latrunculin B (LanB) for 1 hour, followed by T cell stimulation with anti-CD3/CD28 for an additional 3 hours. Cells were harvested 24 hours later, and the percentage of cells expressing HIV GFP was monitored by FACS. Data are presented as fold of activation relative to untreated cells and activated with the indicated T cell activator. Statistical significance is based on calculating mean ± SD from three independent experiments using two-way ANOVA. ***p≤0.05. ** 0.05≤p≤0.1. T cell activation elicits complex and highly dynamic signaling cascades that ultimately lead to the activation of transcription factors, including NF-κB and NF-AT, to increase the expression of T cell receptor target genes . The involvement of these transcription factors in HIV gene expression, at least in part, explains the beneficial effects of T cell activation on HIV gene expression . Since many of the downstream signaling events elicited by TCR engagement depend on the immediate polymerization of cortical actin, we tested if CYTOR affects the actin polymerization response to TCR engagement of Jurkat T cells. Scramble control or CYTOR KD Jurkat cells were placed on a cell stimulatory surface coated with anti-CD3/CD28 antibodies, fixed, and stained for F-actin. Control cells displayed the typical cell spreading and formation of circumferential F-actin-rich rings (actin ring; AR) (Fig 6C, 6D, 6E and 6F). Although CYTOR expression was only moderately reduced in KD cells (Fig 6F), fewer cells responded to TCR stimulation (approx. 40% less cells with AR in CYTOR KD than in control cells; Fig 6E). Detailed analysis of the different cell morphologies revealed that the CYTOR KD particularly resulted in a significant increase in the fraction of cells that were unable to both spread as well polymerize actin into an F-actin ring in response to T cell activation. In contrast, the frequency of cells that failed to spread despite efficient actin polymerization was unaffected (Fig 6H). Increasing CYTOR levels by overexpression did not further increase the frequency of cells that formed ARs, did not alter the morphology of F-actin structures formed in response to TCR activation, and did not result in the formation of ARs in the absence of TCR stimulation (S5 Fig). We conclude that CYTOR is an important regulator of TCR-induced actin polymerization in CD4+ T cells, but its normal endogenous expression levels are not limiting for this response. To assess whether TCR-induced actin remodeling affects HIV gene expression in our experimental system, we measured the induction of HIV gene expression by TCR engagement in 2D10 cells, a CD4+ T cell line that carries a latent GFP cassette under the control of the LTR promoter. Experiments were performed in the absence or presence of the actin polymerization inhibitor, Latrunculin B—an inhibitor that interferes with actin polymerization and is reversible upon washout (Fig 6I). Stimulation with anti-CD3/anti-CD28 resulted in a marked induction of GFP expression and, as observed before, silencing CYTOR expression reduced this induction. Notably, interfering with actin polymerization during the first 3 hours of TCR stimulation in control cells limited the induction of HIV gene expression to the levels observed upon CYTOR KD, and interference with actin dynamics in CYTOR KD did not result in an additional reduction of GFP expression. Finally, overexpressing CYTOR rendered the TCR-mediated induction of GFP expression insensitive to Latrunculin B (Fig 6I) . Together, these results reveal that the regulation of host cell actin dynamics is necessary but not sufficient for the regulation of gene expression of latent HIV. In search of regulators of HIV latency, we profiled changes in the expression of ncRNAs by employing RNA-Seq analysis in resting and stimulated HIV-infected J-Lat 6.3 T cells, comparing RNA expression levels in cells that carry active HIV (GFP+) or latent HIV (GFP-). Our analysis show that different transcriptional profiles exist in cells where HIV is activated versus cells where it remains latent. CYTOR lncRNA was identified as one of these RNAs, and its expression is elevated upon T cell stimulation, where HIV is active. These observations were further confirmed in primary CD4+ T cells (Fig 1). Functional analyses show that following T cell stimulation, over-expression of CYTOR activates HIV gene expression, while its depletion inhibits viral gene expression. Significantly, upon T cell stimulation, depletion of CYTOR promoted entry of HIV into a latent state, while its over-expression delayed entry into latency and enhanced latency reversal (Fig 2). Effects of CYTOR on HIV infection and latency establishment were also confirmed in stimulated primary CD4+ T cells (Fig 3). We are aware that the model of stimulated CD4+ primary cells does not recapitulate the actual state of the reservoir, which is mainly comprise of resting CD4+ T cells that do not support HIV infection. As this is a limitation of the current study, we are trying to adopt a recently developed gene editing approach to lncRNAs to deplete CYTOR in this unique cell population and monitor the effects of latency kinetics without altering its activation . Mechanistically, our observations show that CYTOR directly binds to the HIV promoter and enhances the phosphorylation of the Ser2 CTD of RNAPII through association with P-TEFb to activate viral gene expression (Figs 4 and 5). Changes in histone activation marks around the viral promoter in CYTOR-depleted cells also imply that CYTOR activates the proviral gene expression (Fig 4). In addition to the direct effects of CYTOR on HIV gene expression, we also demonstrate that CYTOR controls global gene expression. CYTOR is recruited to other gene promoters that are regulated by P-TEFb, like myc, NF-κB, and IL2Ra (S3 Fig). Among the identified enriched pathways that potentially are regulated by CYTOR are those that are involved in actin dynamics. Consistently, reduced levels of CYTOR expression are associated with reduced polymerization of cortical actin in response to TCR engagement (Fig 6). In turn, elevated levels of CYTOR do not further increase actin polymerization in response to T cell stimulation and cannot induce morphological responses of T cells in the absence of stimulation (S5 Fig). Thus, CYTOR is an important regulator of TCR-induced actin polymerization in T cells. However, its normal endogenous expression levels are sufficient for a proper response. To test a mechanistic link between actin remodeling, CYTOR levels, and HIV gene expression, we inhibited actin dynamics with specific inhibitors (Fig 6I). Effects of inhibition of actin polymerization phenocopied the effect of CYTOR depletion on HIV gene expression, suggesting that CYTOR may affect HIV gene expression by the regulation of genes that control cellular actin dynamics (Fig 6I). Accordingly, we propose a model where CYTOR exerts its effects on global gene expression and promotes HIV gene expression by both direct and indirect effects (Fig 7). CYTOR directly binds the HIV promoter and recruits the elongation transcription machinery to enhance RNAPII CTD phosphorylation and deposition of active histone markers around the HIV promoter, ultimately activating HIV gene expression. Indirectly, CYTOR controls gene targets that regulate actin dynamics in the nucleus and at the plasma membrane to optimize the response to T cell activation, presumably via the regulation of cellular gene expression. Following T cell activation, levels of CYTOR are elevated in the nucleus. CYTOR is recruited to the HIV promoter and binds to P-TEFb, leading to the activation of viral gene expression. Cellular genes regulated by CYTOR include actin remodeling genes that promote actin polymerization and the indirect activation of HIV gene expression. Like CYTOR, other lncRNAs have been reported to occupy the HIV promoter and modulate its activity at either transcriptional or posttranscriptional levels . Most act as scaffolds that associate with other transcriptional activators or repressors to control HIV gene expression [35–39,58–60]. In the case of CYTOR, its effects on gene expression occur by recruiting the transcription elongation machinery to activate gene expression, either from the viral promoter or other cellular promoters. It will be essential to identify other partners that are associated with CYTOR lncRNA and control HIV promoter activity. As we also aim to dissect the role of CYTOR in gene expression control, specifically for HIV gene regulation, it will be essential to define how events within the nucleus are regulated by CYTOR and translated to the control of downstream effector functions of stimulated T cells. Future studies will further identify the downstream targets of CYTOR that control actin dynamics upon T-cell activation. As additional pathways were identified by our RNA-seq analysis in CYTOR-depleted cells, we visualize that future work will identify novel downstream targets of CYTOR and elucidate their mechanisms of function in regulating HIV gene expression and latency. These may open new ways for developing novel therapeutic tools that will be integrated or substitute current strategies to successfully eliminate the HIV reservoir. Jurkat J-Lat 6.3 T cells are immortalized human T lymphocytes that serve as a model for studying HIV latency, as it harbors a transcriptionally silent integrated HIV provirus that encodes for a GFP reporter instead of Nef, which reactivated following T cell stimulation. 2D10 cells are also Jurkat-based T cells, carrying a mini—HIV cassette coding for Tat and rev and a 2dGFP reporter gene. Jurkat T cells were maintained in RMPI medium (GIBCO) containing with 10% fetal bovine serum (FBS), 2mg/ml L-glutamine, penicillin-streptomycin, and non-essential amino acids (Sigma, M7145). Cells were cultured at 37°C with 5% CO2. Human Embryonic Kidney HEK293T, this cell line was mainly used for the production of viral-like particles were maintained in DMEM complete medium (GIBCO). Cells were cultured at 37°C with 5% CO2. For the isolation of primary human CD4+ T cells, human Buffy Coats from anonymous healthy donors were obtained from the Soroka Medical Center Hospital Blood Bank. At day 0, PBMCs were isolated over a Ficoll gradient (Millipore). PBMCs were maintained at 2 x10 cells/ml overnight at 37°C. CD4+ T cells were isolated by negative selection with the RosetteSep Human CD4+ T Cell Enrichment Cocktail Stemcell Technologies), resulting in homogenous populations of CD4+ T cells with a purity of 90–95% as assured by flow cytometry. CD4+ T cells were cultured in complete RPMI media containing recombinant human IL2 at 20 U/ml (Roche) to a final concentration 10 cells/ml. Cells were then stimulated with anti-CD3/CD28 dynabeads (Invitrogen) and further cultured for 48 hour. The level of activation was monitored by FACS measuring staining with APC anti human CD25 (Biolegend #302609) and Pacific Blue anti-human CD69 (Biolegend #310919). At day two, stimulated cells were counted, centrifuged for 5 minutes at 1500 rpm, and resuspended in fresh RPMI to a final concentration 0.5x10 cell/ml and IL-2 before transduction with high titter HIV carrying CYTOR shRNA at an MOI of 10. 24 hr later (day three), cells were further transduced with HIVGKO lentivirus at MOI of 10. Transduced cells were cultured in complete RPMI media containing recombinant human IL2 and dynabeads at a ratio of 25 μl human beads per 10 million cells and analyzed by FACS at day five. For the IP of P-TEFb, we used the following antibodies: anti-CDK9 (Abcam ab6544) or anti-CYCLIN T1 antibodies (Abcam; ab176702). For ChIP-qPCR for the detection of histone marks activation markers, we used anti-H3K27Ac (ab4729) and anti-H3K4me3 (ab8580). For detecting the different states of the phosphorylation of RNAPII CTD, we used the phosphorylated serine 2 antibody (Ser2P; ab238146) and phosphorylated serine 5 (Ser5P; ab5131). To monitor T cell activation following stimulation, the following antibodies were used: APC anti human CD25 (Biolegend; 302609); Pacific Blue anti human CD69 (Biolegend; 310919). Actin remodeling in response to T cell receptor (TCR) engagement was monitored by forming circumferential F-actin rings as previously described . In brief, stimulatory coverslips were prepared by coating with a 0.01% poly-L-lysine (PLL; Sigma) solution for 10 minutes at room temperature, followed by wet-chamber incubation for 3 hours at 37°C with 7 μg/ml anti-CD3 antibody (50 μl per coverslip, clone HIT3a against CD3E; BD Biosciences) in phosphate-buffered saline (PBS). Stimulatory coverslips were subsequently washed in PBS and stored at 4°C in PBS until use. 5x10 cells per anti-CD3-coated coverslip, respectively) were used to seed coverslips for 4 minutes to allow TCR-mediated actin ring formation. Cells were subsequently fixed in 3% paraformaldehyde for 15 minutes, permeabilized for 2 minutes in 0.1% TritonX-100, and blocked for 30 minutes in 1% Fetal Calf Serum (FCS) in PBS. F-actin was visualized with tetramethyl rhodamine isothiocyanate (TRITC)-conjugated phalloidin (1:1,000, 1 hour, room temperature; Sigma). Samples were mounted on glass slides and analyzed by epifluorescence (Olympus IX81 S1F-3, cellM software) and confocal (spinning-disc PerkinElmer UltraView VoX, Velocity software) microscopes. For quantification of phenotype frequencies, at least 100 transfected cells were counted. Pseudotyped viruses were generated in HEK293T cells as described . Briefly, the plasmid driving the expression of the shRNA transgene was transfected into cells using Lipofectamine 2000 (Invitrogen) together with other lentiviral packaging plasmids coding for Gag, Pol Tat Rev, and the VSV-G envelope. Transfections were done in a 10cm format, and the supernatant containing the virus was harvested 72 hours post-transfection, filtered through a 0.45 μm filter spun at 2000 rpm for 5 min to remove cells debrides and stored at -80°C. 2x10 Jurkat T cells were transduced with the pseudotyped particles for transduction. 16 hours later, the medium containing lentiviral particles was changed. Following transduction, cells were cultured in a medium supplemented with 2 μg/ml of puromycin to eliminate non-transduced cells that did not express shRNA. Upon the death of all the control cells, the medium was changed, and surviving cells were propagated for future experiments. For transducing CD4 primary cells, we used HIVGKO (a gift from Eric Verdin), which codes for a codon-optimized GFP reporter under the control of the HIV-1 promoter and in the context of expression of all viral proteins and a mKO2 reporter under the control of the constitutive promoter EF1α . For knockdown (KD) of CYTOR expression, J-Lat 6.3 cells were transduced with lentiviruses that drive the expression of shRNA that specifically targets CYTOR. Cells were next selected on puromycin, and polyclonal stable cells were monitored for CYTOR expression by RT-qPCR. To achieve CYTOR over-expression, cells were transduced with a lentivirus that drives the expression of CYTOR—exons 1, 4 and 5, the most abundantly expressed form in humans. Following antibiotic selection, resistant J-Lat 6.3 T cells were subjected to RT-qPCR to confirm CYTOR over-expression or knockdown. CYTOR RNA levels were normalized to the gapdh gene. Although we are aware that GAPDH expression is elevated following TCR stimulation, we did analyze several cellular genes in search of a better marker, but concluded that, e.g., genes for actin polymerization machinery are all affected more strongly by T cell activation than gapdh . We therefore used gapdh for normalization. To obtain CYTOR knockdown in stimulated primary CD4+ T cells, cells were isolated from health donors (n = 3) and stimulated with anti-CD3/CD28 beads (1:1 ratio of beads to cell number). Cells were cultured on stimulation media (RPMI+IL2), and on day 3 post isolation and stimulation, cells were subjected to transduction with lentivirus expressing shRNA against CYTOR. The following day cells were transduced with HIVGKO and 48 hour later were analyzed by FACS for HIV-GFP and mKO2 expression. To monitor the effects of CYTOR in promoting HIV latency, we followed the kinetics of entry of stimulated Jurkat 2D10 T cells that express a cassette of the HIV provirus, expressing 2dGFP reporter. Cells where CYTOR is depleted or over-expressed and control cells that express scramble shRNA were activated with P/I and then sorted by FACS to isolate those that express GFP. Cells were then grown, during which their HIV GFP expression was followed by FACS. Control cells expressing scramble shRNA or cells where CYTOR expression was depleted (KD) were cross-linked with 1% formaldehyde for 10 minutes and then washed with PBS and reverse cross-linked with glycine (125mM; 5 minutes). Cells were then lysed for 10 minutes on ice in 130μl sonication buffer (20 mM Tris pH-7.8, 2 mM EDTA, 0,5% SDS, 0.5 mM phenylmethylsulfonyl fluoride (PMSF), and 1% protease inhibitor cocktail), and the nuclear pellets were collected. DNA was fragmented by sonication at the following settings: amplitude 20% for 30 cycles at 10 seconds on/10 seconds off. Samples were centrifuged (15 minutes, 14,000 rpm, 4°C). The soluble chromatin fraction (25 μg) was collected and immunoprecipitated (IP) overnight at 4°C on a rotating wheel in IP buffer (0.5% Triton X-100, 2 mM EDTA, 20 mM Tris pH-7.8, 150 mM NaCl and 10% glycerol) with 2.5 μg of one of the indicated antibodies. The next day, the IP material was incubated with 25 μl dynabeads protein G for two hours to ensure the binding of the antibody to the magnetic beads. DNA was eluted with freshly prepared elution solution (1% SDS and 0.1 M NaHCO3) and heated at 65°C overnight to reverse-crosslink the samples. Precipitated DNA fragments were then extracted using a ChIP DNA clean and concentrator kit (ZYMO Research), and HIV DNA levels were quantified by qPCR with the primers specifically located on the NFκB region at the HIV-LTR promoter. All signals were normalized relative to input DNA. ChIP assays were also performed with an anti-rabbit or mouse IgG as negative control. 3x10 cells were cross-linked with freshly made 1% formaldehyde in PBS for 10 minutes at room temperature while shaking. Crosslinking was quenched with 125 μM glycine for 5 minutes at room temperature. Cells were centrifuged at 1200 rpm for 5 minutes at 4°C and washed twice with PBS on ice. The pellet was re-suspended in 300 μl of sonication buffer (50mM Tris 7.0, 10mM EDTA, 1% SDS, DTT, PMSF, protease inhibitors (Roche), and RNase inhibitor (NEB). Cells were then incubated on ice for 10 minutes and sonicated in Bioruptor at high settings of 3 rounds each of 10 cycles 40 seconds ON/40 seconds OFF. Water was changed to ice-cold between the rounds. Sociated samples were centrifuged at max speed for 10 minutes at 4°C, and then chromatin material was kept at -80°C. For IP, chromatin was diluted in twice the volume of hybridization buffer (500 mM, NaCl, 1% SDS, 100 mM Tris pH-8, 10 mM EDTA, 15% Formamide, protease inhibitors (Roche) and RNase inhibitor (NEB).2 μg of biotinylated RNA was added to 0.5 ml diluted chromatin and mixed by end-to-end rotation at 37°C for 4 hours. Streptavidin-magnetic beads were washed three times in sonication buffer, blocked with 500 ng/μl yeast total RNA and 1 mg/ml BSA for 1 hour at room temperature before resuspended in their original volume. 40 μl of beads were added, and the reaction was incubated for 30 minutes at 37°C. Beads were captured by a magnet and washed five times with the wash buffer (2x SSC, 0.5% SDS, supplemented with fresh DTT and PMSF). Beads were resuspended in 3-times of the original volume with the DNA elution buffer (50 mM NaHCO3, 1% SDS, 200 mM NaCl), and DNA was eluted with a cocktail of 100 μg/ml RNaseA (Sigma) and 0.1 U/μl RNase H (Epicenter). Chromatin was reverse-cross-by treatment with 0.2 U/μl proteinase K at 65°C for 45 minutes. DNA was then extracted with an equal volume of phenol:chloroform: isoamyl alcohol(Invitrogen) and precipitated with ethanol at -80°C overnight. For probe in-vitro transcription of linear RNA synthesis, 1 μg of RNA was transcribed and biotinylated using AmpliScribe-T7-Flashbiotin-RNA transcription kit (Epicentre) according to the manufacturer’s instructions. Eluted DNA was analyzed by qPCR with primers specific to the HIV promoter. 10 Jurkat J-Lat 6.3 cells were washed twice with PBS and resuspended in 800 μl of RNA-IP buffer (0.5% NP-40, 20 mM HEPES pH 7.8, 100 mM KCl, 0.2 mM EDTA supplemented with RNase inhibitor (NEB) and protease inhibitor (Sigma). Cells were cross-linked, and cell lysate was incubated on ice for 10 minutes before isolating nuclei through centrifugation at 2500g for 15 minutes. The supernatant was collected and resuspended in freshly prepared RIP buffer. ChIP material was then sonicated, and the pelleted nuclear membrane and debris were removed by centrifugation at 13,000 rpm for 10 minutes. Isolated ChIP material was incubated with 2.5 μg of indicated antibodies overnight at 4°C. Then, 20 μl of pre-blocked BSA protein A beads were added and incubated for an additional 2 hours at 4°C. 50 μl of cell lysate was collected as input samples. Beads were washed 4 times with washing buffer (0.5% NP-40, 20 mM HEPES pH 7.8, 100 mM KCl, 0.2 mM EDTA supplemented with RNase inhibitor (NEB) and protease inhibitor (Sigma) to remove unbound material. The pellet was resuspended in 100 μl of the lysis buffer and extracted using a TRIZOL reagent (Sigma). RNA was reverse transcribed using qPCRBIO kit (PCPbiosystems), and qPCR was performed using indicated primers against CYTOR or 7SK ncRNA. The amplification of 7SL RNA served as a control RNA that is not associated with P-TEFb. Input RNA was extracted and reverse-transcribed the same way. Dilutions of input were used for standard curve and calculations. 10 Jurkat cells were washed twice with PBS and resuspended in 800 μl of RNA-pull-down buffer (0.5% NP-40, 20 mM HEPES pH 7.8, 100 mM KCl, 0.2 mM EDTA supplemented with RNase inhibitor (NEB) and protease inhibitor (Sigma). Lysates were incubated with an in-vitro transcribed biotinylated CYTOR anti-sense probe (synthesized by IDT), and reactions were pulled down with streptavidin beads. Beads were resuspended in 3 times of their original volume of DNase buffer (100 mM NaCl and 0.1% NP-40), and protein was eluted with a cocktail of 100 μg/ml RNaseA (Sigma) and 0.1 U/μl RNaseH (Epicenter) and 100 U/ml DNase I (Invitrogen) at 37°C for 30 minutes. Eluted proteins were subjected to western blotting with indicated antibodies. Non-specific IgG served as control. Biotinylated scrambled RNA was used as a control for RNA-IP. 7SK RNA confirmed association with P-TEFb. Input is 5% of the total cell lysate . The cytosolic extracts were prepared by resuspending 3x10 cells in 500μl of Buffer A (10 mM KCl, 10 mM MgCl2, 10 mM HEPES, 1 mM EDTA, 1 mM DTT, 0.1% PMSF, and EDTA-free complete protease inhibitor cocktail (Roche) with 0.5% NP-40 for 10 minutes on ice. The nuclei were spun down at 5,000 g for 5 minutes, and the supernatant was saved as the cytosolic extract (CE). The nuclei were washed once with 200 μl of Buffer A with 0.5% NP-40 and re-pelleted. The nuclei were resuspended in 100 μl of Buffer B (450 mM NaCl, 1.5 mM MgCl2, 20 mM HEPES, 0.5 mM EDTA, 1 mM DTT, 0.1% PMSF, and EDTA-free complete protease inhibitor cocktail (Roche) and incubated on ice for 60 minutes. The lysates were clarified by centrifugation at 20,000g for 10 minutes to prepare nuclear extract (NE). RNA from nuclear or cytoplasmic was extracted with Trizol, and RNA was reverse transcribed using a qPCRBIO kit (PCPbiosystems), and qPCR was performed using indicated primers against CYTOR or 7SK ncRNA. CYTOR and 7SK RNA levels were normalized to 7SL RNA in each of the fractions and conditions. For identifying ncRNAs that are differentially expressed upon T cell stimulation in cells that carry active HIV (GFP+) versus latent HIV (GFP-), J-Lat 6.3 cells were stimulated with P/I and sorted based on their GFP expression (n = 4). For analysis of transcriptome upon CYTOR depletion cells, primary CD4+ T cells were stimulated with CD3/CD28 beads and then subjected to CYTOR KD by transducing cells with lentivirus that drive the expression of shRNA that target CYTOR (n = 3). CYTOR overexpression was obtained by transducing stimulated cells with lentivirus that drive CYTOR expression. RNA was purified utilizing the RNeasy Mini kit (QIAGEN) according to the manufacturer’s instructions. The integrity of the isolated RNA was tested using the Agilent High Sensitivity RNA Kit and Tapestation 4200 at the Genome Technology Center at the Faculty of Medicine Bar-Ilan University. Total RNA was used for mRNA enrichment by using the NEBNext mRNA polyA Isolation Module (NEB; E7490L), and libraries for Illumina sequencing were performed using the NEBNext Ultra II RNA kit (NEB; E7770L). Quantification of the library was performed using dsDNA HS Assay Kit and Qubit 2.0 (Molecular Probes, Life Technologies), and qualification was done using the Agilent D1000 Tapestation Kit and Tapestation 4200. 150 nM of each library was pooled together and was diluted to 4nM according to NextSeq manufacturer’s instructions. 1.6 pM was loaded onto the Flow Cell with 1% PhiX library control. Libraries were sequenced on an Illumina NextSeq 500 instrument, 75 cycles of single-read sequencing. For analysis: Quality Control was conducted by evaluating the quality of the FASTQ files using ’Fastqc’ (v0.12.1). Subsequently, the samples underwent quality trimming via the ’Fastp’ software (v0.23.3). To identify potential contaminations, ’fastq-screen’ software was applied. Read alignment involved aligning the reads to the Human GRCh38 genome (Ensembl release 110) using ’STAR’ software (v2.7.10b). Read assignments to coding regions were determined using the ’SubRead’ package (’FeatureCounts’ v2.0.6). Finally, BAM files were sorted using ‘samtools’. Differential expression analysis was conducted in DAVID. PCA was performed to evaluate data dispersion, revealing a batch effect among samples collected on different dates. A correction was implemented in the DESeq2 model to minimize this batch effect. Finally, the indicated groups were compared (without including the control in the model), identifying 90 DEGs. The selected cutoff values were an adjusted p-value <0.05 and a fold-change > 2. For enrichment analysis, the DAVID program was employed to identify enriched pathways and terms associated with the selected genes . These quality control and data analysis steps ensure the reliability and accuracy of the RNA-seq analysis. Primers on the HIV promoter: NFκB forward: 5’ - AGGTTTGACAGCCGCCTA -3’ NFκB Reverse: 5’ - AGAGACCCAGTACAGGCAAAA -3’ gapdh Forward: 5’ - AGCCACATCGCTCAGACAC -3’ gapdh Reverse: 5’ - GCCCAAACGACCAAATCC -3’ Primers for CYTOR: Forward: 5’- AACTTGCCAGCCTCCATC; Reverse: 5’- GAGCTTCCTGTTTCATCTCCC Primers for 7SK: Forward; 5‘- GAGGGCGATCTGGCTGCGACAT Reverse: 5‘- ACATGGAGCGGTGAGGGAGGAA Statistical evaluation was performed with GraphPad Prism 7 using two-way ANOVA with no correction for multiple comparisons. Number of independent data points refers to biological replicates. Each data point, as mentioned in the figure legends, represents the mean of 3–4 independent experiments with the errors calculated based on mean ± SD. Differences were considered statistically significant and denoted as ***p≤0.05; n.s., not significant. We would also like to thank people in the Taube lab who read the manuscript. Also, we thank Dr. Liron Levin, Dr. Aviad Sivan, and Mr. Yehuda Baruch for helping with the bioinformatic analysis. All relevant data can be found at NIH GEO - accession number GSE254771. All other data are within the manuscript and its Supporting information files. This work is supported by the Deutsche Forschungsgemeinschaft (DFG) - Projektnummer 508136175 – FA 378/25-1 to OTF and RT) and the German Centre for Infection Research (DZIF) (TTU 04.820 – HIV reservoir to OTF). OTF is a member of the CellNetworks cluster of excellence (EXC81). Additional funding for RT is from the Bi-National Science Foundation (BSF) - 2021273 to RT and JS and the National Institute of Health -NIH-R21 – 5R21AI170195 for RT and KF. KF is also supported by NIH R01AI167778, and Gilead Mentored Scientist Award from the UCSF AIDS Research Institute (ARI) and a Boost award from the NIH-funded UCSF-Bay Area Center for AIDS Research (P30 AI027763). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. All relevant data can be found at NIH GEO - accession number GSE254771. All other data are within the manuscript and its Supporting information files. |
PMC12118871 | PCYT2 overexpression induces mitochondrial damage and promotes apoptosis in hepatocellular carcinoma cells | Phosphatidylethanolamine cytidyltransferase 2 (PCYT2) is commonly regarded as the rate-limiting enzyme in eukaryotic phosphatidylethanolamine synthesis. However, the role of PCYT2 in the development of hepatocellular carcinoma (HCC) unknow. In this study, the role of PCYT2 overexpression in the development of HCC was examined by culturing HepG2 cells. We compared the expression levels of PCYT2 in L02 cells and HepG2 cells. Then, the HepG2 cells were infected with the lentivirus, establishing PCYT2 overexpression cell models. The proliferation, migration, and apoptotic abilities of PCYT2 overexpression in HepG2 cells was observed using western blotting, CCK-8 assay, Transwell assay, wound healing, and plate cloning methods. Based on this overexpression model, we determined the mitochondrial function and lipid content of HepG2 cells using lipidomics. CDP-ethanolamine (CDP-Etn), a downstream product of PCYT2, was added to the HepG2 cells, inhibiting their proliferation and migration. BALB/c female nude mice inoculated with subcutaneously transplanted tumors were used to explore the role of PCYT2. The results of the in-vitro experiments, shown that the expression of PCYT2 in normal hepatocytes was higher than that in HCC cells, and addition of CDP-Etn and PCYT2 overexpression inhibited the proliferation and migration of HCC cells, promoted the apoptosis of HCC cells, and caused mitochondrial damage. The results of in vivo experiments demonstrated that the tumor volume in the PCYT2 overexpression group was significantly smaller than that in the blank control group. Thus, PCYT2 overexpression inhibits the development of HCC, and its mechanism may be related to the impairment of mitochondrial function.Hepatocellular carcinoma (HCC) is an exceedingly fatal malignancies, with mortalities that approximate the incidence rates worldwide . HCC is usually diagnosed at advanced stages owing to late symptom manifestations with limited therapeutic options, leading to ineffective intervention and poor prognosis . An increasing number of studies have focused on the progression, pathological features, and prognosis of liver cancer [3–5]. HCC epidemiology is rapidly evolving, one of the most common causes is non-alcoholic fatty liver disease , further proving that lipid metabolism plays a crucial role in HCC occurrence. Therefore, the identification of novel therapeutic targets is urgently needed to improve the treatment of patients with HCC . Phosphatidylethanolamine (PE), also known as cephalin, is the most abundant lipid in the cytoplasmic layer of cell membranes and is involved in cellular processes such as membrane fusion , autophagy and apoptosis [9–11]. For eukaryotic PE in vivo, two main synthetic pathways exist : including the Kennedy pathway of CDP-ethanolamine(CDP-Etn) and mitochondrial phosphatidylserine decarboxylation pathway . Phosphatidylethanolamine cytidyltransferase 2(PCYT2) is the rate-limiting enzyme of the CDP-Etn pathway. Previous studies have shown that PCYT2 is highly specific, and is present only in the rough endoplasmic reticulum of eukaryotes . P-Eth is then catalyzed by PCYT2 to form CDP-Etn, leading to PE synthesis . Generally, PCYT2 expression is reduced in various epithelial-derived cancer cell lines compared to normal cells . Compared epithelial-derived cancer cell lines PCYT2 activity with that of breast epithelial cells MCF-10A showed that its PCYT2 activity was inhibited in breast cancer cells (MCF-7) . PCYT2 expression was significantly reduced in invasive human metastatic colon cancer cells compared to that in primary tumor cells , and previous studies have shown that PCYT2 knockdown under nutrient-rich conditions significantly facilitated the proliferation of HeLa and T98G cells and promoted in vivo tumor growth. The inhibition of PCYT2 increases P-Etn levels in cancer cells and stimulates tumor growth . However, in organoid models, PCYT2 knockdown inhibits cell growth . Despite these findings, no evidence exists that suggests that PCYT2 expression is associated with HCC occurrence and development. This study aimed to investigate the role of PCYT2 in human hepatocellular carcinoma cells (HepG2) by inhibiting their proliferation, invasion, and migration abilities and promoting cell apoptosis. CDP-Etn (90756) was purchased from Sigma (Darmstadt, Germany).The ATP assay kit (S0026) and BCA protein assay kit (P0010S) were purchased from Beyotime (Shanghai, China), and RPMI 1640 medium (SH30027.01) was purchased from Gibco (Waltham, MA,USA). Fetal bovine serum(FBS #11012–8611) was purchased from TIANHANG (Zhejiang, China). PCYT2 antibody (ab135290) was provided by Abcam (Cambridge, UK); BAX (#5023), Bcl-2 (#3498), cleaved caspase-3 (#9661) and β-actin (#3700) were purchased from Cell Signaling Technology (Danvers, MA, USA); goat anti-rabbit (E-AB-1034) and goat anti-mouse (E-AB-1035) secondary antibodies were provided by Elabscience (Shanghai, China). L02 cells (normal human liver cells) were cultured in RPMI1640 medium, and HepG2 were cultured in DMEM containing 10% FBS and 1% penicillin-streptomycin. The above two cell types of cells were cultured at 37°C in a humidified incubator atmosphere containing 5% CO2. Lentiviral plasmids overexpressing PCYT2 (LV/In) and negative control (NC/In) were purchased from GENECHEM (Shanghai, China). HepG2 cells were infected with LV/In or NC/In, 48 h later, the cells were screened with DMEM medium containing puromycin (2 µg/mL) for approximately 2 weeks. The expression level of PCYT2 was detected using western blotting after 3–4 generations to confirm PCYT2 expressing up-regulation. RIPA buffer was used to extract total cellular protein. Protein samples were electrophoresed using 9–11% SDS-PAGE, then transferred to PVDF membranes (Merck, Darmstadt, Germany) that were blocked with 5% skimmed milk in TBST, and incubated overnight utilizing primary antibodies, including anti-PCYT2 (1:250), anti-β-actin (1:3000), anti-BAX (1:1000), and anti-Bcl-2 (1:1000). Subsequently, samples were incubated 1–2 h using an appropriate peroxidase-linked secondary antibodies. Consequently, employing β-actin we normalized the protein levels. Images were visualized using the Chemi-Doc MP system (Bio-Rad). HepG2 cells were seeded into 96-well plates and incubated with a Cell Counting kit (CCK-8) solution for 40 min at 37°C. Absorbance was measured at 450 nm using a Spectra MAX M5 microplate spectrophotometer to detect cell viability. To determine cell migration, the Transwell assay was performed by adding 500 µL of complete medium to a 24-well plate, then placing the Transwell chamber on the plate. The cell suspension was prepared by adding basal medium and 200 µL (3 × 10 cells/200 µL) to each top chamber. Subsequently, the cells were incubated at 37°C for 1 d. The cells that successfully migrated and attached to the surface of the underlying membrane were fixed with paraformaldehyde and stained with 0.1% crystal violet. Five to six fields of view (original magnification: 200×) were randomly selected for cell counting under a light microscope. Dilute the Matrigel with basal medium (1:8), lay it flat on a Transwell membrane, and incubate at 37°C for 4 h. The subsequent experimental steps were identical to those used for the migration assay, including cell culture, paraformaldehyde fixation and 0.1% crystal violet staining. Two milliliter of cell suspension was added to a six-well plate (3 × 10 cells/well) and cultured at 37°C in a humidified incubator atmosphere at 5% CO2; 24 h later, the center of each well was scratched with a 100 µL plastic micropipette and the medium was substituted with basal medium. Each well was imaged under a 200 × light microscope at three randomly selected identical locations after culturing for 24 and 48 h. Finally, we measured cell migration ability by comparing the distance between the edges of each wound at 24 and 48 h. A cell suspension (2 mL, 5 × 10 cells/mL) was added to each well of a six-well plate and incubated for 2 weeks. When the clones were visible to the naked eye, they were washed twice with phosphate-buffered saline (PBS), and the cells were fixed using –4% paraformaldehyde for 10 min. Paraformaldehyde was washed away and 0.1% crystal violet staining was performed for 15 min. The clones were washed twice with PBS, dried at room temperature and the number of colonies was statistically analyzed. Cellular ATP levels were measured using an ATP kit (Beyotime). Sample tubes containing 20 μL of sample or standards were placed in a luminometer (SuPerMax 3100) and rapidly mixed with a micropipette. After > 2 s, relative light unit values were measured. The tiled cells (1 × 10) were removed, immersed in PBS, and the cell surface was rinsed. Then 2.5% of pre-cooled glutaraldehyde was added to the tiled cells at 4°C, fixed at 4°C for 2 h or overnight, aspirated, and soaked in PBS twice for 10 min each. Finally, ethanol gradient dehydration (30, 50, 70, 80, 90, 100%), critical point drying, coating, and electron microscopy was performed to observe mitochondrial morphology. Five-week-old BALB/c female nude mice were purchased from Jiangsu Jicui Pharmaceutical Biotechnology Co, passed SPF level training and assessment, and were routinely reared using standard SPF conditions. PCYT2 overexpressed HepG2 cells and control cells were subcutaneously injected into the left and right sides of the nude mice (approximately 1 × 10 cells on each side, n = 6 tumors in each group). Following 20–25 d, the tumors achieved a certain size and the micewere anesthetized with isopentane and sacrificed; the tumors were collected for follow-up evaluation. The animal experiments were approved by the Animal Ethics Committee of Anhui Medical University and conducted in accordance with the guidelines for the care and use of laboratory animals. Data were analyzed using GraphPad Prism software (version 8.0), and the results were expressed as mean ± standard deviation (SD). T-test was performed to determine the significant of differences between two groups. A one-way analysis of variance was used for comparison across groups. Statistical significance was set at P < 0.05. The Cancer Genome Atlas (TCGA) database, Cancer Research Project, which is a collaboration between the National Cancer Institute and National Human Genome Research Institute, provides a large, free reference database for cancer research by collecting and collating cancer-related data. By analyzing the TCGA database, we attempted to determine the relationship between the PCYT2 expression level and HCC. Our findings demonstrated that overall survival of patients with HCC was closely correlated with PCYT2 expression. We observed that the survival percentage of patients gradually decreased over time; however, patients with high PCYT2 expression levels had a higher survival percentage than those with low PCYT2 expression (Fig 1A). This indicates that high PCYT2 expression levels are beneficial for the overall survival of patients with HCC. Western blotting was conducted on cultured L02 and HepG2 cells to verify their PCYT2 expression level (Fig 1B and 1C), we found that PCYT2 expression was higher in L02 cells than in HepG2 cells. To verify the effect of PCYT2 on HCC development, we transfected HepG2 cells with a lentivirus and performed western blotting, successfully establishing PCYT2 overexpression in HepG2 cells (Fig 2A). The expression levels of Bcl-2 (an anti-apoptotic factor), Bax (an apoptogenic factor) and cleaved-caspase-3(an apoptogenic factor)(S1 Fig.) were detected using western blotting, and the results revealed that the LV group exhibited increased HepG2 cell apoptosis (Fig 2B–2D). The CCK-8 assay indicated that PCYT2 overexpression significantly reduced HepG2 cells viability (Fig 2E). Utilizing the Transwell assay, we discovered that the HepG2 cells invasion and migratory ability was suppressed post-lentiviral transfection (Fig 2F–2H). The scratch wound healing assay also determined that the migratory ability of the LV group was inhibited (Fig 2I). Additionally, PCYT2 overexpressing HepG2 cells formed less colonies than the negative control group (Fig 2J and 2K). In summary, we demonstrated that PCYT2 overexpression inhibited the proliferation, invasion, and migration of HepG2 cells. (A–D) The level of PCYT2 (A) as well as the protein expression of Bax and Bcl-2 was measured per group (NC, normal control; LV, lentivirus transfection to over-express PCYT2) using western blotting and representative protein quantification (n = 3 per group). (E) The proliferation ability of each group was detected using CCK-8 (n = 6 per group). (F–H) A Transwell assay was conducted per group to investigate the effects of cellular migration and invasion. Magnification: 100 × . (I–K) The scratch wound healing and colony formation assays of cells post-PCYT2 overexpression. The magnification for scratch wound healing assay is 100 × . Data are presented as means ± SD. *P < 0.05, **P < 0.01, ***P < 0.001. During cell apoptosis, the mitochondria undergoes many changes, such as respiratory chain depolymerization, oxidative phosphorylation, decreased ATP synthesis, and increased reactive oxygen species. We measured the concentration of ATP in the cells of the NC and LV groups (Fig 3A), and photographed the mitochondria using transmission electron microscopy (Fig 3B). The results showed that the LV group exhibited mitochondrial damage and its ATP content was lower than that in the NC group. Compared to the NC group, in the LV group, the mitochondria was swollen and its number was significantly reduced (Fig 3B), further suggesting that PCYT2 overexpression promotes apoptosis. (A) The ATP levels of each group (NC, normal control; LV, lentivirus transfection to over-express PCYT2) were detected using commercial reagent kits (n = 3 per group). (B) Images of mitochondrial damage post-PCYT2 overexpression were captured via transmission electron microscopy. Data are presented as means ± SD. **P < 0.01. PCYT2’s inhibition of cellular proliferation and migration was mediated by altering downstream metabolite levels, CDP-Etn was introduced to HepG2 cells, and cell viability was measured using CCK-8. Therefore, HepG2 + CDP-Etn cells have lower cell viability than HepG2 cells (Fig 4A). In addition, TUNEL staining was performed to observe apoptosis in HepG2 and HepG2 + CDP-Etn cells; the results indicated that CDP-Etn promoted apoptosis in HepG2 cells, affirming those of the CCK-8 assay (Fig 4B and 4C). Furthermore, Transwell assay (Fig 4D) and scratch experiment (Fig 4E) was conducted to detect cell migration, suggesting that supplementation with CDP-Etn inhibited cell migration. Subsequently, plate cloning showed that fewer colonies formed in HepG2 + CDP-Etn cells than in HepG2 cells (Fig 4F and 4G). Thus, we confirm that CDP-Etn inhibited the proliferation and migration ability of HepG2; that is, PCYT2 inhibited proliferation and migration by altering downstream PE metabolism. (A) The proliferation ability of HepG2 cells with or without CDP-Etn supplementation were detected using CCK-8 (n = 3 per group). (B–C) HepG2 cells underwent TUNEL staining, and the nucleus underwent DAPI staining. Magnification: 100 × (D–G) Transwell (D), scratch wound healing (E) and colony formation assays (F–G) were performed post-CDP-Etn supplementation. Magnification for Transwell and scratch wound healing assay is 100 × . Data are presented as means ± SD. *P < 0.05, **P < 0.01. To investigate whether PCYT2 affects hepatocarcinogenesis and development in vivo, we subcutaneously implanted PCYT2-overexpressing HepG2 cells and blank control HepG2 cells into 12 nude BALB/c mice. The tumor volume was measured at 2 d intervals(Fig 5C). After 20 d, the mice were anesthetized with isoflurane to observe the tumor volume using an in vivo imaging system (Fig 5A). The nude mice were sacrificed and tumors were removed for imaging (Fig 5B) and weighing (Fig 5D). The diameter of the largest tumor was 1.9 cm. The results showed that the fluorescence intensity of the tumors in the LV group was weaker than that in the NC group and the tumor volume and weight in the NC group were greater than that in the LV group, suggesting that PCYT2 overexpression inhibit tumor growth in vivo. (A–B) Representative image of in vivo imaging system (A) and tumors isolated from mice xenograft model (B), which were established by subcutaneously implanting PCYT2-overexpressing HepG2 cells (LV) and blank control HepG2 cells (NC). (C, D) Cancer volumes were measured every other day, and the average tumor weights in each group were measured. n = 6 per group, data are presented as means ± SD. ***P < 0.001, ****P < 0.0001. PCYT2 is a rate-limiting enzyme in PE synthesis that is commonly used in the study of obesity-related diseases, such as non-alcoholic fatty liver disease and type 2 diabetes [12,20–22]. Recently, interest in the role of PCYT2 in cancer has been growing . Previous studies show that PCYT2 has different roles in various cancers and cancer settings. For instance, in metastatic colorectal cancer (CRC), PCYT2 is significantly downregulated and functions as a tumor metastasis inhibitor . In human breast cancer cells (MCF-7), the level of PCYT2 in cancer cells is elevated in response to the stressful environment . Our findings show that PCYT2 expression is abnormally downregulated in HepG2 cells, which is consistent with that of previous studies where in PCYT2 expression was downregulated in MCF-7 and invasive human metastatic CRC . Although PCYT2 regulates several human cancers , its role in HCC cells remains unknown. Based on the literature and our findings, PCYT2 in human cancers appears to have a consistent expression profile in human cancers, irrespective of tumor origin or location . Regarding the mechanism whereby PCYT2 influences cancer development, a previous report showed that PCYT2 downregulation-induced phosphoethanolamine (PEtn) accumulation correlated with tumor growth under nutrient starvation, thereby PCYT2 overexpression reduced PEtn levels and tumor growth . However, in the present, we found that CDP-Etn supplementation inhibited HCC migration, invasion and proliferation. In our previous study, we showed that the levels of BAX and cleaved caspase-3 were significantly increased whereas Bcl-2 was significantly reduced in the livers of type 2 diabetic mice and L02 cells after stimulation with high glucose and free fatty acids (HG&FFA)(12). CDP-Etn (100 μM) protected cells from HG&FFA-induced apoptosis by reducing BAX and cleaved caspase-3 levels as well as and increasing Bcl-2 levels . Whether PCYT2 downregulation induced PEtn accumulation contributes to HCC development further study. Increasing evidence suggests that PCYT2 is aberrantly expressed in various models of liver disease and may predict clinical outcomes in patients. PCYT2 is instrumental in the deregulation of these processes leading to the development of obesity, insulin resistance, liver steatosis and dyslipidemia . CDP-Etn supplementation has been reported to alleviate PCYT2 deficiency engendering age-dependent and insulin-resistant non-alcoholic steatohepatitis to improve patient prognosis [20,29–31]. Chronic administration of peroxisome proliferators can increase the content of hepatic PC and PE for hepatomegaly and proliferation as well as cause liver cancer in rodents . Based on the current studies, we hypothesized that PCYT2 may be involved in the regulation of cellular processes in HCC. Therefore, we utilized in vivo and in vitro validation methods to assess the expression and mechanistic roles of PCYT2 in liver cancer cells. Herein, PCYT2 overexpression was determined to inhibit HCC cell proliferation, migration and invasion both in vitro and in vivo. And, the number and morphology of mitochondria in HCC cells overexpressing PCYT2 were significantly different from those in HCC cells without any treatment, such as a decrease in the number of mitochondria and swelling of the mitochondria. These changes suggest that PCYT2 affects the mitochondrial function of cells. Notably, when the HepG2 cells received CDP-Etn supplementation, their proliferation, migration, and invasion were inhibited in vitro. Notably, the ATP level decreased in HCC cells following the overexpression of PCYT2, and the cells were found to be accompanied by mitochondrial damage using transmission electron microscopy. However, previous reports have indicated that PCYT2 is present only in the endoplasmic reticulum of hepatocytes . Additionally, the phenotype of liver PCYT2 knockout mice showed no signs of liver injury, however, they experienced massive accumulation of liver triglycerides (TAG) . Therefore, we hypothesized that the influence of PCYT2 on mitochondrial function is mediated by metabolites such as TAG, DAG, and PE. However, further studies are needed to clarify whether the PCYT2 exerted inhibition of HCC cells alleviates mitochondrial damage. As understanding, the mechanism whereby PCYT2 overexpression causes mitochondrial damage will deepen our understanding of PCYT2 regulation in HCC cells. In conclusion, this study provided compelling data demonstrating the aberrant expression and functional role of PCYT2 in HepG2 cells. PCYT2 expression levels were lower in HepG2 than in L02. Furthermore, PCYT2 overexpression in HepG2 cells induced mitochondrial damage; inhibited proliferation, invasion, and migration; and promoted cell apoptosis. |
PMC1462997 | Derivation of normal macrophages from human embryonic stem (hES) cells for applications in HIV gene therapy | Many novel studies and therapies are possible with the use of human embryonic stem cells (hES cells) and their differentiated cell progeny. The hES cell derived CD34 hematopoietic stem cells can be potentially used for many gene therapy applications. Here we evaluated the capacity of hES cell derived CD34 cells to give rise to normal macrophages as a first step towards using these cells in viral infection studies and in developing novel stem cell based gene therapy strategies for AIDS. Undifferentiated normal and lentiviral vector transduced hES cells were cultured on S17 mouse bone marrow stromal cell layers to derive CD34 hematopoietic progenitor cells. The differentiated CD34 cells isolated from cystic bodies were further cultured in cytokine media to derive macrophages. Phenotypic and functional analyses were carried out to compare these with that of fetal liver CD34 cell derived macrophages. As assessed by FACS analysis, the hES-CD34 cell derived macrophages displayed characteristic cell surface markers CD14, CD4, CCR5, CXCR4, and HLA-DR suggesting a normal phenotype. Tests evaluating phagocytosis, upregulation of the costimulatory molecule B7.1, and cytokine secretion in response to LPS stimulation showed that these macrophages are also functionally normal. When infected with HIV-1, the differentiated macrophages supported productive viral infection. Lentiviral vector transduced hES cells expressing the transgene GFP were evaluated similarly like above. The transgenic hES cells also gave rise to macrophages with normal phenotypic and functional characteristics indicating no vector mediated adverse effects during differentiation. Phenotypically normal and functionally competent macrophages could be derived from hES-CD34 cells. Since these cells are susceptible to HIV-1 infection, they provide a uniform source of macrophages for viral infection studies. Based on these results, it is also now feasible to transduce hES-CD34 cells with anti-HIV genes such as inhibitory siRNAs and test their antiviral efficacy in down stream differentiated cells such as macrophages which are among the primary cells that need to be protected against HIV-1 infection. Thus, the potential utility of hES derived CD34 hematopoietic cells for HIV-1 gene therapy can be evaluated.Human embryonic stem cells (hES cells) show great promise for many novel cellular therapies due to their pluripotent nature . These cells have the capacity to give rise to mature cells and tissues that arise from all three germ layers during embryonic development [2-4]. Several pluripotent hES cell lines have so far been derived from the inner cell mass of human blastocysts and can be cultured indefinitely in an undifferentiated state [5-7]. Thus, these cells provide a renewable source of pluripotent stem cells from which many types of differentiated cells could be produced for experimental and therapeutic purposes. Cell differentiation protocols currently exist for the derivation of neurons, cardiomyocytes, endothelial cells, hematopoietic progenitor cells, keratinocytes, osteoblasts, and hepatocytes to name a few . In addition to providing for potential cellular replacement therapies, opportunities exist in programming hES cells to correct a genetic defect and/or to express a therapeutic transgene of interest. Using such approaches, many possibilities exist for treating a number of genetic and immune system disorders . Many novel applications can be foreseen for hES cells in infectious disease research. AIDS is a potential disease that can benefit from exploiting hES cells for cell replacement therapy as they have the capacity to differentiate into various hematopoietic cells. HIV continues to be a major global public health problem with infections increasing at an alarming rate . Given the present lack of effective vaccines and the ineffectiveness of drug based therapies for a complete cure, new and innovative approaches are essential. Gene therapy through intracellular immunization offers a promising alternative approach and possible supplement to current HAART therapy [12-14]. HIV mainly targets cells of the hematopoietic system, namely, T cells, macrophages, and dendritic cells . As infection progresses, the immune system is rendered defenseless against other invading pathogens and succumbs to opportunistic infections. There is a great deal of progress in the area of stem cell gene therapy for AIDS . A primary goal of many ongoing studies is to introduce an effective anti-HIV gene into hematopoietic stem cells [16-18]. As these cells possess the ability to self renew, they have the potential to continually produce HIV resistant T cells and macrophages in the body thus providing long term immune reconstitution. These approaches use CD34 hematopoietic stem cells for anti-HIV gene transduction via integrating viral vectors such as lentiviral vectors [16-18]. Lentiviral vectors have several advantages over conventional retroviral vectors since higher transduction efficiencies can be obtained and there is less gene silencing. The CD34 cells currently used for many therapies are primarily obtained from bone marrow or mobilized peripheral blood . Thus, CD34 progenitor cells are an essential ingredient for HIV gene therapy. In view of the need for CD34 cells for HIV gene therapy as well as for other hematopoietic disorders, if one can produce these cells in unlimited quantities from a renewable source, it will overcome the limitations of securing large numbers of CD34 cells for therapeutic purposes. In this regard, progress has been made in deriving CD34 cells from hES cells (hES-CD34). Different methods currently exist to derive CD34 cells from hES cells with varying efficiencies [20-27]. Recent reports have indicated the capacity of hES cell derived CD34 cells to give rise to lymphoid and myeloid lineages thus paving the way for utilization of these cells for hematopoietic cell therapy [20,27-29]. For the effective utilization of hES-CD34 cells for HIV gene therapy, a number of parameters need to be examined. First, one has to demonstrate that hES-CD34 cells can give rise to macrophages and helper T cells which are the main cells that need to be protected against HIV infection. Recent evidence has shown that hES-CD34 cells can give rise to myelomonocytic cells . However, thorough phenotypic or functional characterization of these cells is lacking. It is also not clear if these cells are susceptible to HIV infection. Similarly, although the hES-CD34 cells were shown to have lymphoid progenitor capacity, only B cell and natural killer (NK) cell differentiation has been examined so far . The capacity to generate T cells remains to be evaluated. With this background, as a first step, our primary goal in these studies is to examine the capacity of hES-CD34 cells to give rise to phenotypically and functionally normal macrophages and whether such cells are susceptible to productive HIV infection. Since lentiviral vectors have been shown to successfully transduce hES cells [30-33], we further investigated the ability of transduced hES cells to differentiate into transgenic macrophages that can support HIV-1 infection. Demonstration of HIV-1 productive infection in these cells will permit future efficacy evaluations of anti-HIV genes in this system. Here we show that normal and lentiviral vector transduced hES-CD34 cells can give rise to phenotypically and functionally normal macrophages that support HIV infection thus paving the way for many novel approaches to evaluate their potential for HIV gene therapy. Undifferentiated hES cell colonies grown in media supplemented with 4 ng/ml bFGF displayed normal morphology of pluripotent human embryonic stem cells with tight and discreet borders on the MEF feeder layers (Fig 1A). Similarly, lentiviral vector transduced hES cell colonies, also displayed normal morphology and growth characteristics (Fig 1A). As expected, the vector transduced colonies displayed green fluorescence due to the presence of the GFP reporter gene. When cultured on irradiated S17 mouse bone marrow stromal cells, both nontransduced and transduced hES cells developed into embryonic cystic bodies (Fig 1A). FACS analysis of single cell suspensions of the cystic bodies showed levels of CD34 cells which ranged from 7–15%. Figure 1B displays a representative FACS profile of hES-CD34 cells. Purified CD34 cells were later cultured in semi-solid methylcellulose medium to derive myeloid colonies. Both nontransduced (denoted as ES in figures) and vector transduced (denoted as GFP ES in figures) hES cell derived CD34 cells gave rise to normal myelomonocytic colonies similar to human fetal liver derived CD34 cells (denoted as CD34 in figures) (Fig 1A). When pooled colonies were cultured further in liquid cytokine media for 12–15 days for differentiation, the cells developed into morphologically distinct macrophages (Fig 1A). When compared, the morphology of macrophages derived from all stem cell progenitor populations appeared similar. These results were found to be consistent in replicative experiments. The transgene GFP expression was also maintained during the differentiation of hES cells into mature macrophages. GFP expression in cystic body derived CD34 cells was around 80% (data not shown) with similar levels seen in differentiated macrophages (Fig 2). Derivation of macrophages from lentiviral vector transduced and normal hES cells. A) Transduced and non-transduced H1 hES cells were cultured on mouse S17 bone marrow stromal cell layers to derive cystic bodies. Cystic body derived CD34 cells were purified by positive selection with antibody conjugated magnetic beads and placed in methocult media to obtain myelomonocytic colonies. Pooled colonies were cultured in liquid cytokine media supplemented with GM-CSF and M-CSF to promote macrophage growth. For comparison, fetal liver derived CD34 cells were cultured similarly to derive macrophages. Representative ES cell colonies, cystic bodies, methocult colonies, and derivative macrophages are shown with GFP expressing cells fluorescing green under UV illumination. B) Representative FACS profile of hES cell derived CD34 cells stained with PE conjugated antibodies. Percent positive CD34 cells are shown with isotype control shown in the left panel. Phenotypic FACS analysis of hES cell derived macrophages. A) Macrophages derived from transduced and nontransduced hES CD34 and fetal liver CD34 cells were stained with antibodies to CD14, HLA-DR, CD4, CCR5, and CXCR4 and the expression of these surface markers was analyzed by FACS. B) Isotype controls for PE and PE-CY5 antibodies. Percent positive cells are displayed in the plots for each respective cell surface marker staining. Dot plots are representative of triplicate experiments. Macrophages play a critical role in immune system function and are also major target cells for many viral infections including HIV-1. Distinct surface phenotypic markers exist on these cells and, thus far, there has been no thorough evaluation of hES cell derived macrophages. Therefore we analyzed hES cell derived macrophages for the presence of characteristic cell surface markers and compared these to the phenotypic profile displayed on fetal CD34 cell derived macrophages. The surface markers analyzed were CD14, a monocyte/macrophage specific marker, HLA-DR (a class II MHC molecule found on antigen presenting cells), CD4, the major receptor for HIV-1 infection, and CCR5 and CXCR4, chemokine receptors which are critical coreceptors essential for HIV-1 entry. EGFP expression was also analyzed to determine the levels of transduction and any transgene silencing that may occur during differentiation. Fetal liver (CD34), nontransduced (ES), and vector transduced (GFP ES) hES cell derived macrophages were all positive for the monocyte/macrophage marker CD14 (99.3%, 88.7%, and 99.2%, respectively) (Fig 2A). However, the mean fluorescent intensity (MFI) was found to be lower on hES cell derived macrophages. Surface expression of HLA-DR was observed at similar levels between macrophages derived from fetal liver CD34 cells (99.6%), nontransduced hES cells (92.8%), and transduced hES cells (98.2%) (Fig 2A). CD4 levels were comparable for all stem cell derived macrophages (99.2%, 83.3%, and 88.7%, respectively) (Fig 2A). CCR5 and CXCR4 cell surface expression was also observed for fetal liver CD34 cell (99.6% and 99.3%), nontransduced hES cell (91.9% and 92.6%), and transduced hES cell (98.9% and 99.3%) derived macrophages (Fig 2A). As compared to fetal liver CD34 cell derived macrophages, hES cell derived macrophages displayed a higher level of expression of CXCR4. Isotype controls for both PE and PECY5 stains are shown in Fig 2B. The above phenotypic data are representative of triplicate experiments. The antigen presenting cell surface specific marker HLA-DR (MHC II) on normal macrophages is critical for presenting antigen to CD4 T cells. A second co-stimulatory molecule, B7.1 is present at low basal levels on resting macrophages and is necessary to activate T cells. Its expression is elevated upon activation with certain stimuli such as LPS. Our results of LPS stimulation of respective macrophages have shown upregulation of B7.1 with values for fetal liver CD34 cell (CD34) (27.9% to 75.4%) nontransduced (ES) (17.8% to 49.4%) and transduced (GFP ES) (35.6% to 65.7%) hES cell derived macrophages (Fig 3A). These values represent a significant upregulation of B7.1 for all three macrophage populations. Functional analysis of hES cell derived macrophages for B7.1 costimulatory molecule upregulation and phagocytosis of E. coli particles: A) Mature macrophages were stimulated with LPS to determine B7.1 upregulation. Twenty-four hours post-stimulation, macrophages were labeled with a PE-CY5 conjugated anti-B7.1 antibody and analyzed by FACS. B7.1 upregulation data are representative of triplicate experiments. Isotype control is shown in the left panel. B) To assess phagocytic function, E. coli Bioparticleswere added directly to the cultured macrophages. Twenty four hours post-addition, cells were analyzed by FACS. Percent positive cells are displayed in the plots for each experiment. These data are representative of triplicate experiments. Another important function of macrophages is their ability to phagocytose foreign material and present antigenic peptides on their cell surface. To evaluate phagocytic function, fluorescently labeled E. coli Bioparticleswere added to macrophage cultures followed by FACS analysis. Nontransduced (94.6%) as well as lentiviral vector transduced (98.7%) hES cell derived macrophages were found to be capable of phagocytosing the Bioparticlesin comparison to fetal liver CD34 cell derived macrophages (95.8%) (Fig 3B). These values are representative of triplicate experiments. Magi-CXCR4 cells with no phagocytic capacity were used as non-phagocytic cell controls and similarly exposed to E. coli Bioparticles(Fig 3B). No uptake of the bacteria could be seen. Thus, uptake of E. coli Bioparticlesby macrophages is indicative of active ingestion. Macrophages, as effector cells, play a key role in the inflammatory response. Activated macrophages secrete various cytokines, two of the major ones being IL-1 and TNF-α. To determine if hES cell derived macrophages have such a capacity, cells were stimulated with LPS. On days 1, 2, and 3 post-stimulation, culture supernatants were analyzed by ELISA to detect IL-1 and TNF-α. As seen in figure 4A, there were no significant differences in IL-1 secretion between the three sets of macrophages. Similarly, nontransduced and transduced hES cell derived macrophages were also capable of TNF-α secretion upon LPS stimulation. However, levels of the respective cytokines detected were slightly lower than those from fetal liver CD34 cell derived macrophages (Fig 4B). The values of cytokine secretion levels represent triplicate experiments. Cytokine IL-1 and TNFα secretion by stimulated hES cell derived macrophages: Macrophages derived from transduced and nontransduced hES and fetal liver CD34 cells were stimulated with 5 μg/ml LPS. On days 1, 2, and 3 post-stimulation, supernatants were collected and assayed by ELISA for (A) IL-1 and (B) TNFα. Experiments were done in triplicate. The above data have shown that hES cell derived macrophages are very similar to normal human macrophages based on phenotypic and functional analysis. In addition to being important cells of the immune system, macrophages are among the major target cells for certain viral infections, particularly for HIV-1. We wanted to determine if hES cell derived macrophages were susceptible to HIV-1 infection compared to standard macrophages. In these studies, we only used an R5-tropic strain of HIV-1 since macrophages are natural targets for this virus. Our results from challenge studies of these cells clearly indicated the capacity of hES cell derived macrophages in supporting a productive infection. Levels of virus increased up to 15 days similar to non-hES derived macrophages showing that the initial viral input was amplified in productive viral infection. However, the levels of viral yield were found to be slightly lower for the ES cell derived macrophages. In the case of GFP-ES macrophages, there was a decline in viral titer. This could be due to possible lower numbers of cells present in the initial cultures. As a first step towards the use of hES cells for hematopoietic stem cell and HIV gene therapies, we have shown here that phenotypically and functionally normal macrophages could be derived from hES-CD34 cells. Both non transduced and lentiviral vector transduced hES cells were found to be capable of generating CD34 cells that give rise to macrophages which could support productive HIV-1 infection. Current sources of CD34 cells consist of human bone marrow, cytokine mobilized peripheral blood, fetal liver, and cord blood . However, the number of cells that can be obtained for manipulations is not unlimited. Therefore, deriving CD34 cells for therapeutic and investigative purposes from hES cells with unlimited growth potential has the advantage of a consistent and uniform source. The ability to obtain phenotypically normal and functionally competent macrophages from hES cells is important to evaluate their potential therapeutic utilities in the future. Additionally, testing of transgenic hES cells derived via lentiviral vector gene transduction is also helpful to determine the stability of the transgene expression and their capacity for differentiation into end stage mature cells such as macrophages. Based on these considerations, both non- transduced and lentiviral vector transduced hES cells were evaluated for their capacity to give rise to CD34 progenitor cells. In colony forming assays using semisolid methylcellulose medium, the morphology of myelomonocytic colonies derived from hES CD34 cells appeared similar to that of fetal liver CD34 cells. When subsequently cultured in cytokine media that promotes macrophage differentiation, morphologically normal macrophages were obtained with hES-CD34 cells similar to that of fetal liver CD34 cells. At higher magnification, the macrophages displayed flat projecting cellular borders with fried egg appearance with distinct refractory lysosomal granules in the cytoplasm (data not shown). Lentiviral vector transduced hES cells also did not display any abnormal growth or differentiation characteristics as compared to nontransduced hES-CD34 cells indicating no adverse effects due to vector integration and expression. Transduced cells gave rise to cystic bodies with similar CD34 cell content and profiles upon development. The transduced hES-CD34 cells also gave rise to apparently normal macrophages that expressed the transgene as shown by GFP expression. These results are consistent with those of others that showed normal differentiation of hES cells to other cell types following lentiviral transduction . A requirement for successful cellular and HIV-1 gene therapy is that mature end stage cells derived from CD34 progenitor cells be phenotypically and functionally normal to maintain and restore the body's immunological function. Accordingly, hES cell derived macrophages were evaluated to determine if they met these criteria. Macrophages display distinct cell surface markers upon end stage differentiation. To determine whether hES cell derived macrophages display these surface markers, FACS analysis was performed to detect the presence of CD14, HLA-DR (MHCII), CD4, CCR5, and CXCR4. As observed in Fig 2A, both nontransduced and transduced hES cell derived macrophages expressed all of these markers with some differences in their levels of expression. HLA-DR, CD4, and CCR5 expression profiles were comparable between all cell types analyzed. Even though all cell types analyzed stained positive for CD14, relative expression of CD14 was slightly lower on hES cell derived macrophages compared to fetal liver CD34 cell derived macrophages. On the contrary, the levels of CXCR4, a chemokine receptor involved in cellular homing, were found to be higher on hES-CD34 cell derived macrophages. This may be due to inherent differences in the cell types and/or due to their physiological state at the time of harvest . Additional hES cell lines need to be evaluated in the future to establish if these differences are consistent. A major functional role of macrophages in vivo is their ability to serve as professional antigen presenting cells. During this process macrophages present antigen peptide fragments complexed with both classes of MHC molecules and deliver a costimulatory signal through the expression of B7 molecules. Upon stimulation with LPS, hES-CD34 cell derived macrophages had shown upregulation of the costimulatory molecule B7.1 similar to cells derived from fetal liver. Furthermore, the hES-CD34 cell derived macrophages also showed a normal capacity to ingest foreign particles in phagocytosis assays using E.coli Bioparticles. In addition to antigen presentation and phagocytosis, macrophages also play a critical role in inflammation and secrete cytokines in response to external stimuli. When exposed to LPS, the hES-CD34 cell derived macrophages secreted two important cytokines IL-1 and TNF-α similar to that of fetal liver derived cells. The above data has established that phenotypically and functionally normal macrophages could be derived from hES-CD34 cells. Macrophages in addition to playing important physiological roles are also major cell targets for certain viral infections, particularly HIV-1. Here we evaluated the susceptibility of hES-CD34 cell derived macrophages to be productively infected with HIV-1. Similar to that of fetal liver CD34 cell derived cells, the hES-CD34 macrophages also supported HIV-1 infection although the levels of viral yield differed somewhat. However this should not be a major concern for testing anti-HIV genes in these cells. In all the above experiments, the vector transduced transgenic macrophages also behaved similarly to that of nontransduced cells showing that they were also physiologically normal. The lack of vector toxicity on cellular maturation is encouraging for future work with transduced hES-CD34 cells to derive other important differentiated cells like T cells and dendritic cells relevant for HIV studies. Although there are numerous studies on hES cell differentiation into many important end stage mature cells, systematic work on hES cell hematopoietic differentiation and thorough characterization of end stage mature cells that participate in critical immune responses has just begun [21,27-29]. Our current results established that physiologically normal macrophages could be derived from hES cells and that these cells have the potential for use in cellular and gene therapies. To our knowledge this is the first demonstration that hES cell derivatives can be used for infectious disease research. Due to the extensive ability for hES cells to self-renew, large numbers of differentiated cells can be derived so that infection studies and evaluation tests can be carried out in a more standardized way. Our results showing that both normal and transgenic derivative macrophages support HIV-1 infection points out to their utility for testing anti-HIV constructs transduced into hES-CD34 cells and pave the way for their application in stem cell based HIV gene therapy. So far a number of studies including our own have tested many gene therapeutic constructs in CD34 cells from conventional sources. These constructs include anti-HIV ribozymes, RNA decoys, transdominant proteins, bacterial toxins, anti-sense nucleic acids, and most recently siRNAs [36-50]. In addition, a number of cellular molecules that aid in HIV-1 infection such as cellular receptors and coreceptors CD4, CCR5 and CXCR4 have also been successfully tested in CD34 cell derived macrophages and T cells . Some of these approaches have progressed into clinical evaluations as well . Based on our current results, many of these novel anti-HIV constructs can also be tested in hES-CD34 cells for their potential application. Although there are advantages of using hES cell derived CD34 cells for potential cellular therapies, transplantation of these cells constitutes an allogenic source with immune rejection as a major issue. However, a recent study using human leukocyte reconstituted mice suggested that hESCs and their derivative cell types were less prone to invoking an allogeneic response . Other recent studies demonstrated successful engraftment of primary and secondary recipients with hES cell derived hematopoietic cells in both immunodeficient mice and in vivo fetal sheep models adding further support that any obstacles could be overcome . Moreover, multiple novel strategies to avoid immune-mediated rejection of hES cell-derived cells have been proposed . It is not too far in the future that even autologous hES cells may be derived from specific individuals for deriving CD34 cells which can be used for cell replacement therapy. Phenotypically normal and functionally competent macrophages could be derived from hES-CD34 cells. Since these cells are susceptible to HIV-1 infection, they provide a uniform source of macrophages for viral infection studies. Based on these results, it is also now feasible to transduce hES-CD34 cells with anti-HIV genes such as inhibitory siRNAs and test their antiviral efficacy in down stream differentiated cells such as macrophages which are among the primary cells that need to be protected against HIV-1 infection. Thus, the potential utility of hES derived CD34 hematopoietic cells for HIV-1 gene therapy can be evaluated. The NIH approved human ES H1 cell line was obtained from WiCell (Madison, Wisconsin). hES cell colonies were cultured on mouse embryonic fibroblasts (MEF) (Chemicon, Temecula, CA) in the presence of DMEM-F12 (Invitrogen, Carlsbad, CA) supplemented with 20% KNOCKOUT serum replacement with 1 mM L-glutamine, 1% Nonessential Amino Acids, 0.1 mM β-mercaptoethanol, 0.5% penicillin/streptomycin, and 4 ng/ml human basic fibroblast growth factor. Culture medium was replaced daily with fresh complete DMEM-F12. Mature colonies were subcultured weekly by digesting with collagenase IV as previously described . A VSV-G pseudotyped lentiviral vector (SINF-EF1a-GFP) containing a GFP reporter gene (kindly supplied by R. Hawley, George Washington University) was used for hES cell transductions as previously described (30, 58). Generation of the pseudotyped vector in 293T cells and its concentration by ultracentrifugation were described previously . For vector transduction, the undifferentiated hES cells were prepared into small clumps of 50–100 cells with enzyme digestion as done for routine passaging of cells. The cell clumps were incubated with the vector for 2 hrs in the presence of polybrene 6 ug/ml. A secondary cycle of transduction was done by adding fresh vector and incubating for another 2 hrs. The general vector titers were 1 × 10and the multiplicity of infection was 10. The transduction efficiency was about 50%. The transduced colonies were cultured on MEF like above. Undifferentiated hES cells were cultured on S17 mouse bone marrow stromal cell monolayers to derive cystic bodies containing CD34+ hematopoietic progenitor stem cells. hES cell cultures were treated with collagenase IV(1 mg/ml) for 10 minutes at 37°C and subsequently detached from the plate by gentle scraping of the colonies. The hES cell clusters were then transferred to irradiated (35 Gy) S17 cell layers and cultured with RPMI differentiation medium containing 15% FBS (HyClone, Logan, UT), 2 mM L-glutamine, 0.1 mM β-mercaptoethanol, 1% MEM-nonessential amino acids, and 1% penicillin/streptomycin. Media was changed every 2 to 3 days during 14–17 days of culture on S17 cells . After allowing adequate time for differentiation, hES cystic bodies were harvested and processed into a single cell suspension by collagenase IV treatment followed by digestion with trypsin/EDTA supplemented with 2% chick serum (Invitrogen, Carlsbad, CA) for 20 minutes at 37°C. Cells were washed twice with PBS and filtered through a 70 uM cell strainer to obtain a single cell suspension. To assess the levels of CD34 cells in the bulk cell suspension, cells were labeled with PE conjugated anti-CD34 antibody (BD Biosciences, San Jose, CA) and analyzed by FACS. To purify the CD34 cells, Direct CD34 Progenitor Cell Isolation Kit (Miltenyi Biotech, Auburn, CA) was used following the manufacturer's protocol. Isolated CD34 hematopoietic progenitor stem cells were then analyzed by FACS as mentioned above to determine cell purity. For comparative experiments, human CD34 hematopoietic progenitor cells were also purified from fetal liver tissue as described above. CD34 cells were cultured initially in semisolid media to derive myelomonocytic colonies followed by liquid culture in cytokine supplemented media as described below. Purified CD34+ progenitor cells (~2.5 × 10to 4.0 × 10) were placed directly into Methocult semisolid medium (Stem Cell Technologies, Vancouver, BC), mixed, and cultured in 35 mm plates. Myeloid colonies were allowed to develop for 12–15 days. Upon differentiation and proliferation, myelomonocytic colonies were harvested by the addition of 5 ml DMEM containing 10% FBS, 10 ng/ml each GM-CSF and M-CSF. Cells (~10) were placed in a 35 mm well and allowed to adhere for 48 hours. At two and four days post-harvest, medium was replaced with fresh complete DMEM supplemented with 10 ng/ml GM-CSF and M-CSF. By 4–5 days, cells developed into mature macrophages which were used for subsequent phenotypic and functional characterization. To determine if nontransduced and lentiviral vector transduced hES cell derived macrophages display normal macrophage surface markers, FACS analysis was performed using respective fluorochrome conjugated antibodies. Fetal liver derived CD34+ cells as well as nontransduced and transduced hES cell derived macrophages were evaluated in parallel. Cells were scraped from their wells, washed two times with PBS, and stained with the following antibodies: PE-CD14, PE-HLA-DR, PECY5-CD4, PECY5-CCR5, PECY5-CXCR4 (BD Biosciences, San Jose, CA). A blocking step was first performed by incubating the cells with the respective isotype control for 30 minutes at 4C before staining with the respective cell surface marker antibodies. Isotype control staining was used to determine background levels. FACS analysis was performed on a Beckman-Coulter EPICS XL-MCL flow cytometer with data analysis using EXPO32 ADC software (Coulter Corporation, Miami, FL). A minimum of 8,000 cells were analyzed in each FACS evaluation. Physiological roles of macrophages include phagocytic and immune related functions. To determine if hES cell derived macrophages were functionally normal, a stimulation assay to determine upregulation of the costimulatory molecule B7.1 was performed. Activated macrophages upregulate the expression of B7.1 upon activation with various stimuli. Accordingly, fetal liver CD34, nontransduced hES, and GFP-alone transduced hES cell derived macrophages were stimulated by the addition of LPS (5 ug/ml) to the cell culture medium. Twenty-four hours post-stimulation, cells were stained with an anti-B7.1 antibody labeled with PE-Cy5 (BD Biosciences, San Jose, CA) and analyzed by FACS. To assess the hES cell derived macrophages' phagocytic function, 5 ug/ml of fluorescently labeled E. coli Bioparticles(Invitrogen, Carlsbad, CA) were added directly to the cell culture medium. Four hours later, macrophages were washed six times with PBS and fresh medium with 10 ng/ml GM-CSF and M-CSF was added. Twenty-four hours later, cells were analyzed by FACS for the presence of ingested Bioparticleswhich can be detected in the PE (FL2) channel. Lentiviral vector transduced Magi-CXCR4 cells, a HeLa cell derivative with no phagocytic capacity, were used as non-phagocytic cell controls and similarly exposed to E. coli Bioparticles Human ES cell derived macrophages were also analyzed for their ability to secrete two major cytokines, IL-1 and TNF-α, upon external stimulation. Accordingly, macrophages were stimulated with 5 ug/ml of LPS during culture. On days 1, 2, and 3 post-stimulation, cell culture supernatant samples were collected and analyzed by a QuantikineELISA kit (R&D Systems, Minneapolis, MN). Non-stimulated supernatants were also analyzed for basal levels of cytokine secretion. To determine if hES cell derived macrophages can be infected with HIV-1 and support viral replication, cells were challenged with a macrophage R5-tropic BaL-1 strain of HIV-1. An m.o.i. of 0.01 in the presence of 4 ug/ml polybrene was used. At different days post-infection, culture supernatants were collected and assayed for p24 antigen by ELISA. To quantify viral p24 levels, a Coulter-p24 kit (Beckman Coulter, Fullerton, CA) was used. The author(s) declare that they have no competing interests. JA and SB contributed equally to this work. SB was responsible for deriving CD34 cells from the hESC and culturing macrophages. JA performed the phenotypic, functional and infection assays on the differentiated macrophages. DSK provided hES cell protocols and supplied lentiviral vector transduced cells. RA was responsible for the overall experimental design and implementation of the project. hES cell derived macrophages support productive HIV-1 infection: Macrophages derived from transduced and nontransduced hES CD34 and fetal liver CD34 cells were infected with macrophage R5-tropic HIV-1 BaL-1 strain at an m.o.i. of 0.01. Culture supernatants were collected on different days post infection and assayed for viral p24 antigen by ELISA. Data is representative of triplicate experiments. |
PMC1160574 | Derivation of Multipotent Mesenchymal Precursors from Human Embryonic Stem Cells | Human embryonic stem cells provide access to the earliest stages of human development and may serve as a source of specialized cells for regenerative medicine. Thus, it becomes crucial to develop protocols for the directed differentiation of embryonic stem cells into tissue-restricted precursors. Here, we present culture conditions for the derivation of unlimited numbers of pure mesenchymal precursors from human embryonic stem cells and demonstrate multilineage differentiation into fat, cartilage, bone, and skeletal muscle cells. Our findings will help to elucidate the mechanism of mesoderm specification during embryonic stem cell differentiation and provide a platform to efficiently generate specialized human mesenchymal cell types for future clinical applications.Embryonic stem (ES) cells are pluripotent cells derived from the inner cell mass of the blastocyst that can be maintained in culture for an extended period of time without losing differentiation potential. The successful isolation of human ES cells (hESCs) has raised the hope that these cells may provide a universal tissue source to treat many human diseases. However, directed differentiation of hESCs into specific tissue types poses a formidable challenge. Protocols are currently available for only a few cell types, mostly of neural identity [1–3], and differentiation into many of the cell types derived from the paraxial mesoderm has not been reported, with the exception of a recent study indicating osteoblastic differentiation . Mesenchymal stem cells (MSCs) have been isolated from the adult bone marrow , adipose tissue , and dermis and other connective tissues . Harvesting MSCs from any of these sources requires invasive procedures and the availability of a suitable donor. The number of MSCs that can be obtained from a single donor is limited, and the capacity of these cells for long-term proliferation is rather poor. In contrast, hESCs could provide an unlimited number of specialized cells. In this study, we present techniques for the generation and purification of mesenchymal precursors from hESCs and their directed differentiation in vitro into various mesenchymal derivatives, including skeletal myoblasts. Our isolation method for mesenchymal precursors is the first example, to our knowledge, of efficiently deriving structures of the paraxial mesoderm from ES cells, and further highlights the potential of hESCs for basic biology and regenerative medicine. Undifferentiated hESCs, H1 (WA-01, XY, passages 40–65) and H9 (WA-09, XX, passages 35–45), were cultured on mitotically inactivated mouse embryonic fibroblasts (Specialty Media, Phillipsburg, New Jersey, United States) and maintained under growth conditions and passaging techniques described previously . OP9 cells were maintained in alpha MEM medium containing 20% fetal bovine serum (FBS) and 2 mM L-glutamine. Mesenchymal differentiation was induced by plating 10 × 10 to 25 × 10 cells/cm on a monolayer of OP9 cells in the presence of 20% heat-inactivated FBS in alpha MEM medium. Flow-activated cell sorting (FACS) (CD73-PE; PharMingen, San Diego, California, United States) was performed on a MoFlo (Cytomation, Fort Collins, Colorado, United States). All human ES cell–derived mesenchymal precursor cell (hESMPC) lines in this study are of polyclonal origin. Primary human bone marrow–derived MSCs and primary human foreskin fibroblasts (both from Poietics, Cambrex, East Rutherford, New Jersey, United States) were grown in alpha MEM medium containing 10% FBS and 2 mM L-glutamine. hESMPCs are grown to confluence followed by exposure to 1 mM dexamethasone, 10 μg/ml insulin, and 0.5 mM isobutylxanthine (all from Sigma, St. Louis, Missouri, United States) in alpha MEM medium containing 10% FBS for 2–4 wk. Data were confirmed in hESMPC-H1.1, -H1.2, -H1.3, and -H9.1 (hESMPC-H1.4 was not tested). Differentiation of hESMPCs was induced in pellet culture by exposure to 10 ng/ml TGF-β3 (R & D Systems, Minneapolis, Minnesota, United States) and 200 μM ascorbic acid (Sigma) in alpha MEM medium containing 10% FBS for 3–4 wk. Data were confirmed in hESMPC-H1.1, -H1.3, and -H9.1 (hESMPC-H1.2 and -H1.4 were not tested). hESMPCs were plated at low density (1 × 10 to 2.5 × 10 cells/cm) on tissue-culture-treated dishes in the presence of 10 mM β-glycerol phosphate (Sigma), 0.1 μM dexamethasone, and 200 μM ascorbic acid in alpha MEM medium containing 10% FBS for 3–4 wk. Data were confirmed in hESMPC-H1.1, -H1.3, and -H9.1 (hESMPC-H1.2 and -H1.4 were not tested). Confluent hESMPCs were maintained for 2–3 wk in alpha MEM medium with 20% heat-inactivated FBS. More rapid induction was observed in the presence of medium conditioned for 24 h by differentiated C2C12 cells. Coculture of hESMPCs and C2C12 cells was carried out in alpha MEM with 3% horse serum and 1% FBS . Data were confirmed in hESMPC-H1.3, -H1.4, and -H9.1 (hESMPC-H1.1 and -H1.2 were not tested). Immunocytochemistry for all surface markers was performed on live cells. Monoclonal antibodies VCAM, STRO-1, ICAM-1(CD54), CD105, CD29, and MF20 were from Developmental Studies Hybridoma Bank (University of Iowa, Iowa City, Iowa, United States); CD73, CD44, and ALCAM(CD166) were from BD Biosciences Pharmingen (San Diego, California, United States). All other immunocytochemical analyses were performed after fixation in 4% paraformaldehyde and 0.15% picric acid, followed by permeabilization in 0.3% Triton X100. Polyclonal antibodies used were MyoD (Santa Cruz Biotechnology, Santa Cruz, California, United States) and nestin (gift from R. McKay); monoclonal antibodies were vimentin, alpha smooth muscle actin, fast-switch myosin, pan-cytokeratin (all from Sigma), and human nuclear antigen (Chemicon, Temecula, California, United States). Alkaline phosphatase reaction was performed using a commercially available kit (Kit-86; Sigma) and the mineral was stained with silver nitrate according to the von Kossa method. Fat granules were visualized by Oil Red O staining solution (Sigma). Alcian Blue (Sigma) was used to detect extracellular matrix proteoglycans in chondrogenic cultures. Total RNA was extracted by using the RNeasy kit and DNase I treatment (Qiagen, Valencia, California, United States). Total RNA (2 μg each) was reverse transcribed (SuperScript; Invitrogen, Carlsbad, California, United States). PCR conditions were optimized and linear amplification range was determined for each primer by varying annealing temperature and cycle number. PCR products were identified by size, and identity was confirmed by DNA sequencing. Primer sequences, cycle numbers, and annealing temperatures are provided in Table S1. Total RNA (5 μg) from primary MSCs, from hESMPC-H9.1, hESMPC-H1.2, and three samples of undifferentiated hESCs (H1; passages 42–46), were processed by the Memorial Sloan-Kettering Cancer Center Genomics Core Facility and hybridized on Affymetrix (Santa Clara, California, United States) U133A human oligonucleotide arrays. Data were analyzed using MAS5.0 (Affymetrix) software. Transcripts selectively expressed in each of the mesenchymal cell populations (MSC, hESMPC-H9.1, and hESMPC-H1.2) were defined as those called “increased” by the MAS5.0 algorithm in each of three comparisons with independent samples of undifferentiated hESCs. A Venn diagram was generated to visualize overlap in gene expression. Further statistical analyses were performed as described below. Mesenchymal differentiation of hESCs (lines H1 [WA-01] and H9 [WA-09]) was induced by plating undifferentiated hESCs on a monolayer of murine OP9 stromal cells , in the presence of 20% heat-inactivated FBS in alpha MEM medium. OP9 cells have been previously shown to induce blood cell differentiation from mouse ES cells . After 40 d of coculture, cells were harvested and sorted by FACS for CD73, a surface marker expressed in adult MSCs (Figure 1A). An average of 5% CD73+ cells was obtained from the mixed culture of OP9 and differentiated hESC progeny. CD73+ cells were replated in the absence of stromal feeders on tissue culture plates and expanded in alpha MEM medium with 20% FBS for 7–14 d. We next established the membrane antigen profile of the resulting population of flat spindle-like cells. The H1- and H9-derived CD73+ cells expressed a comprehensive set of markers that are considered to define adult MSCs, including CD105(SH2), STRO-1, VCAM (CD106), CD29(integrin β1), CD44, ICAM -1(CD54), ALCAM(CD166), vimentin, and alpha smooth muscle actin (Figure 1B and 1C). The cells were negative for hematopoietic markers such as CD34, CD45, and CD14. They were also negative for neuroectodermal, epithelial, and muscle cell markers including nestin, pancytokeratin, and desmin (data not shown). The human identity of these presumed mesenchymal cells (termed hESMPC-H1.1, -H1.2, -H1.3, -H1.4, and -H9.1) was confirmed for all experiments by immunocytochemistry for human nuclear antigen to rule out the possibility of contamination with OP9 cells (Figure S1). (A) FACS (MoFlo, Cytomation) for the isolation of CD73+ precursors (right) and isotype control (left). (B) Flow cytometry analysis of the CD73+ hESMPC population for various markers characteristic of MSCs, including CD44, CD73, CD105, CD166, VCAM, ICAM-1, CD29, and STRO-1. (C) Immunocytochemistry of hESMPCs for MSC markers (VCAM, STRO-1, CD73, and CD105). The cells also express vimentin and alpha smooth muscle actin. Scale bar = 50 μm. (D) Venn diagram presenting the overlap among transcripts selectively expressed in hESMPC-H1.2, hESMPC-H9.1, and primary adult human MSCs. To further characterize hESMPCs, we performed genome-wide expression analysis using oligonucleotide arrays (Affymetrix U133A). The expression profiles of hESMPC-H1.2 and hESMPC-H9.1 were compared with that of human primary adult MSCs. Housekeeping genes for each of the mesenchymal cell populations were eliminated by subtracting those transcripts also expressed in at least one of three independent samples of undifferentiated hESCs. Based on this analysis, 1,280 transcripts were selectively expressed in hESMPC-H1.2, 932 transcripts in hESMPC-H9.1, and 1,218 transcripts in primary adult MSCs. A remarkable overlap of 579 transcripts shared among the three mesenchymal populations was observed (Figure 1D). Using the genes that were selected in the initial filter, we performed a statistical analysis on the expression levels to determine whether the genes were expressed significantly differently in the two cell types. We used a Bayesian extension to the standard t-test to assess this difference. Of the 579 genes, 412 of them were significantly different, at a false discovery rate cutoff of 0.05. The relative fold changes were also extremely large in many of the cases. We also looked at the variance of the expression levels within the cell types. For the MSCs, 94% had a coefficient of variation less than 20% for the expression (log transformed); for the ES-derived cells, 72% had a coefficient of variation less than 20%. Numerous known MSC markers were included in the list of 412 genes, such as the mesenchymal stem cell protein DSC54 (13.9-fold increase, p < 0.001), neuropilin 1 (30.4-fold increase, p < 0.001), hepatocyte growth factor (48.1-fold increase, p < 0.001), forkhead box D1 (14.8-fold increase, p < 0.001), and notch homolog 2 (2.9-fold increase, p < 0.001) . Table S2 lists the p-values from the test, the mean and standard deviation of the expression levels, and the relative fold change of all 412 genes between the two types. Known markers of MSCs, such as mesenchymal stem cell protein DSC54, were all included within the 579 shared transcripts. These findings support the immunocytochemical data and suggest that hESMPCs and primary MSCs are highly related. MSCs are characterized functionally by their ability to differentiate into mesenchymal tissues, such as fat, cartilage, and bone. Therefore, we tested whether hESMPCs have the same potential (Figure 2). (A) Adipocytic differentiation in the presence of dexamethasone, insulin, and isobutylxanthine. Adipocytic characterization by Oil Red O staining and RT-PCR analysis for PPARγ. (B) Chondrocytic differentiation in the presence of TGF-β3 and ascorbic acid. Chondrocytic characterization by Alcian Blue staining and RT-PCR for aggrecan and collagen II. (C) Osteogenic differentiation in the presence of β-glycerolphosphate, dexamethasone, and ascorbic acid. Osteocytic characterization by von Kossa staining and RT-PCR for bone-specific alkaline phosphatase (ALP) and bone sialoprotein (BSP). (D) Phase-contrast image of hESMPCs and RT-PCR for the ES cell markers Nanog and Oct-4 in hESMPC-H1.1 and -H9.1 compared with undifferentiated H1 hESCs. Scale bar = 50 μm for all panels. Adipocytic differentiation of hESMPCs was induced under conditions described previously for primary adult MSCs . Appearance of cells harboring fat granules was observed after 10–14 d in culture. After 3 wk of induction, more than 70% of the cells displayed Oil Red O+ fat granules, and PPARγ, a marker of adipocytic differentiation, was detected by RT-PCR. (Figure 2A). Chondrocytic differentiation was achieved using the pellet culture system . After 28 d in culture, more than 50% of all cells exhibited robust staining for Alcian Blue, a marker specific for extracellular matrix proteoglycans. Chondrocytic differentiation was confirmed by the gene expression of collagen II and aggrecan, two components of extracellular matrix selectively expressed by chondrocytes, using RT-PCR (Figure 2B). Osteogenic differentiation was induced in the presence of β-glycerolphosphate . Osteogenesis was demonstrated by specific staining for calcium deposition in the matrix (von Kossa, Figure 2C; or Alizarin Red, Figure S2A) and increased expression of bone-specific alkaline phosphatase and bone sialoprotein at day 28 of treatment (Figures 2C and S2B). At day 28, Alizarin Red staining was detected in approximately 70% of all cells. Throughout these studies, human adult MSCs and foreskin fibroblasts were used as positive and negative controls, respectively. In addition to adipocytic, chondrocytic, and osteogenic differentiation, reports suggested that adult MSCs can form skeletal muscle . Although generation of skeletal muscle cells from adult MSCs remains controversial, we tested whether hESMPCs exhibit this potential. Under the conditions previously described , hESMPC-H1.1 and -H9.1 did not yield significant numbers of MyoD+ cells after 15–20 d in culture. However, when confluent cells were maintained in culture in the presence or absence of 5-AzaC without passage for more than 21 d, expression of specific skeletal muscle markers such as MyoD and fast-switch myosin was observed (Figure 3A). More rapid myogenic differentiation was obtained in the presence of 24-h-conditioned medium from the murine myoblastic cell line C2C12 previously induced to form myotubes . Direct coculture of hESMPCs with C2C12 cells led to the formation of hESMPC-derived myotubes, as visualized by expression of human nuclear antigen (Figure 3B), similar to those formed by host C2C12 cells. After 1 wk of coculture, myotubes composed of human nuclei accounted for more than 10% of the total number of human cells present, and each human myotube was composed of up to ten human nuclei. Human cell contribution to myotubes in coculture was confirmed by expression of human muscle-specific transcripts such as MyoD, myosin heavy chain IIa, and myogenin (data not shown). These data demonstrate that hESMPCs can give rise to mesenchymal derivatives typically obtained from primary adult MSCs, as well as to cells expressing markers of skeletal muscle. (A) Immunocytochemistry for MyoD (red) and fast-switch myosin (green). RT-PCR for MyoD in human skeletal muscle as a positive control (hSM), and in hESMPC-H9.1 cells differentiated for 10 d in the presence of C2C12-conditioned medium (hESMPC). (B) Myotube formation induced at high cell densities in the presence of C2C12 cells. Myotube characterization by immunocytochemistry for MF20 against sarcomeric myosin (green) and human nuclear antigen (hNA, red). Left panel: Control undifferentiated hESCs (H9) do not fuse with C2C12. Right panel: Under identical culture conditions, hESMPCs (line 9.1) efficiently fuse with C2C12 cells, forming myotubes containing human nuclei. RT-PCR for human specific muscle transcripts myosin heavy chain IIa (MYHC-2) and MyoD in C2C12 cells, in human skeletal muscle as positive control (huSM), and in hESMPC-H9.1 cells cocultured with C2C12 cells. One concern for the clinical application of hESC-derived progeny in regenerative medicine is the risk of teratoma formation due to the presence of residual undifferentiated ES cells among the differentiated progeny. We did not detect markers of undifferentiated hESCs, such as Nanog or Oct-4 , in any of the hESMPCs by RT-PCR (see Figure 2D) and immunocytochemistry (data not shown), suggesting the lack of any undifferentiated ES cells in hESMPC cultures. However, future in vivo studies are required to rule out the potential of these cells for teratoma formation. Previous studies have demonstrated the derivation of neural cells [1–3], hematopoietic and endothelial lineages , and cardiomyocytes from hESCs. This study presents the induction of paraxial mesoderm with the generation of multipotent mesenchymal precursors. We calculate that under these conditions a single undifferentiated hESC yields an average of one CD73+ cell at day 40 of differentiation, suggesting a balance between cell proliferation and cell selection. There were no obvious differences in marker and gene-expression profile or in differentiation behavior among the five hESMPC lines generated. However, some of the lines (e.g., hESMPC9.1) exhibited a tendency of spontaneous osteogenic differentiation after long-term propagation. Directed differentiation of hESCs into somatic stem-cell-like precursors represents a substantial advancement in harnessing the developmental potential of hESCs. The high purity, unlimited availability, and multipotentiality of hESMPCs will provide the basis for future therapeutic efforts using these cells in preclinical animal models of disease. Such in vivo studies will also be required to properly assess the safety profile of these cells. Furthermore, our system also offers a novel platform to study basic mechanisms of mesodermal induction and differentiation during early human development. The Gene Expression Omnibus (GEO) (http://www.ncbi.nlm.nih.gov/geo) accession number for all raw microarray data used in this study is GSE2248. The Unigene (http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=unigene) accession numbers for the gene products discussed in this paper are aggrecan (Hs.2159 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=2159; bone sialoprotein (Hs.518726 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=518726; bone-specific alkaline phosphatase (Hs.75431 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=75431; collagen II (Hs.408182 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=408182; forkhead box D1 (Hs.519385 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=519385; hepatocyte growth factor (Hs.396530 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=396530; mesenchymal stem cell protein (DSC54, Hs.157461 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=157461; MyoD (Hs.520119 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=520119; myogenin (Hs.2830 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=2830; myosin heavy chain IIa (Hs.513941 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=513941; Nanog (Hs.329296 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=329296]) ; neuropilin 1 (Hs.131704 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=131704; notch homolog 2 (Hs.549056 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=549056; Oct-4 (Hs.504658 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=504658; and PPARγ (Hs.162646 [http://www.ncbi.nlm.nih.gov/UniGene/clust.cgi?ORG=Hs&CID=162646]). The discovery and isolation of human embryonic stem cells (cells that are capable of renewing themselves and turning into the many different cell types that make up the human body) has the potential to revolutionize the treatment of many diseases that require the replacement of abnormal or missing cells. In particular, it would be very valuable to be able to replace tissues that are derived from one particular tissue type—mesenchyme—which bone, cartilage, fat and muscle develop from. However, before such treatments can happen, it will be necessary to work out exactly how embryonic cells become other cells, and whether it is possible to make these changes happen in the laboratory. They took two lines of completely undifferentiated human embryonic stem cells and by culturing them in the presence of mouse cells stimulated them to turn into mesenchymal cells. They then treated these cells with compounds to make them change into specialized bone, cartilage, fat, and muscle cells. They were able to confirm that these cells were all human (important because the early part of the experiment is done in the presence of mouse cells) and also that there was no evidence that the cells became cancerous. It is theoretically possible to produce lines of bone, cartilage, fat, and muscle cells from human embryonic stem cells. However, the process will need more refinement before the cell lines could be used for treatment; ideally, for example, all the culturing would be done without any mouse cells. The United States National Institutes of Health has a group of Web pages on stem cells: http://stemcells.nih.gov/info/faqs.asp The International Society for Stem Cell Research has a list of frequently asked questions about stem cells: http://www.isscr.org/science/faq.htm We thank R. McKay for nestin antibody; P. Song and the Sloan-Kettering Genomics and Flow Cytometry Core Facilities for technical assistance; and R. Stan, V. Tabar, M. Tomishima, Y. Elkabetz, and S. Desbordes for critical review of the manuscript. This work was supported in part by the Kinetics Foundation. The funder had no role in the study design, data analysis, decision to publish, or manuscript preparation and content. embryonic stem flow-activated cell sorting fetal bovine serum human embryonic stem cell human embryonic stem cell–derived mesenchymal precursor cell mesenchymal stem cell Citation: Barberi T, Willis LM, Socci ND, Studer L (2005) Derivation of multipotent mesenchymal precursors from human embryonic stem cells. PLoS Med 2(6): e161. |
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