Citation Davids B-O, Balasubramaniam M, Sapp N, Prakash P, Ingram S, Li M, Craigie R, Hollis T, Pandhare J, Dash C. 2021. Human three prime repair exonuclease 1 promotes HIV-1 integration by preferentially degrading unprocessed viral DNA. J Virol 95:e00555-21. https://doi.org/10.1128/JVI.00555-21.
Received 2021 Apr 1; Accepted 2021 May 25. Copyright © 2021 American Society for Microbiology.Three prime repair exonuclease 1 (TREX1) is the most abundant 3′→5′ exonuclease in mammalian cells. It has been suggested that TREX1 degrades HIV-1 DNA to enable the virus to evade the innate immune system. However, the exact role of TREX1 during early steps of HIV-1 infection is not clearly understood. In this study, we report that HIV-1 infection is associated with upregulation, perinuclear accumulation, and nuclear localization of TREX1. However, TREX1 overexpression did not affect reverse transcription or nuclear entry of the virus. Surprisingly, HIV-1 DNA integration was increased in TREX1-overexpressing cells, suggesting a role of the exonuclease in the post-nuclear entry step of infection. Accordingly, preintegration complexes (PICs) extracted from TREX1-overexpressing cells retained higher levels of DNA integration activity. TREX1 depletion resulted in reduced levels of proviral integration, and PICs formed in TREX1-depleted cells retained lower DNA integration activity. Addition of purified TREX1 to PICs also enhanced DNA integration activity, suggesting that TREX1 promotes HIV-1 integration by stimulating PIC activity. To understand the mechanism, we measured TREX1 exonuclease activity on substrates containing viral DNA ends. These studies revealed that TREX1 preferentially degrades the unprocessed viral DNA, but the integration-competent 3′-processed viral DNA remains resistant to degradation. Finally, we observed that TREX1 addition stimulates the activity of HIV-1 intasomes assembled with the unprocessed viral DNA but not that of intasomes containing the 3′-processed viral DNA. These biochemical analyses provide a mechanism by which TREX1 directly promotes HIV-1 integration. Collectively, our study demonstrates that HIV-1 infection upregulates TREX1 to facilitate viral DNA integration.
IMPORTANCE Productive HIV-1 infection is dependent on a number of cellular factors. Therefore, a clear understanding of how the virus exploits the cellular machinery will identify new targets for inhibiting HIV-1 infection. The three prime repair exonuclease 1 (TREX1) is the most active cellular exonuclease in mammalian cells. It has been reported that TREX1 prevents accumulation of HIV-1 DNA and enables the virus to evade the host innate immune response. Here, we show that HIV-1 infection results in the upregulation, perinuclear accumulation, and nuclear localization of TREX1. We also provide evidence that TREX1 promotes HIV-1 integration by preferentially degrading viral DNAs that are incompatible with chromosomal insertion. These observations identify a novel role of TREX1 in a post-nuclear entry step of HIV-1 infection.
KEYWORDS: Three prime repair exonuclease 1, TREX1, human immunodeficiency virus, HIV, integration, preintegration complex, PIC, intasome, exonuclease, reverse transcription
Three prime repair exonuclease 1 (TREX1) is the most abundant 3′→5′ exonuclease in mammalian cells (1,–3). The primary function of TREX1 is to degrade cellular DNA originating from aberrant DNA replication, recombination, and repair (4, 5). In addition, TREX1 degrades DNA intermediates of endogenous retroviruses and retroelements, the remnants of ancient retroviral infections that are vertically transmitted in the population (4, 5). Removal of these endogenous DNA products is critical to abrogate activation of the cellular DNA sensing pathway and prevent an autoimmune response (4, 5). Accordingly, genetic mutations in human TREX1 have been implicated in autoimmune and auto inflammatory disorders such as Aicardi-Goutières syndrome (AGS), familial chilblain lupus (FCL), systemic lupus erythematosus (SLE), and retinal vasculopathy with cerebral leukodystrophy (RVCL) (6, 7). Interestingly, in recent years, TREX1 has garnered a great deal of attention for its role during early steps of human immunodeficiency virus type 1 (HIV-1) infection.
HIV-1 is the causative agent for AIDS (8). The virus has infected >73 million people and has been responsible for more than 37 million deaths worldwide (UNAIDS, 2018). HIV-1 primarily infects CD4 + T lymphocytes and enters the cell by engaging with the CD4 receptor and the CCR5 or CXCR4 coreceptor (9). Binding of the viral envelope glycoprotein to these receptors orchestrates fusion of the viral and cellular plasma membranes and release of the viral capsid into the cytoplasm (10). The incoming viral capsid contains two copies of the single-stranded viral RNA genome and a number of viral/cellular proteins that are necessary for productive infection (11,–16). First, the reverse transcription complex (RTC) converts the single-stranded viral RNA genome into a double-stranded DNA copy (17). Then, the preintegration complex (PIC) carries out integration of the viral DNA into the host chromosome (18,–21).
It is well established that the early steps of HIV-1 infection involve a number of viral nucleic acid intermediates. For instance, the reverse transcription process includes single-stranded RNA (ssRNA), double-stranded RNA (dsRNA), single-stranded DNA (ssDNA), double-stranded DNA (dsDNA), and DNA/RNA hybrids (17). Additionally, the reverse transcribed viral dsDNA undergoes 3′ processing to produce a 5′ overhang and recessed 3′ DNA ends (19). Despite the presence of these viral nucleic acid substrates in the infected cell, chronic HIV-1 infection does not trigger an interferon stimulatory DNA (ISD) response (22). Although the underlying mechanism for a lack of an ISD response is not fully understood, there is evidence that the viral capsid could shield the RNA genome and the PIC may protect the viral DNA from the cellular sensors (23,–26). Interestingly, it has also been suggested that TREX1 may play a key role in the cellular ISD response to HIV-1 infection.
TREX1’s role during HIV-1 infection was first recognized in a study of the human SET complex (27). The SET complex is an endoplasmic reticulum (ER)-associated multiprotein DNA repair complex, containing TREX1 and two other nucleases, that localizes to the nucleus in response to oxidative stress (28, 29). In that study, knockdown of TREX1 inhibited HIV-1 infection at a step after the reverse transcription of the viral genome (27). Importantly, disruption of the interaction between the SET complex and HIV-1 PIC was shown to increase autointegration of the viral DNA concomitant with decreased proviral integration, even though a direct role of TREX1 was not demonstrated (27). A later study demonstrated that TREX1 plays a key role in suppressing interferon response during HIV-1 infection (30). This study reported that depletion of TREX1 in mouse embryonic fibroblasts (MEFs) resulted in the accumulation of cytosolic HIV-1 DNA and induction of an ISD response. Interestingly, the ISD response in TREX1-depleted cells was abrogated by a reverse transcriptase (RT) inhibitor (30). Therefore, it was concluded that TREX1 prevents accumulation of reverse transcribed HIV-1 DNA to avert the interferon response (30). Subsequently, TREX1’s role in the degradation of cytosolic viral DNA was reported in other retroviruses such as murine leukemia virus (MLV) and simian immunodeficiency virus (SIV) (23). A potential role of TREX1 during HIV-1 transmission was also described in human cervicovaginal explants and humanized mice (31). Furthermore, a study showed that TREX1 regulates the HIV-1-associated ISD response in THP-1 monocytic cells (32). Interestingly, a recent report found that HIV-1 infection of T cells failed to mount an RT inhibitor-sensitive immune response despite an intact cyclic GMP-AMP synthase (cGAS) sensing pathway (33). Collectively, these studies suggest that TREX1 may play a key role in the ISD response to HIV-1 infection in a cell-type-dependent manner. However, the exact role of TREX1 during early steps of HIV-1 infection is not fully understood.
In this study, we observed that TREX1 expression is elevated in HIV-1-infected cells. Since TREX1 has been implicated in HIV-1 infection, we probed the exact role of this cellular exonuclease during early steps of infection. We found that TREX1 enhances HIV-1 integration without a measurable effect on reverse transcription and nuclear entry of the virus. Extraction of HIV-1 PICs from infected cells and measurement of DNA integration activity provided biochemical evidence of a direct role of TREX1 in HIV-1 integration. Exonuclease activity assays showed preferential degradation of the unprocessed viral DNA relative to that of the 3′-processed DNA by TREX1. Since 3′-processed viral DNA is critical for HIV-1 integration, these results suggest a novel mechanism by which TREX1 promotes integration. Collectively, our results show that upregulation of TREX1 in HIV-1-infected cells is linked to increased viral DNA integration and uncover a role of TREX1 at a post-nuclear early step of HIV-1 infection.
It has been reported that TREX1 prevents accumulation of cytosolic viral DNA and enables HIV-1 to evade the ISD response (30,–32). However, the details of molecular and biochemical interactions between the virus and TREX1 during early steps of infection are not fully understood. To better understand TREX1’s role during HIV-1 infection, we first measured TREX1 protein levels in human T cell lines Sup-T1 and Jurkat, since these cells are highly permissive to HIV-1 infection and are extensively used to study viral replication (34). These cells were inoculated with different amounts of HIV-1 particles, and protein expression was measured in the cellular lysates 48 h postinfection (hpi) by Western blotting ( Fig. 1 ). Detection of HIV-1 capsid protein (p24) in infected Sup-T1 ( Fig. 1A ) and Jurkat cells ( Fig. 1C ) confirmed productive infection. As expected, higher levels of p24 protein were detected in cells inoculated with an increased amount of virus. Western blot analysis also showed that TREX1 is endogenously expressed in both Sup-T1 and Jurkat cells ( Fig. 1A and andC). C ). Interestingly, the levels of TREX1 in these cells were significantly elevated after HIV-1 infection compared to that in the uninfected control cells ( Fig. 1A to toD). D ). Accordingly, the expression of TREX1 was further increased in cells inoculated with larger amount of virus ( Fig. 1A to toD D ).
HIV-1 infection upregulates TREX1 expression. (A to H) Effects of HIV-1 inoculum on TREX1 expression in T cell lines. Sup-T1 (A) and Jurkat (C) cells were spinoculated with two different concentrations of HIV-1 particles. Productive infection was assessed by probing for HIV-1 p24 protein in the cellular lysates by Western blotting. Representative Western blots showing endogenous TREX1 protein (molecular weight [MW], 33 kDa) expression in uninfected and infected cells, with β-actin (MW, 42 kDa) serving as a protein loading control. Densitometric analysis of TREX1 protein expression in uninfected and infected Sup-T1 (B) and Jurkat (D) cells. (E to H) Effects of HIV-1 infection on TREX1 as a function of time. Sup-T1 and Jurkat cells were spinoculated with 150 ng p24 equivalent (E and F) or 300 ng p24 equivalent (G and H) of HIV-1 particles, and the cells were collected at 6, 12, 24, and 48 hpi. TREX1 expression in these cells was assessed by Western blotting and quantified by densitometry. The TREX1 level in each time point sample was normalized to respective β-actin level, and the fold change in TREX1 expression over time was plotted relative to that at 0 hpi. (I and J) Effects of HIV-1 infection on TREX1 expression in PBMCs. PHA-activated PBMCs (n = 3) were spinoculated with HIV-1 particles, and the cellular lysates were probed by Western blotting for HIV-1 p24 (MW, 24 kDa), TREX1 (MW, 33 kDa), and β-actin (MW, 42 kDa) levels. (I) Representative Western blot showing TREX1 levels in uninfected and infected cells. (J) Densitometric analysis of TREX1 expression in uninfected and infected PBMCs from three donors. Data shown in panels B, D, E to H, and J are mean values from three independent experiments, with error bars representing the standard errors of the mean (SEMs). *, P < 0.05 for the comparison of TREX1 levels between uninfected and infected cells in panels B, D, and J and between 0 hpi and later time points in panels E to H.
To further examine the effects of HIV-1 infection on TREX1 expression, we probed TREX1 levels in infected cells in a time-dependent manner. Sup-T1 and Jurkat cells were inoculated with two different amounts of HIV-1 particles (150-ng and 300-ng equivalent of p24), and cells were collected at 6, 12, 24, and 48 hpi. Western blot analysis illustrated that TREX1 expression was upregulated in infected cells even at the earliest point of 6 hpi ( Fig. 1E to toH). H ). TREX1 levels were further increased in these cells with increasing time of infection. The upregulation of TREX1 expression was maximum at 24 hpi followed by a slight decrease at 48 hpi ( Fig. 1E to toH). H ). The time-dependent upregulation of TREX1 was consistent in cells inoculated with 150 ng ( Fig. 1E and andF) F ) and 300 ng of HIV-1 particles ( Fig. 1G to toH). H ). Taken together, these results show that TREX1 expression is significantly upregulated early during HIV-1 infection and remained elevated up to 48 hpi.
Finally, we measured TREX1 levels in human peripheral blood mononuclear cells (PBMCs), which are primary cells constituting CD4 + lymphocytes that serve as the in vivo target cells of HIV-1 (35, 36). PBMCs were isolated from whole-blood samples of three healthy donors, activated by phytohemagglutinin (PHA), and cultured in the presence of interleukin 2 (IL-2). Then, the cells were inoculated with different amounts of infectious HIV-1 particles and were harvested at 48 hpi. Cellular lysates from both uninfected and infected PBMCs were analyzed by immunoblot analysis. Presence of HIV-1 p24 protein indicated productive infection of the PBMCs ( Fig. 1I ). Similar to the data obtained with T cell lines, TREX1 was endogenously expressed in these primary cells. Furthermore, we also observed that the TREX1 level was significantly upregulated in the PBMCs upon HIV-1 infection compared to that in the uninfected control cells ( Fig. 1I and andJ). J ). Collectively, these results corroborate the observations in T cell lines ( Fig. 1A to toH) H ) and establish that HIV-1 infection induces TREX1 protein levels—an observation that has not been previously reported.
TREX1 was first purified as an exonuclease from the nuclei of mammalian cells (2, 37). However, TREX1 is distributed in the cytoplasm, specifically, it is localized with the ER and in the perinuclear regions (5, 38). There is also strong evidence that TREX1 can be mobilized to the nucleus in response to DNA damage and repair (5, 28, 29). Therefore, TREX1 is distributed both in the cytoplasm and nuclei of mammalian cells to carry out key functions. It is important to note that the early steps of HIV-1 infection are also carried out both in the cytoplasm and the nucleus of the target cell. Therefore, we probed the distribution of TREX1 using both confocal microscopy and immunoblotting in uninfected and infected cells to better understand the effects of HIV-1 on subcellular localization of TREX1.
First, we used HEK293T cells, since these cells do not endogenously express TREX1 protein. HEK293T cells were transfected with a green fluorescent protein GFP-Trex1 fusion construct, and expression of GFP-TREX1 was monitored by green fluorescence using confocal microscopy ( Fig. 2A ). As expected, we observed that the GFP-TREX1 protein was associated with the ER and predominantly localized near the perinuclear regions ( Fig. 2A and andB). B ). Specifically, colocalization was assessed using Pearson’s correlation coefficient for green (GFP-TREX1) and red (calreticulin or lamin). These analyses illustrated significant colocalization between GFP-TREX1 and calreticulin, with a Pearson’s coefficient value of ∼0.53 ( Fig. 2C ), a value much greater than 0.30—the threshold for colocalization. On the other hand, expression of the GFP protein alone without the TREX1 fusion as a control showed a Pearson’s coefficient
HIV-1 infection is associated with perinuclear accumulation and nuclear localization of TREX1. (A and B) HEK293T cells were transfected with a Gfp-Trex1 fusion construct and were subjected to confocal microscopy. GFP-TREX1 was detected by green fluorescence, the ER-associated calreticulin or nuclear envelope-associated lamin was detected by red fluorescence, and DAPI (blue) was probed as a nuclear marker. (C and D) To quantify GFP, red fluorescent protein (RFP), and blue intensity, 10 ROIs were calculated for each condition and the mean intensity value was exported and analyzed by Pearson’s correlation coefficient. Pearson’s correlation coefficient was used to determine the extent of colocalization of TREX1 with calreticulin (n = 10) or lamin (n = 10). (E) TZM-bl cells were transfected with a Gfp-Trex1 fusion construct and were subsequently inoculated with HIV-1 particles. At 24 hpi, both uninfected and infected cells were subjected to confocal microscopy. (F) Colocalization of GFP-TREX1 with HIV-1 p24 was calculated by Pearson’s coefficient. (G) Nuclear intensity of GFP-TREX1 in uninfected and infected cells. To establish statistical significance, a t test was performed for relative uninfected versus infected cells (n = 10). (H) Immunoblot analysis of cytoplasmic and nuclear distributions of TREX1 in uninfected and infected cells. Total cellular lysates, cytoplasmic fractions, and nuclear fractions of uninfected and infected TZM-bl cells were probed for GAPDH as a cytoplasmic protein, lamin as a nuclear protein, and TREX1. HIV-1 p24 protein was probed for productive infection. Representative immunoblot from three independent experiments is shown.
Next, we probed the distribution and localization of TREX1 in cells inoculated with HIV-1 particles. For this experiment, we used TZM-bl cells that express the CD4 receptor and CXCR4/CCR5 coreceptors utilized by HIV-1 for cellular entry. TZM-bl cells were transfected with the Gfp-Trex1 construct, and 24 h posttransfection, these cells were inoculated with VSV-G pseudotyped HIV-1 particles; presence of HIV-1 p24 protein was used as a marker for productive infection. Localization of GFP-TREX1 in infected cells and uninfected control cells was probed by confocal microscopy. Similar to the results in Fig. 2A to toD D with HEK293T cells, GFP-TREX1 was localized with the ER network near the perinuclear regions of uninfected TZM-bl cells, as determined by Pearson’s correlation coefficient (data not shown). Interestingly, in infected cells, GFP-TREX1 was also distributed near the perinuclear regions ( Fig. 2E ). Additionally, in these cells, a significant colocalization between GFP-TREX1 and HIV-1 p24 protein was observed, with a significant Pearson’s coefficient ( Fig. 2F ). Most importantly, we also observed that the GFP-TREX1 protein was relocalized to the nuclei of the infected cells. For instance, quantification of the GFP intensity in the nucleus revealed that the nuclear intensity of GFP-TREX1 was significantly increased in infected cells compared to that in the uninfected cells ( Fig. 2G ). Taken together, these studies suggest that TREX1 is associated with the ER and distributed in the perinuclear regions. However, in HIV-1-infected cells, TREX1 is also localized to the nucleus.
Finally, we probed the distribution of endogenous TREX1 protein in uninfected and infected TZM-bl cells by immunoblot analysis ( Fig. 2H ). We prepared total cell extracts, cytoplasmic fractions, and nuclear fractions of cells 24 to 48 hpi. Detection of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as a cytoplasmic marker and lamin as a nuclear marker was used to clarify the subcellular fractionation. Productive infection was analyzed by the detection of HIV-1 p24 protein. Our cytoplasmic and nuclear fractions were relatively devoid of cross-contaminating proteins ( Fig. 2H ). Interestingly, the viral p24 protein was detected in both the cytoplasmic and nuclear fractions of the infected cells. These results are consistent with the emerging role of HIV-1 capsid protein both in pre-nuclear entry and post-nuclear entry steps of infection. As expected, endogenous TREX1 protein was predominantly detected in the cytoplasmic fraction of uninfected cells. Most importantly, a small but detectable amount of TREX1 protein was present in the nuclei of HIV-1-infected cells, even though the majority of the protein was retained in the cytoplasmic fraction. These biochemical results are consistent with the distribution and localization of GFP-TREX1 observed with our imaging studies ( Fig. 2A to toG). G ). Collectively, imaging and biochemical results provide strong evidence that TREX1 is predominantly distributed in the perinuclear region of uninfected cells, but a portion of the protein is mobilized to the nucleus upon HIV-1 infection.
The effect of higher levels of TREX1 on HIV-1 infection in human T cells has not been studied. Nevertheless, there is evidence that depletion of TREX1 results in the accumulation of reverse transcribed HIV-1 DNA in MEFs and THP-1 monocytic cells (30, 32). Interestingly, a recent study in murine T cells showed that knockout (KO) of TREX1 has minimal effect on HIV-1 DNA levels during infection (33). Conversely, higher levels of TREX1 in THP-1 cells marginally decreased HIV-1 DNA early during infection (32). However, at 24 hpi, higher levels of TREX1 failed to reduce the levels of HIV-1 DNA (32). These studies suggest a cell-type-dependent effect of TREX1 on HIV-1 reverse transcription. Therefore, to understand how higher levels of TREX1 affect early steps of HIV-1 infection, we carried out overexpression studies in T cell models. First, we generated Sup-T1 cells that stably overexpress GFP-TREX1 (TREX1-OE). The TREX1-OE cells were cultured in the presence of the antibiotic G418 to select GFP-positive cells. Then, the top 10% of cells with the highest levels of GFP expression were collected by fluorescence-activated cell sorting (FACS) ( Fig. 3A ). These cells were then cultured, and TREX1 expression in the cellular lysates was verified by Western blotting ( Fig. 3B ). As expected, the TREX1-OE cells expressed robust levels of TREX1 in the form of the GFP fusion protein, whereas no such protein was detected in the parental cells ( Fig. 3B ). Subsequently, TREX1-OE and parental cells without the GFP-TREX1 protein were inoculated with HIV-1 particles. DNA from these cells were subjected to quantitative PCR (qPCR) to quantify late reverse transcription products. Copy numbers of viral DNA were calculated from a standard curve generated using a molecular clone of HIV-1. Resulting data from these qPCR analyses revealed that the amount of reverse transcription products in TREX1-OE cells was not significantly changed compared to that in the parental cells ( Fig. 3C ). For example, the average amount of viral DNA in parental cells was approximately 5.2 × 10 4 copies, which was slightly increased to 5.5 × 10 4 copies in the TREX1-OE cells. The qPCR assay is specific for measuring reverse transcription, since cells treated with the reverse transcriptase inhibitor efavirenz (EFV) showed significantly fewer copies of viral DNA ( Fig. 3C ).
TREX1 overexpression has a minimal effect on HIV-1 reverse transcription. (A) Generation of TREX1-OE Sup-T1 cells. TREX1-OE Sup-T1 cells were generated by electroporation with Gfp-Trex1 fusion plasmid construct. The GFP-positive cells were detected by FACS, and the top 10% cell population exhibiting the highest GFP fluorescence intensity was sorted by FACS. These cells were then expanded by further culturing. (B) GFP-TREX1 expression in Sup-T1 cells. GFP-TREX1 expression level in the TREX1-OE Sup-T1 cells was assessed by Western blot analysis of the cell lysate. The parental Sup-T1 cell lysate was analyzed as a control. Representative immunoblots showing GFP-TREX1 fusion protein (MW, 57 kDa) and β-actin (loading control with MW of 42 kDa). (C) HIV-1 reverse transcription in TREX-OE Sup-T1 cells. Parental and TREX1-OE Sup-T1 cells were inoculated with HIV-1 (150 ng p24 equivalent). As controls, parental Sup-T1 cells were mock inoculated or inoculated with HIV-1 in the presence of efavirenz. Total DNA isolated from these cells was analyzed by qPCR using primers specific for amplification of late RT products (viral DNA). Levels of viral DNA were quantified by calculating copies of viral DNA from a standard curve generated in parallel using 10-fold serial dilutions of known copy numbers (10 0 to 10 8 ) of an HIV-1 molecular clone. (D and E) Detection of GFP-TREX1 in HEK293T cells. HEK293T cells were transfected with the control Gfp plasmid or the Gfp-Trex1 plasmid. GFP-TREX1 expression in the transfected cells was assessed by fluorescence microscopy (D) and Western blot analysis (E). (F) HIV-1 reverse transcription in HEK293T cells expressing GFP-TREX1. HEK293T cells expressing GFP or GFP-TREX1 were inoculated with VSV-G pseudotyped HIV-1. Reverse transcription products in these cells were measured as described in Fig. 3C . (G and H) HIV-1 reverse transcription in TREX1-OE Jurkat cells. (G) TREX1-OE Jurkat cells were generated by electroporation with Gfp-Trex1 fusion plasmid construct. GFP-TREX1 expression level in these cells was assessed by Western blot analysis; representative immunoblot showing GFP-TREX1 fusion protein (MW, 57 kDa) and β-actin (MW, 42 kDa). (H) Parental and TREX1-OE Jurkat cells were inoculated with HIV-1 (150 ng p24 equivalent). Total DNA isolated from these cells was analyzed by qPCR for the copies of late RT products. Data shown are representative of at least three independent experiments, with error bars representing the SEMs.
A lack of reduction in viral DNA levels in TREX1-OE Sup-T1 cells is surprising, since TREX1 depletion is associated with accumulation of HIV-1 DNA (30). Thus, we probed whether the GFP protein contributes to this observation given that TREX1-OE cells express TREX1 as a GFP fusion protein. We used HEK293T cells that lack endogenous levels of TREX1 and transfected the pGfp-Trex1 fusion construct or pGfp control plasmid into these cells ( Fig. 3D and andE). E ). Expression of GFP-TREX1 fusion protein and GFP protein was confirmed by GFP fluorescence ( Fig. 3D ) and Western blot analysis ( Fig. 3E ). Then, the GFP-TREX1-expressing and GFP-expressing HEK293T cells were inoculated with VSV-G pseudotyped HIV-1 particles. DNA isolated from these cells was subjected to qPCR analysis to quantify reverse transcription products. These analyses revealed that the levels of HIV-1 reverse transcription products remained comparable in infected cells expressing either GFP protein or GFP-TREX1 fusion protein ( Fig. 3F ). These results suggested that presence of GFP in the GFP-TREX1 fusion protein minimally affects TREX1’s effect on HIV-1 reverse transcription.
Next, we carried out TREX1-OE experiments in Jurkat cells—another T cell line permissive to HIV-1 infection. The pGfp-Trex1 fusion construct or the pGfp control was transfected into Jurkat cells, and TREX1-OE was confirmed by Western blotting ( Fig. 3G ). TREX1-OE and control Jurkat cells were inoculated with HIV-1 particles, and DNA from these cells was isolated. Quantification of reverse transcription products by qPCR revealed that the amount of HIV-1 DNA in TREX-OE cells was not significantly changed compared to that in the control cells ( Fig. 3H ). A lack of effect of higher levels of TREX1 in accumulation of HIV-1 DNA in Jurkat cells is consistent with the data obtained with Sup-T1 cells ( Fig. 3C ). Collectively, these results establish that higher levels of TREX1 minimally reduce the amount of reverse transcribed HIV-1 DNA in permissive T cell lines.
After HIV-1 reverse transcription, the viral DNA is integrated into the host genome to establish productive infection (18, 20,–22). Since TREX1 protein levels minimally affected HIV-1 reverse transcription ( Fig. 3 ), we next probed the effects of TREX1-OE on viral DNA integration to better understand the effects of HIV-1 infection-induced upregulation of TREX1. Parental Sup-T1 and TREX1-OE cells were inoculated with HIV-1 particles, and DNA isolated from these cells was subjected to qPCR to measure viral DNA integration. We quantified HIV-1 integration into the host genome by Alu-based nested PCR that amplifies the junction of the host DNA and integrated provirus (39,–41). The numbers of copies of the integrated viral DNA were determined through a standard curve generated in the second-round qPCR. Results from these analyses showed that HIV-1 integration was significantly higher in TREX1-OE cells than in the parental cells ( Fig. 4A ). Presence of the integrase inhibitor raltegravir (RAL) potently inhibited HIV-1 integration, demonstrating that these assays are specific for measuring integrated virus. The increased levels of HIV-1 integration were also observed in HEK293T cells transiently expressing TREX1 as a GFP fusion relative to the levels in control cells expressing only GFP ( Fig. 4D ). Similarly, significantly higher levels of HIV-1 integration into the host chromosomes were also observed in TREX1-OE Jurkat cells ( Fig. 4G ). Taken together, the proviral integration data from two different T cell lines and HEK293T cells strongly suggest that higher levels of TREX1 promote HIV-1 integration, consistent with increased viral integration observed in TREX1-OE THP-1 monocytic cells (32).
TREX1-OE enhances HIV-1 integration. Effects of TREX-OE on HIV-1 integration. Sup-T1 cells (A to C) and Jurkat cells (G to I) were inoculated with native envelope-containing HIV-1 particles, whereas HEK293T cells (D to F) were inoculated with VSV-G pseudotyped HIV-1 particles. Total DNA from these cells was subjected to Alu-based nested PCR for measuring viral DNA integration into the host genome. Copies of integrated viral DNA in the samples were calculated by interpolating data from a standard curve generated in parallel using 10-fold serial dilutions of known copies of an HIV-1 molecular clone. Effects of TREX-OE on HIV-1 nuclear import. Total DNA from uninfected and infected Sup-T1 (B), HEK293T (E), and Jurkat (H) cells were subjected to qPCR to amplify the junctions of the HIV-1 2-LTR circles. Copy numbers of the 2-LTR circles in the samples were calculated by interpolating data from a standard curve generated in parallel using 10-fold serial dilutions of known copy numbers of the p2LTR plasmid. Effects of TREX-OE on efficiency of HIV-1 nuclear import. Efficiency of viral nuclear import in Sup-T1 (C), HEK293T (F), and Jurkat (I) cells was determined by calculating the ratio of the copy numbers of 2-LTR circles (panels B, E, and H) to the copy number of the reverse transcripts ( Fig. 3C, F , and andH, H , respectively). The data are presented as relative percentage of parental infected cells. Data are mean values from at least three independent experiments, with error bars representing the SEMs. *, P < 0.05 for the comparison between parental cells and TREX-OE cells.
HIV-1 integration into the host genome is dependent on nuclear import of the reverse transcribed viral DNA (18, 20,–22). Since TREX1-OE enhanced HIV-1 integration ( Fig. 4A, D , and andG), G ), without significantly increasing reverse transcription ( Fig. 3C, F , and andH), H ), we next measured the viral nuclear entry step. We quantified the number of 2-long terminal repeat (2-LTR) circles as a surrogate for HIV-1 nuclear entry, since these circles exclusively accumulate in the nuclei of the infected cells through nonhomologous end joining (42,–46). Therefore, these circles are utilized as useful markers of HIV-1 nuclear entry even though they may represent nonproductive products (47). The copy numbers of 2-LTR circles were quantified through a standard curve generated using a plasmid containing the 2-LTR sequence (p2-LTR). Interestingly, quantification of 2-LTR circles in the infected Sup-T1 ( Fig. 4B ), HEK293T ( Fig. 4E ), and Jurkat ( Fig. 4H ) cells revealed that the copies of 2-LTR circles in TREX1-OE cells were not significantly changed compared to that in the control cells. Furthermore, calculation of the relative ratios between the copies of 2-LTR circles and reverse transcription products ( Fig. 4C, F , and andI) I ) also confirmed a lack of significant effect of TREX1-OE on HIV-1 nuclear entry. These results collectively establish that higher levels of TREX1 enhance HIV-1 integration without significantly affecting the viral nuclear entry step.
It has been reported that TREX1 depletion is associated with an accumulation of HIV-1 DNA in MEFs (30). However, in THP-1 monocytic cells, TREX-1 knockout (KO) minimally reduced levels of HIV-1 DNA (32). Furthermore, a recent report showed that HIV-1 DNA levels were not significantly affected in murine T cells lacking TREX1 protein (33). To better understand the effects of TREX1 depletion on early steps of HIV infection in human T cells, we carried out TREX1 knockdown (KD) experiments in Sup-T1 and Jurkat cells. We relied on KD studies instead of KO experiments due to the technical challenges of generating and culturing sufficient numbers of viable and healthy TREX1-KO cells.
Depletion of TREX1 was achieved through transduction of lentiviral particles producing short hairpin RNA (shRNA) specifically targeting the TREX1 mRNA. Western blot analysis confirmed the significant reduction in TREX1 levels in Sup-T1 ( Fig. 5A ) and Jurkat ( Fig. 5F ) cells. We observed that TREX1-KD efficiency in Sup-T1 cells was much greater than in Jurkat cells. Then, TREX1-KD and parental cells were inoculated with HIV-1 particles, and 24 hpi, cells were harvested. DNA isolated from these cells was subjected to qPCR to measure copies of reverse transcription ( Fig. 5B and andG), G ), 2-LTR levels ( Fig. 5D and andI), I ), and integrated HIV-1 DNA ( Fig. 5C and andH). H ). Results from these analyses illustrated that the level of reverse transcribed HIV-1 DNA in TREX1-KD cells is not significantly changed in Sup-T1 cells compared to that in the parental control cells ( Fig. 5B ). Similarly, in Jurkat cells, TREX1-KD did not significantly alter the levels of HIV-1 DNA ( Fig. 5G ). The lack of increased viral DNA in TREX1 KD T cell lines is counter to the observations in MEFs and THP-1 cells showing accumulation of HIV-1 DNA upon TREX-1 depletion (30, 33).
TREX1-KD reduces HIV-1 integration. Generation of TREX1-KD T cell lines. Depletion of TREX1 was carried out by transduction of lentiviral particles producing shRNA targeting TREX1 mRNA. TREX1-KD was confirmed by probing TREX1 levels in Sup-T1 (A) and Jurkat (F) cells by Western blotting. These cells were inoculated with HIV-1 particles. At 24 hpi, total DNA from these cells was subjected to qPCR. Quantification was carried out by interpolating the qPCR data from a standard curve generated in parallel using 10-fold serial dilutions of known copy numbers of a relevant molecular clone as described before. (B and G) Effects of TREX-KD on HIV-1 reverse transcription. (C and H) Effects of TREX-KD on HIV-1 integration. (D and I) Effects of TREX-KD on HIV-1 nuclear import. (E and J) Effects of TREX-KD on efficiency of HIV-1 nuclear import. Ratio of 2-LTR circle copy numbers (data from panels D and I) to the reverse transcript copy numbers (data from panels B and G). Data shown are mean values from at least three independent experiments, with error bars representing the SEMs. *, P < 0.05 for the comparison between parental cells and TREX-KD cells.
To further understand TREX1’s role during HIV-1 infection, we quantified the copies of integrated HIV-1 DNA in the genomes of TREX1-KD cells. Our qPCR results revealed that HIV-1 DNA integration was significantly lower in TREX1-KD cells than in the parental cells ( Fig. 5C and andH). H ). Furthermore, qPCR analysis of the copies of 2-LTR circles in these cells revealed that the number of 2-LTR copies were not significantly changed in TREX1-KD cells ( Fig. 4D and andI). I ). Finally, calculation of the relative ratios of the copies of 2-LTR circles to the reverse transcribed DNA indicated that TREX1-KD has minimal impact on HIV-1 nuclear entry ( Fig. 5E and andJ). J ). In summary, these observations demonstrated that TREX1-KD significantly reduces HIV-1 integration but not due to the consequence of reduced reverse transcription and/or nuclear entry.
To understand the mechanism by which TREX1 promotes integration, we biochemically studied HIV-1 PICs—the viral replication complexes that carry out integration of the viral DNA into the host genome (18, 20,–22). PICs extracted from acutely infected cells have been demonstrated to retain DNA integration activity in vitro (39, 48). Since higher levels of TREX1 enhanced HIV-1 integration without affecting nuclear entry ( Fig. 4 ) or reverse transcription ( Fig. 3 ), we probed whether the PICs from TREX1-OE cells retain higher integration activity. To test this, we inoculated TREX1-OE and parental Sup-T1 cells with DNase I-treated HIV-1 particles and prepared cytoplasmic extracts containing PICs (39, 40, 49). Then, the DNA integration activity of the PICs was assessed in an in vitro integration assay, and the copies of viral DNA integrated into the linear target DNA were quantified by qPCR (49, 50). As expected, PICs from parental Sup-T1 cells efficiently integrated the viral DNA into the target ( Fig. 6A ). The DNA integration activity was robustly inhibited by RAL, demonstrating the specificity of the in vitro integration assay ( Fig. 6A ). Interestingly, PICs from the TREX1-OE cells exhibited higher integration activity relative to the PICs extracted from the parental cells ( Fig. 6B ). For example, with the PICs from parental cells, approximately 2.2 × 10 6 copies of integrated viral DNA were quantified ( Fig. 6B ). However, with the PICs from TREX1-OE cells, a significantly higher number (∼3.8 × 10 6 copies) of integrated viral DNA was quantified ( Fig. 6B ).
TREX1 promotes DNA integration activity of HIV-1 PICs. (A) Quantification of PIC activity in vitro. Cytoplasmic PICs (Cy-PICs) were isolated from Sup-T1 cells inoculated with HIV-1 particles (1,500 ng p24 equivalent) as per our published method (49, 50). The in vitro integration activity of PICs was assessed by nested PCR in the presence or absence of the integrase inhibitor RAL. Control lysate refers to lysate from uninfected cells. The copy numbers of integrated viral DNA were determined by interpolating qPCR data from a standard curve generated in parallel using 10-fold serial dilutions of known copy numbers of an HIV-1 molecular clone. (B to D) Effects of TREX1-OE on the integration activity, viral DNA content, and specific activity of HIV-1 PICs. Parental and TREX1-OE Sup-T1 cells were inoculated with HIV-1 particles, and cytoplasmic PICs were extracted from these cells. (B) Integration activity of these PICs was measured in vitro, and the copy numbers of integrated viral DNA were determined as described above. (C) Copies of viral DNA in PICs isolated from parental and TREX1-OE Sup-T1 cells were quantified by qPCR using a standard curve. (D) Specific integration activity of PICs was determined by calculating the ratio of PIC-associated integration activity and PIC-associated viral DNA content. The data are presented relative to that of parental Sup-T1 cells. (E to G) Effects of TREX1-KD on the integration activity, viral DNA content, and specific activity of HIV-1 PICs. Parental and TREX1-KD Sup-T1 cells were inoculated with HIV-1 particles, and cytoplasmic PICs were extracted from these cells. (E) Integration activity of the PICs. (F) Copies of viral DNA in PICs isolated from parental and TREX1-KD Sup-T1 cells. (G) Specific integration activity of PICs The data are presented relative to that of parental Sup-T1 cells. (H to L) Effects of exogenous recombinant TREX1 (rTREX1) addition on HIV-1 PIC activity. (H) Quantification of PIC activity in vitro using plasmid DNA as the target. PICs from parental acutely infected Sup-T1 cells were assessed in vitro using a plasmid DNA target, and the copy numbers of integrated viral DNA were determined as described above. (I) PIC activity in the presence of various concentrations of rTREX1. PIC-associated integration activity in the absence or presence of increasing concentrations of rTREX1 protein was assessed, and the copy numbers of integrated viral DNA were determined. (J to L) Effects of rTREX1 addition on integration activity, viral DNA content, and specific activity of HIV-1 PICs. (J) PIC activity assessed in the presence or absence of 2 nM rTREX1 and copies of integrated viral DNA quantified. (K) Copies of viral DNA in PICs incubated with or without 2 nM rTREX1 protein were quantified by qPCR. (L) Specific activity was calculated as the ratio of PIC-associated integration activity and the PIC-associated viral DNA in the presence or absence of rTREX1. Data shown are mean values from at least three independent experiments, with error bars representing the SEMs. *, P < 0.05 for the comparison between untreated PICs and rTREX1-treated PICs.
Next, we examined whether the higher integration activity of PICs from TREX1-OE cells was due to the formation of an increased number of PICs. To test this, we measured the number of PIC-associated viral DNA by qPCR in the parental and TREX1-OE Sup-T1 cells. These analyses revealed that the levels of viral DNA in the PICs from TREX1-OE cells were not significantly changed compared to the levels in PICs from the parental cells ( Fig. 6C ). To account for the slight increase in copy number of the viral DNA in the TREX1-OE PICs, we calculated the specific activity of PICs, i.e., the ratio between integration activity and PIC-associated viral DNA copies ( Fig. 6D ). Results from these calculations revealed that the specific activity of PICs from TREX1-OE cells was significantly higher than that of the PICs from the parental cells ( Fig. 6D ). Collectively, these results strongly supported that higher levels of TREX1 promote integration activity of PICs.
Finally, to better understand TREX1’s role in PIC activity, we extracted PICs from TREX1-KD Sup-T1 cells and measured in vitro DNA integration activity, amount of viral DNA, and PIC-specific activity ( Fig. 6E to toG). G ). These analyses revealed that TREX1 depletion resulted in a marked reduction in PIC-associated viral DNA integration activity ( Fig. 6E ). As expected, the number of PIC-associated viral DNAs was not significantly changed in TREX-1 KD cells compared to that in the parental cell line ( Fig. 6F ). Nevertheless, PIC-specific activity calculations revealed a significant reduction in PIC activity due to TREX1 depletion ( Fig. 6G ). These biochemical analyses provide strong evidence that TREX1 levels promote HIV-1 DNA integration.
Our results suggested that higher levels of TREX1 enhance HIV-1 integration ( Fig. 4 ), most likely through enhanced PIC-associated viral DNA integration activity ( Fig. 6B and andE). E ). To further probe that TREX1 is directly and functionally involved in promoting PIC function, we measured DNA integration activity of PICs isolated from acutely infected parental Sup-T1 cells in the presence of purified TREX1 protein. We established a modified in vitro integration assay that utilized a circular plasmid DNA as the target rather than the linear DNA used in the previous experiments ( Fig. 6A ). This is mainly because the addition of TREX1 would degrade the linear target DNA through exonuclease activity. However, a circular plasmid DNA would be resistant to degradation by TREX1 (51) and therefore serves as an appropriate target for measuring PIC activity in the presence of TREX1. Measurement of PIC-associated integration activity showed efficient integration of the viral DNA into the plasmid target. As expected, the PIC activity was significantly inhibited in the presence of RAL ( Fig. 6H ). Next, we added a range of concentrations of purified TREX1 protein to the reaction mixture containing PICs extracted from infected parental Sup-T1 cells. Quantification of viral DNA integration illustrated that PIC activity was enhanced in the presence of TREX1 in a dose-dependent manner ( Fig. 6I ). Specifically, addition of 2 nM TREX1 to the PICs significantly increased the number of integrated viral DNA copies compared to that in the control PICs without added TREX1 ( Fig. 6I and andJ). J ). Interestingly, PIC-associated viral DNA levels were also significantly reduced with the addition of 2 nM TREX1 ( Fig. 6K ). This reduction is most likely due to degradation of the viral DNA in the PICs by the exonuclease activity of TREX1. Finally, calculation of specific activity established that addition of TREX1 significantly enhanced PIC-associated DNA integration activity ( Fig. 6L ). Taken together, these biochemical studies show that TREX1 directly promotes the DNA integration activity of HIV-1 PICs.
HIV-1 integration is dependent on a 3′ processing step, where the viral integrase enzyme removes GT dinucleotides from the 3′ ends of the reverse-transcribed viral DNA (19). Therefore, to better understand how TREX1 promotes PIC function, we employed exonuclease activity assays to test whether TREX1 preferentially degrades the unprocessed viral DNA that cannot integrate into the host genome (19). We prepared dsDNA substrates containing the unprocessed and 3′-processed viral DNA ends ( Fig. 7 ). The ssDNA oligonucleotides containing the U3 and U5 sequences of the viral DNA ends were fluorescently labeled at the 5′ ends. These substrates were annealed with the respective unlabeled cDNA strands and incubated with purified recombinant TREX1 enzyme. The reaction products were then loaded on a denaturing urea gel, separated by electrophoresis, and imaged to detect the DNA degradation products. Results from these assays show that TREX1 efficiently degraded the DNA substrates containing both the unprocessed U3 and U5 viral sequences ( Fig. 7A and andB, B , left). For instance, at 2 min (120 s), most of the unprocessed U3 and U5 substrates were degraded by TREX1 ( Fig. 7A and andB, B , left). However, the exonuclease activity of TREX1 was markedly less efficient in degrading the substrates containing the 3′-processed U3 and U5 viral DNA ends ( Fig. 7A and andB, B , right). Interestingly, the reduced level of degradation of the 3′-processed substrates was evident at each time point of the exonuclease activity measurement. To quantify the substrate preference, we calculated the percentage of substrates cleaved as a function of time. Interestingly, at the earliest time point of 15 s, ∼90% of the unprocessed U3 viral DNA was degraded ( Fig. 7C ). However, at the same time point, only ∼50% of the 3′-processed U3 viral DNA was degraded. Similarly, a significant difference in degradation was also detected with substrates containing U5 DNA ( Fig. 7D ). The reduced levels of degradation of the 3′-processed U3 and U5 substrates remained significant through the course of exonuclease activity measurement ( Fig. 7C and andD). D ). Importantly, the difference in the degradation was specific to the exonuclease activity of TREX1, since the catalytic mutant TREX1 D18N protein did not degrade any of the viral substrates ( Fig. 7E and andF). F ). Collectively, these observations provide biochemical evidence that TREX1 preferentially degrades the unprocessed viral DNA compared to the 3′-processed DNA.
TREX1 preferentially degrades unprocessed viral DNA substrates. Exonuclease activity of TREX1 was measured using viral DNA substrates containing the U3 (A, C, and E) and the U5 (B, D, and F) regions of the viral sequences; 20-mer (left) or 18-mer (right) 5′ fluorescein-labeled oligonucleotides were annealed with a 20-mer unlabeled complementary strand. Exonuclease activity was measured as a function of time and subjected to electrophoresis on 20% 7 M urea-polyacrylamide gels. The degradation products were imaged on a Molecular Imager. (C and D) Percentage of substrates degraded by TREX1. Data in panels A and B were analyzed by densitometry and plotted as percentage of remaining uncleaved U3 (C) and U5 (D) substrates as a function of time. (E and F) Exonuclease assays with catalytically inactive TREX1 (D18N) mutant. Increasing amounts of TREX1 D18N enzyme were incubated with the U3 (E) and the U5 (F) containing viral sequences. Samples were removed from 37°C after 60 min and subjected to electrophoresis on 20% 7 M urea-polyacrylamide gels. The degradation products were separated by electrophoresis under denaturing conditions. Data shown in panels A, B, E, and F are representative of at least three independent experiments. Data shown in panels C and D are from three independent experiments. *, P < 0.05 for the comparison between unprocessed and 3′-processed substrates at each time point.
TREX1 is a 3′ exonuclease and can remove 3′ nucleotides from both ends of dsDNA substrates (51, 52). Therefore, we determined whether simultaneous degradation from both the 3′ ends of the viral DNA contributed to the reduced exonuclease activity of the 3′-processed substrates. To test this, we utilized viral DNA substrates containing a 3′ phosphorothioate modification in the complementary strand. Phosphorothioate modification replaces one of the nonbridging oxygen atoms of the internucleotide linkage with sulfur and thus reduces degradation by nucleases (53). As expected, exonuclease activity confirmed that ssDNA substrates containing 3′ phosphorothioate modifications are resistant to degradation by TREX1 (data not shown). Next, the appropriate 3′ phosphorothioate ssDNA was annealed to the 5′-fluorescently labeled complementary viral (U3 or U5) DNA strand. These dsDNA substrates were incubated with TREX1, and the degradation products were analyzed. The degradation rate of phosphorothioate-modified dsDNA substrates by TREX1 was much lower than that of the unmodified substrates (data not shown). Therefore, the time course of exonuclease activity measurements was increased to 60 min for these phosphorothioate-modified substrates. Interestingly, the DNA degradation patterns of the 3′ phosphorothioate substrates ( Fig. 8 ) were similar to those of the unmodified substrates ( Fig. 7 ). For instance, the unprocessed U3 and U5 viral DNA substrates were degraded by TREX1 efficiently ( Fig. 8A and andB, B , left). However, similar to the data in Fig. 7 , degradation of the 3′-processed viral DNAs remained significantly lower as a function of time ( Fig. 8A and andB, B , right). Furthermore, calculation of the percentage of uncleaved substrates revealed that within 15 min, ∼50% of the unprocessed U3 and U5 substrates were degraded ( Fig. 8D and andE). E ). However, at the same time point, only ∼25% of the 3′-processed U3 substrates and ∼20% of the 3′-processed U5 substrates were degraded ( Fig. 8D and andE). E ). Interestingly, a significantly low percentage of 3′-processed viral DNA substrates was degraded at the 60-min time point compared to the percentage degraded of the unprocessed viral DNA ( Fig. 8D and andE), E ), akin to data in Fig. 7 .
Exonuclease activity of TREX1 on substrates with 3′-phosphorothioate modification. Exonuclease activity of U3 (A), U5 (B), and nonviral (C) DNA substrates. Exonuclease activity was measured as a function of time containing either 20-mer (left) or 18-mer (right) 5′ fluorescein-labeled oligonucleotides (250 nM), annealed with a 20-mer 3′ phosphorothioate-modified complementary strand. (D to F) Percentage of uncleaved substrates by TREX1 exonuclease activity. Data in panels A to C were analyzed by densitometry and plotted as percentage of substrates remaining; the U3 (D), the U5 (E), and nonviral (F) DNA substrates. Exonuclease assays with increasing amounts of TREX1. Increasing amounts of rTREX1 were incubated with the U3 (G), the U5 (H), and nonviral (I) substrates, and the degradation products were analyzed on 20% 7 M urea-polyacrylamide gels. Data shown in panels A to C and G to I are representative of three independent experiments. Data shown in panels D to F are from three independent experiments. *, P < 0.05 for the comparison between unprocessed and 3′-processed substrates at each time point.
Next, we tested whether the preference in the exonuclease activity of TREX1 is specific for the viral DNA substrates. We carried out exonuclease assays using 3′ phosphorothioate-modified nonviral DNA substrates containing a 20-nucleotide (nt) blunt end and an 18-nt 3′-recessed end. These substrates were specifically designed to mimic the unprocessed and 3′-processed viral DNA substrates used in Fig. 7 . Interestingly, exonuclease activity measurements revealed that TREX1 efficiently degraded both the nonviral DNA substrates efficiently irrespective of whether the DNA contained a blunt end ( Fig. 8C , left) or a 3′-recessed end ( Fig. 8C , right). Calculation of the percentage of uncleaved substrates confirmed comparable levels of degradation of these nonviral DNA substrates by TREX1 ( Fig. 8F ). The minor difference in degradation of these substrates as a function of time was not statistically significant ( Fig. 8F ).
Finally, we measured exonuclease activity of TREX1 in a concentration-dependent manner to further define the preference for the unprocessed viral DNA substrates ( Fig. 8G to toI). I ). Interestingly, the 3′-processed DNA substrates containing the U3 and U5 viral sequences resisted complete degradation, even at the highest TREX1 concentration ( Fig. 8G and andH, H , right). In contrast, the unprocessed viral DNA substrates were almost completely degraded at the highest concentration of the enzyme ( Fig. 8G and andH, H , left). The dose-dependent studies also revealed that both nonviral DNA substrates were efficiently degraded by TREX1 ( Fig. 8I ). Furthermore, the exonuclease activity was comparable for the substrate with a blunt or recessed 3′ end ( Fig. 8I ). These results also indicate that TREX1’s preference of degradation is specific to viral DNA substrates. Collectively, these studies provide strong biochemical evidence that the unprocessed viral DNAs are preferred for degradation by TREX1. In contrast, the 3′-processed viral DNA remains relatively resistant to the exonuclease activity of TREX1.
To better understand the biochemical role of TREX1 in the integration process, we studied the in vitro activity of HIV-1 intasomes. Intasome is a collective term utilized for stable viral nucleoprotein complexes formed during the retroviral integration (21, 54, 55). These nucleoprotein complexes are formed by a pair of viral DNA ends bridged by the viral integrase enzyme. The first intasome on the integration pathway is the stable synaptic complex (SSC), within which, two nucleotides are removed from the 3′ ends of the viral DNA to form the cleaved synaptic complex (CSC). The CSC then noncovalently captures the target DNA to form the target capture complex (TCC), within which, a pair of transesterification reactions occurs to form the strand transfer complex (STC).
Biochemical studies have established that SSC and CSC forms of HIV-1 intasomes can be assembled. Furthermore, these biochemically assembled intasomes retain the DNA strand transfer step of HIV-1 integration in the absence of other factors that are associated with the PICs formed in the infected cells (55,–57). Therefore, to probe whether the effects of TREX1 on PIC-mediated viral DNA integration is specific, we carried out in vitro integration assays using these intasomes. For this assay, we used a circular DNA substrate (pGEM) given that TREX1 degrades double-stranded DNA. First, activity measurement of HIV-1 SSCs showed efficient strand transfer of the viral DNA into the plasmid target ( Fig. 9A ). Next, a range of concentrations of purified TREX1 was added to the reaction mixture containing SSCs, and activity was measured. Relative activity data showed that DNA strand transfer activity of HIV-1 SSCs was significantly enhanced in the presence of TREX1. These data also illustrated a dose-dependent increase in intasome activity with increasing amount of TREX1 ( Fig. 9A ). Specifically, addition of 25 nM TREX1 increased the strand transfer activity almost 2-fold compared to that in the control ( Fig. 9A ). As expected, strand transfer activity was not observed with the negative-control reactions such as intasome only, substrate only, and TREX1 only. Finally, these measurements are specific to DNA strand transfer activity of SSCs, since the activity was significantly inhibited by a specific strand transfer inhibitor, RAL ( Fig. 9A ). These results provide further evidence for a direct role of TREX1 in HIV-1 integration and are consistent with the PIC data ( Fig. 6 ).
TREX1 enhances strand transfer activity of HIV-1 intasomes assembled with unprocessed viral DNA ends. The strand transfer activity of HIV-1 SSC intasomes (containing the unprocessed viral DNA) (A) and HIV-1 CSC intasomes (containing the 3′-processed viral DNA) (B) were measured using a circular DNA as a target by qPCR. Biochemically assembled intasomes (20 nM) were mixed with pGEM (300 ng) in a reaction mixture containing 20 mM HEPES (pH 7.5), 20% glycerol, 100 mM NaCl, 10 mM DTT, 4 μM ZnCl2, and 5 mM MgCl2 at 37°C for 1 h. The reaction was stopped by ethanol precipitation, and viral DNA strand transfer to the target was quantified by amplifying the junctions formed between the viral DNA and pGEM. The strand transfer inhibitor RAL was used to evaluate intasome specific activity. Data shown are from three independent experiments, with error bars representing the SEMs. *, P < 0.05 for the comparison between intasomes without and with TREX1 addition (A) and comparison between intasomes without and with RAL (B).
Finally, we carried out strand transfer activity measurements of HIV-1 CSC intasomes assembled with 3′-processed viral DNA. As expected, the CSCs were competent for inserting the viral DNA into the target plasmid, and the activity was significantly inhibited by RAL ( Fig. 9B ). Furthermore, the control reactions such as intasomes without target DNA, target pGEM substrate without intasomes, and TREX1 protein without the intasomes showed no measurable strand transfer activity. Interestingly, addition of purified TREX1 protein showed no measurable increase in the activity of the CSC intasomes, even at the highest concentrations of TREX1 ( Fig. 9B ). A lack of effect on the viral DNA strand transfer by CSCs is in contrast to the stimulating effects of added TREX1 on the activity of HIV-1 SSCs ( Fig. 9A ). Collectively, these results illustrate that the TREX1 addition specifically enhances DNA strand transfer activity of HIV-1 intasomes assembled with the unprocessed viral DNA and solidifies a biochemical role of TREX1 on HIV-1 integration.
The key function of TREX1 is to degrade cytosolic DNA originating from both exogenous and endogenous sources (2, 37). The presence of exogenous DNA, whether from a transfected plasmid, synthetic DNA, or intracellular pathogens such as bacteria or virus, activates the cellular ISD response (58). However, endogenous DNA originating from DNA replication/recombination/repair or retroelements does not activate the ISD response (4, 5). This is achieved by TREX1-mediated degradation of endogenous DNA to a level that cannot be detected by cellular DNA sensors and activate autoimmune reaction to self-DNA (4, 5). Accordingly, inhibition of the ISD response in TREX1-null mice by a reverse transcriptase inhibitor has solidified TREX1’s role in the degradation of DNA from retroelements (59). Endogenous retroelements represent ancient retroviruses that are permanently integrated into the host genome (4, 5). Therefore, it has been hypothesized that TREX1 protects retroviruses such as HIV-1 against the cellular ISD response. In this context, our results demonstrating upregulation of TREX1 in HIV-1-infected cells ( Fig. 1 ) warranted further studies to pinpoint the role of this host exonuclease during retroviral infection. Additionally, in infected cells, perinuclear accumulation of TREX1 was significantly enhanced, concurrent with colocalization with HIV-1 p24 ( Fig. 2 ). Most importantly, a significant albeit small amount of TREX1 protein was mobilized to the nuclei of infected cells. Collectively, these observations prompted us to study a potential role of TREX1 in pre- and post-nuclear entry steps of HIV-1 infection.
TREX1 has been implicated in HIV-1 reverse transcription, since depletion of the protein resulted in the accumulation of viral DNA in the infected cells (30). Therefore, we expected a smaller amount of viral DNA in cells with higher levels of TREX1. Surprisingly, the numbers of copies of reverse-transcribed HIV-1 DNA were not reduced in TREX1-OE cells compared to that in the cells with endogenous levels of TREX1 ( Fig. 3 ). Interestingly, a lack of reduction in the levels of HIV-1 strong-stop DNA has also been recently reported in TREX1-OE THP-1 cells (32). Accordingly, reverse transcription was not significantly altered in TREX-KD cells ( Fig. 5 ), establishing that the exonuclease minimally affects pre-nuclear entry steps of infection. Additionally, we predict that endogenous TREX1 could maintain the HIV-1 DNA levels below the detection limit of cellular DNA sensors—a hypothesis consistent with the lack of ISD response during HIV-1 infection (22). On the other hand, upregulation of TREX1 in infected cells pointed to a potential role of the cellular nuclease during early steps of infection. Therefore, we probed the nuclear entry of the virus in TREX1-OE and TREX1-KD cells by quantifying 2-LTR circles that are exclusively formed in the nucleus of the infected cell (60, 61). Interestingly, the number of 2-LTR circles was not significantly changed in either TREX1-OE ( Fig. 4 ) or TREX-1 KD ( Fig. 5 ) cells, consistent with a recent study of HIV-1 nuclear entry in THP-1 cells (32). Collectively, these results establish that TREX1 minimally affects nuclear entry of the reverse-transcribed HIV-1 DNA. It is important to note that nuclear entry of HIV-1 is dependent on the viral capsid protein (62) and a number of capsid-binding host factors (18, 63,–65). Since there is no evidence that TREX1 interacts with HIV-1 capsid or capsid-binding host factors, a lack of effect of TREX1 on viral nuclear entry is not surprising.
Given a lack of effect of TREX1 on HIV-1 reverse transcription and nuclear entry, we next probed TREX1’s role in post-nuclear entry steps of infection, specifically, the integration of viral DNA into the host genome. Interestingly, the numbers of copies of integrated viral DNA were significantly enhanced in TREX1-OE cells ( Fig. 4 ), and depletion of TREX1 resulted in a significant reduction in proviral integration ( Fig. 5 ). TREX1’s effect on HIV-1 integration was not a consequence of an effect on reverse transcription and/or nuclear entry, since there was no measurable effect of TREX1-OE or TREX1-KD on these two early steps of replication ( Fig. 4 and and5). 5 ). These studies suggested a role of TREX1 after the nuclear entry step of HIV-1 infection, most likely at the integration step. HIV-1 integration in vivo is carried out in the context of the PIC (66, 67). The PIC contains the viral DNA and associated cellular/viral factors, including the integrase enzyme that carries out integration of the viral DNA into the host chromatin (18,–21). Therefore, we predicted that TREX1 enhanced HIV-1 PIC function to promote viral DNA integration. Extraction of PICs and in vitro quantification of DNA integration activity revealed that PICs from TREX1-OE cells exhibited markedly higher activity ( Fig. 6B to toD). D ). Accordingly, PICs extracted from TREX1-KD cells retained significantly lower integration activity ( Fig. 6E to toG). G ). A direct role of TREX1 in HIV-1 integration was further supported by the enhanced integration activity of PICs in the presence of purified TREX1 protein ( Fig. 6I ). Calculation of PIC-specific activity also revealed that integration efficiency was significantly enhanced in the presence of TREX1 ( Fig. 6L ). The marked enhancement of PIC-specific activity was most likely due to a significant reduction in PIC-associated viral DNA in the presence of TREX1 protein ( Fig. 6K ). The reduction in viral DNA also implied that the viral DNA in a subset of PICs could be degraded by TREX1 exonuclease activity, whereas the viral DNA within the functional PICs is protected. The markedly greater effects of TREX1 on PIC activity in vitro is in stark contrast to the modest 2-fold effect of TREX1-OE on HIV-1 integration in cell-based assays ( Fig. 4 ). We predict that this could be a consequence of endogenous TREX1 being fully sufficient to degrade the unprotected or nonproductive viral DNA that could be associated with nonfunctional PICs. This contention is based on our observation that TREX-OE showed minimal effect in reducing the levels of reverse-transcribed viral DNA ( Fig. 3 ). Even though our results do not identify the factors that protect the viral DNA, it is plausible that binding of integrase to the viral DNA ends could protect it from degradation by TREX1 (48). Recently, we reported that HIV-1 capsid protein is part of the functional PICs (68). Therefore, PIC-associated capsid protein could protect the viral DNA from TREX1 by rendering a steric block. Nevertheless, the unprotected viral DNAs will be incompetent for integration. Thus, we predict that removal of such nonproductive viral DNA could enrich the functional PICs (with protected viral DNA ends) to efficiently access the target DNA and promote HIV-1 integration.
Integration of HIV-1 DNA into the host genome is dependent on the sequential 3′ processing and strand transfer steps carried out by the virally encoded integrase enzyme (19). During the 3′-processing step, the terminal GT dinucleotides are removed from the viral DNA to produce recessed 3′ CAOH ends (19). Subsequently, the 3′-processed viral DNA is integrated into the host genome by the strand transfer step, where integrase inserts the 3′ CAOH ends into the opposing strands of host chromosomal DNA (19). In contrast, the unprocessed viral DNA without the reactive 3′ CAOH ends is considered an abortive product, since it cannot integrate into the host genome (19). Therefore, we hypothesized that TREX1 preferentially degrades the unprocessed viral DNA to promote HIV-1 integration. This rationale was initially borne out by our results showing that addition of TREX1 reduced viral DNA in the PICs ( Fig. 6I ). Our exonuclease activity assays provided further biochemical support for this hypothesis ( Fig. 7 and and8). 8 ). For instance, TREX1 efficiently degraded the unprocessed blunt end viral DNA ( Fig. 7A and andB, B , left), but the 3′-processed DNA ends were not degraded efficiently and remained markedly resistant ( Fig. 7A and andB, B , right). This substrate preference of TREX1 was not due to the recessed 3′ ends, since nonviral substrates with either a blunt end or 3′-recessed end were efficiently degraded ( Fig. 8C ). Furthermore, the preference in degradation was specific to TREX1 exonuclease activity, given that catalytically inactive TREX1 D18N did not degrade any of these substrates ( Fig. 7E and andF). F ). Finally, results from exonuclease assays using 3′ phosphorothioate-modified substrates confirmed that the preference was not due to simultaneous degradation from both ends of the 3′-processed viral DNA ( Fig. 8A and andB). B ). These biochemical results demonstrate that the 3′-processed viral DNA ends with a recessed 3′ OH and a 2-nt 5′ overhang are resistant to TREX1 degradation.
TREX1 is well established to efficiently degrade a variety of dsDNA substrates (5, 37, 51, 69). Therefore, the inability of TREX1 to degrade the 3′-processed viral DNA ends was unexpected. However, there is evidence that the rate of exonuclease activity of TREX1 is reduced on dsDNA in regions of secondary structure (2, 37) and shows a preference for particular DNA sequences (70). Additionally, it has been reported that HIV-1 DNA ends may contain specific secondary structural determinants (48). Therefore, we hypothesized that the U3 and U5 sequences of the HIV-1 DNA may possess unique sequence and/or secondary structural features independently or in complex with the integrase enzyme that resist degradation by TREX1. To test this hypothesis, we probed effects of TREX1 on HIV-1 intasome activity. Intasomes are viral replication complexes that are formed by the HIV-1 integrase enzyme and the viral DNA. Interestingly, addition of TREX1 stimulated the strand transfer activity of HIV-1 SSC intasomes that contain the unprocessed viral DNA but not of CSCs assembled with the 3′-processed viral DNA ( Fig. 9 ). It should be noted that a recessed 3′ OH is essential for the strand transfer activity of HIV-1 intasomes in vitro. Therefore, the assembled CSCs with 3′-processed viral DNA are primed for strand transfer activity. In contrast, in the SSCs, the terminal 3′ CA dinucleotides of the viral DNA must be removed prior to strand transfer. Therefore, based on our results, we predict that TREX1 could interact with HIV-1 integrase of the SSC to induce conformational changes that stimulate integrase-mediated removal of the 3′ CA dinucleotides. This prediction is based on the presence of a polyproline helix (PPII helix) domain in TREX1 that interacts with proteins containing src homology 3 (SH3) domain (71). Accordingly, the C-terminal domain (CTD) of HIV-1 integrase that interacts with the viral DNA has a five-stranded beta-barrel that resembles an SH3 domain (72,–74). Alternatively, the 3′ exonuclease activity of TREX1 could remove a couple of nucleotides from the 3′ end of the unprocessed viral DNA to produce a recessed and reactive 3′ OH, akin to the integrase-mediated 3′ processing step. However, future studies are warranted to test these hypotheses and establish a functional interaction between TREX1 and HIV-1 intasome.
Recently, there were reports suggesting that HIV-1 reverse transcription may not be completed in the cytoplasm of the host cell (75, 76). Thus, there is a possibility that cytosolic TREX1 cannot access and degrade the unprocessed viral DNA located in the nucleus. It is important to point out that TREX1 has well-established functions both in the cytoplasm and nuclei of mammalian cells. As a component of the SET complex (38), TREX1 translocates to the nucleus in response to granzyme A-activated DNA damage (38, 77). In the nucleus, TREX1 removes ssDNA nicks to prevent rejoining of the nicked DNA ends (38, 77). Furthermore, there is evidence that in the nucleus, TREX1 associates with the DNA replication foci to attenuate chronic activation of the ataxia telangiectasia mutated-dependent DNA damage response (5). Interestingly, our imaging and biochemical studies highlight that TREX1 is ER associated and localized at the perinuclear regions ( Fig. 2 ). Additionally, in infected cells, TREX1 is colocalized with the HIV-1 capsid protein (p24) near the periphery of the nuclear envelope ( Fig. 2 ). Therefore, it is likely that TREX1 can access and degrade the unprocessed viral DNA during viral nuclear entry. Furthermore, our data show that a portion of the TREX1 protein is mobilized to the nuclei of infected cells ( Fig. 2 ). The nuclear presence of TREX1 could be utilized by the virus to degrade the unprocessed viral DNA after the nuclear entry step of infection. This could also reduce and/or remove the nonfunctional PICs in the nucleus and promote specific activity of functional PICs. Furthermore, based on the stimulating effects of TREX1 on strand transfer activity of HIV-1 SSCs ( Fig. 9 ), it is feasible that the nuclear TREX1 can remodel the HIV-1 intasome to enhance specific DNA integration activity of PICs.
Taken together, our results show that HIV-1 infection is associated with elevated levels of TREX1. We also provide evidence for a direct and functional role of TREX1 in HIV-1 integration. The stimulating effects of TREX1 are dependent on removal of nonfunctional PICs and reveal a novel mechanism by which TREX1 promotes early steps of HIV-1 infection. These observations are also significant in demonstrating an ISD-independent role of the cellular exonuclease during HIV-1 infection.
Raltegravir (RAL) and efavirenz (EFV) were obtained from the NIH AIDS reagent program, Division of AIDS, NIAD, NIH. Geneticin G148, puromycin, doxycycline, and Lipofectamine 2000 were obtained from Thermo Fisher scientific (Waltham, MA, USA). Polyethylenimine (PEI) 25000 was obtained from Polysciences, Inc. (Warrington, PA, USA).
Virus particles were generated from full-length HIV-1 infectious molecular clone pNL4-3 or pNL4-3.Luc.R-.E-, which is defective for expression of Nef, Env, and Vpr (78). The plasmid pMD 2.G encoding VSV-G was used to generate VSV-G pseudotyped HIV-1 particles (79). pMD2.G was a gift from Didier Trono through Addgene (Cambridge, MA, USA). The pGFP-TREX1 plasmid used to overexpress TREX1 was obtained from Judy Lieberman through Addgene. The TREX1 empty vector control plasmid was generated by first digesting pGFP-TREX1 with BglII and XbaI, followed by blunt-end generation with DNA polymerase I Klenow fragment and subsequent intramolecular ligation with T4 DNA ligase. To generate an inducible lentiviral TREX1 construct, a 1,013-bp DNA fragment containing the TREX1 coding sequence was obtained from Genewiz and was cloned into the pTRIPZ-FBXW7 plasmid provided by Christophe Nicot (University of Kansas Medical Center), from which the FBXW7 insert was removed by restriction digestion. The TREX1 insert was engineered to contain an upstream Agel site, Kozak sequence, a human codon-optimized flag sequence, a five amino acid linker, and the TREX1 open reading frame (ORF) flanked by an MluI site. Cloning of the TREX1 coding sequence and other elements was confirmed by sequencing. The psPAX2 packaging plasmid and pMD2.6 envelope plasmid were obtained from Addgene, Cambridge, MA.
All enzymes were obtained from New England BioLabs (Ipswich, MA, USA). Competent Stbl2, DH5α, or NEB-Stbl bacterial cells were used for transformation, plasmids were isolated using Zyppy plasmid prep kit (Zymo Research, Irvine, CA, USA), and recombinant plasmids were verified by restriction enzyme digestion and Sanger DNA sequencing.
TREX1 was probed with the rabbit TREX1 antibody purchased from Cell Signaling Technology (Danvers, MA, USA). β-actin was probed with the mouse β-actin antibody purchased from Santa-Cruz Biotechnology (Dallas, TX, USA). GFP was probed with rabbit GFP antibody purchased from Cell Signaling Technology. Antibody to HIV-1 capsid protein (CA) was obtained from the NIH AIDS reagent program, Division of AIDS, NIAD, NIH. Antibodies to lamin and calreticulin were obtained from Abcam (Cambridge, MA, USA). The GAPDH antibody was obtained from Cell Signaling Technology, and Flag-conjugated horseradish peroxidase (HRP) was obtained from Sigma-Aldrich (St. Louis, MO, USA).
Jurkat, Sup-T1, and human embryonic kidney 293T (HEK293T) cell lines were obtained from the American Type Culture Collection (Manassas, VA, USA). The TZM-bl reporter cell line was from John C. Kappes, Xiaoyun Wu, and Tranzyme, Inc., through the NIH AIDS Reagent Program, Division of AIDS, NIAID, NIH. Jurkat and Sup-T1 cells were grown in RPMI 1640 medium; HEK293T and TZM-bl cells were grown in Dulbecco modified Eagle medium (DMEM). All cell culture media were supplemented with 10% heat-inactivated fetal bovine serum (hi-FBS), 2 mM glutamine, 1,000 U/ml penicillin, and 100 mg/ml streptomycin, all obtained from Thermo Fisher Scientific (Waltham, MA, USA). All cells were cultured in a 37°C incubator with 5% CO2.
Whole blood from healthy donors was purchased from the New York Blood Center (New York, NY, USA) using the Meharry Medical College Institutional Review Board (IRB) terms and regulations. PBMCs from blood were isolated using the Ficoll-based method as previously described with slight modifications (49). In brief, human blood was diluted 1:1 with phosphate-buffered saline (PBS) supplemented with 2% FBS, and 25 ml diluted blood was overlaid on 12.5 ml Ficoll‐Paque premium reagent (GE Healthcare Life Sciences, Pittsburgh, PA, USA) in SepMate tubes purchased from STEMCELL Technologies (Vancouver, BC, Canada) and centrifuged at 750 × g. Thereafter, the interphase cells (PBMCs) were transferred carefully to a new 50-ml tube, centrifuged several times, and washed with PBS supplemented 2% with FBS to remove unwanted cell types. Red blood cells (RBCs) were removed through the addition of RBC lysis buffer as per the manufacturer’s instructions (Thermo Fisher scientific).
PBMCs were cultured at a density of 2 × 10 6 cells/ml in RPMI medium supplemented with 20% heat-inactivated fetal bovine serum, 2 mM glutamine, 1,000 U/ml penicillin, and 100 mg/ml streptomycin, all obtained from Thermo Fisher scientific.
The Sup-T1 cell line stably expressing GFP-TREX1 was generated by electroporation using the Neon transfection system (Thermo Fisher, Waltham, MA, USA). Sup-T1 cells (2 × 10 5 ) were electroporated with pEGFP-TREX1 plasmid (3 μg) using the following electroporation conditions: voltage, 1,400 V; pulse width, 20 ms; number of pulses, 2. These cells were then cultured in RPMI selection medium containing G418 (500 μg/ml) and monitored daily for GFP fluorescence under a fluorescence microscope. Once a majority of the cells were positive for GFP fluorescence, they were subjected to fluorescence-activated cell sorting (FACS). The top 10% cell population with the highest GFP intensity was selected and sorted. The sorted cells were subsequently cultured in selection medium containing G418 (500 μg/ml). Before using these cells, TREX1 overexpression in these cells was verified by immunoblot and fluorescence microscopy.
Jurkat cells overexpressing TREX1 were generated through transduction of lentiviral particles containing a TREX1 expression construct. To generate these lentiviral particles, HEK293T cells (2.2 × 10 6 cells) in a 10-cm dishes were transfected with psPAX2, pMD 2.G, and pTRIPZ flag-TREX1 plasmid using a 1:3 ratio of DNA to PEI. After 8 h, the medium was changed; 60 h after the transfection, culture supernatants containing the lentivirus particles were harvested, cleared of debris by low-speed centrifugation, and filtered through a 0.45-μM filter. The number of infectious particles was determined using the qPCR lentivirus titer kit obtained from Applied Biological Materials Inc. (Richmond, BC, Canada). Jurkat cells (6 × 10 6 cells) were inoculated with lentivirus particles (multiplicity of infection [MOI] of ∼5) by spinoculation (480 × g) at 25°C for 2 h. At 24 h postransduction, the cells were gently pelleted, washed once with PBS, and gently resuspended in media containing 2.5 μg/ml of puromycin for selection and 1 μg/ml doxycycline for TREX1 induction. TREX1 expression in these cells was verified by immunoblotting prior to use.
Sup-T1 or Jurkat cells depleted of endogenous TREX1 were generated by transduction with lentiviral particles producing short hairpin RNA (shRNA) specifically targeted for TREX1 mRNA. Briefly, lentiviral particles were produced by cotransfecting HEK293T cells (3 × 10 6 ) with pLKO shTREX1 (Dharmacon Inc., Chicago, IL, USA), psPAX2 packaging plasmid (Addgene, Cambridge, MA, USA), and pMD2.6 envelope plasmid (Addgene). At 12 h posttransfection, cells were carefully washed once with PBS, and then complete DMEM medium was added. The virus-containing supernatant was collected at 60 h after transfection, cleared of debris by low-speed centrifugation, and subsequently filtered through a 0.45-μM filter. Sup-T1 and Jurkat cells (1 × 10 6 ) were transduced with pLKO shTREX1 lentiviral particles by spinoculation (480 × g) at 25°C for 2 h and then were incubated at 37°C with 5% CO2. At 24 h postransduction, the cells were gently pelleted, washed once with PBS, and gently resuspended in RPMI medium supplemented with 1 μg/ml puromycin. One week after selection, the concentration of puromycin was increased to 2.5 μg/ml. Before using these cells, TREX1 expression in these cells was verified by immunoblotting.
To detect proteins by immunoblotting, whole-cell lysates were prepared in radioimmunoprecipitation assay (RIPA) buffer supplemented with protease inhibitor cocktail and phenylmethylsulfonyl fluoride (PMSF), 10 μg/ml (all obtained from Sigma-Aldrich, St. Louis, MO, USA) according to the manufacturer’s protocols. Protein concentrations were determined using bicinchoninic acid (BCA) protein assay reagent (Thermo Fisher Scientific, Waltham, MA, USA) as per the manufacturer-recommended protocol. Equivalent amounts of protein from whole-cell lysates were electrophoresed on NuPAGE 4% to 12% bis-Tris protein gels (Thermo Fisher Scientific) and electrophoretically transferred to nitrocellulose membranes using a Trans-Blot SD semi-dry transfer cell (Bio-Rad Laboratories, Hercules, CA, USA). The membranes were blocked in blocking buffer (5% [wt/vol] nonfat milk in Tris-buffered saline containing 0.05% Tween 20 [TBST], pH 8.0]). Thereafter, the membranes were probed with the appropriate antibody diluted in blocking buffer (rabbit anti-TREX1 at 1:1,500 [vol/vol], mouse anti-β-actin at 1:10,000 [vol/vol], rabbit anti-GFP at 1:10,000, and mouse anti-CA at 1:1,000 [vol/vol]) and subsequently with secondary antibody conjugated to horseradish peroxidase (anti-rabbit antibody at 1:10,000 [vol/vol]; anti-mouse antibody at 1:10,000 [vol/vol]). The membranes were washed with TBST buffer several times, and immunocomplexes were detected by clarity enhanced chemiluminescence (ECL) method (Bio-Rad Laboratories). Densitometry analysis was performed through either LI-COR Image Studio Digits version 5.2 software (LI-COR, Lincoln, NE, USA) or using ImageJ (v1.51). Data were normalized to levels of loading controls such as β-actin, GAPDH, or other required proteins.
To examine the cellular distribution and localization of TREX1, we employed confocal microscopy-based imaging. HEK293T or TZM-bl cells (5 × 10 5 cells) were seeded on poly- l -lysine-coated coverslips in a 6-well culture plate. Then, cells were transfected with GFP-TREX1 or GFP plasmid using Lipofectamine 2000 (Thermo Fisher Scientific, Waltham, MA, USA). Twenty-four hours posttransfection, VSV-G pseudotyped or wild-type HIV-1 stocks prepared in DMEM containing Polybrene (6 μg/ml) were added to cells. Twenty-four hours postinoculation, the medium was aspirated and cells were washed once with PBS. Cells were fixed using 3.7% (wt/vol) paraformaldehyde (PFA) for 15 min at room temperature. Fixing solution was carefully removed, and the cells were then washed three times with PBS. Cells were then incubated in permeabilization buffer (0.1% Triton X-100 in PBS) for 5 min at room temperature. Subsequently, the cells were washed three times with PBS before 0.1 M glycine was added to quench unreacted aldehydes. After 10 min, the glycine was removed, and blocking solution (5% bovine serum albumin [BSA] in PBS) was added. After 30 min, the blocking solution was removed, and the anti-HIV-1 p24 KC57-phycoerythrin (PE) conjugate (ARP-13449) at a 1:100 dilution in 3% BSA in PBS was added and incubated at 4°C overnight. The next day, the primary antibody was removed and the samples were carefully washed three times for 5 min with PBS. The coverslips were then carefully removed, mounted to slides with Diamond antifade mounting medium with 4′,6-diamidino-2-phenylindole (DAPI) (Thermo Fisher Scientific), and dried overnight at room temperature in the dark. Imaging was performed using a Nikon A1R confocal laser scanning microscope. The excitation/emission wavelengths were set at 405/425 to 475 nm for DAPI, 488/500 to 550 nm for green fluorescence, and 561/570 to 620 nm for red fluorescence. Regions of interest (ROIs) were drawn around the nuclei to determine the perinuclear localization of TREX1. ROIs were drawn at areas of TREX1 puncta to determine colocalization with DAPI via Pearson’s correlation coefficients using Nikon Elements Advanced Research imaging software and GraphPad prism (GraphPad Software, La Jolla, CA).
Virus stocks of VSV-G pseudotyped and wild-type HIV-1 were generated using the molecular clone of HIV-1 pNL4.3 by transfection of HEK293T cells using polyethylenimine (PEI) according to our published methods (68, 80). Briefly, VSV-G pseudotyped HIV-1 particles were produced by cotransfecting HEK293T cells with one part pMD2.G (VSV-G) and 3 parts pNL4-3.Luc.R-.E- plasmids. Wild-type envelope HIV-1 particles were produced by transfecting HEK293T cells with pNL4.3 plasmid. At 12 h posttransfection, cells were carefully washed with PBS, and medium was replaced with complete DMEM. For generating infectious HIV-1 virions, we used the supernatants of chronically infected ACH-2 cells that produce the X4 tropic HIV-1 LAI virions, per our published methods (81, 82). In brief, ACH-2 cells were cultured in RPMI medium in the presence of phorbol myristate acetate (PMA) and tumor necrosis factor alpha (TNF-α), and the virus-containing supernatant was collected 48 h after transfection. The supernatant was cleared of large cellular debris by low-speed centrifugation and filtered through a 0.45-μM filter. The virus particles generated from HEK293T cell transfection were treated with 40 U/ml of DNase I (Calbiochem/EMD, San Diego, CA, USA) to remove any contaminating plasmid DNA. Infectivity of the virus was determined by luciferase reporter assay using TZM-bl cells, which harbor a firefly luciferase reporter gene under the control of the HIV-1 Tat-inducible LTR promoter (83). Virus concentrations were determined using the p24-specific enzyme-linked immunosorbent assay (ELISA) methods as per our published protocol (68, 80).
Infection experiments were carried out by inoculating these cells with pseudotyped or wild-type (WT) HIV-1 particles (68, 80). HEK293T or TZM-bl (2.5 × 10 5 cells per well) were seeded in 6-well plates, cultured overnight, and then washed with fresh medium. Virus stocks prepared in DMEM containing Polybrene (6 μg/ml) and either dimethyl sulfoxide (DMSO), 1 μM RAL, or 5 μM EFV were added to the cells. Sup-T1 or Jurkat cells (4 × 10 6 cells per well) in 6-well plates were spinoculated at 480 × g with HIV-1 virions and either DMSO, 1 μM RAL, or 5 μM EFV at 25°C for 2 h. The cells were then cultured for 24 to 48 h at 37°C with 5% CO2 and harvested for further use. PBMCs were cultured at a density of 2 × 10 6 cells/ml in RPMI medium and activated with 5 μg/ml PHA, all obtained from Thermo Fisher Scientific (Waltham, MA, USA). Two days after activation, cells were pelleted by centrifugation at 100 × g for 4 min and resuspended in RPMI medium containing 5 μg/ml PHA and 20 U/ml of IL-2 (Sigma-Aldrich, St. Louis, MO, USA). Activated PBMCs (5 × 10 6 cells) were infected with HIV‐1 LAI particles by spinoculation in the presence of Polybrene (Bio-Rad Laboratories, Hercules, CA, USA), as described before (81, 82). Productive infection of cells was measured by detecting intracellular HIV‐1 p24 protein either by immunoblotting or FACS, as per our published methods (49, 81, 82). Total DNA from the infected and uninfected control cells was isolated using a Quick-DNA miniprep kit according to the manufacturer-recommended protocol (Zymo Research, Irvine, CA, USA) for further analysis.
To measure reverse transcription products and 2-LTR circles, we employed a qPCR-based strategy (68, 80). Reverse transcription products were quantified by utilizing a SYBR green-based qPCR which contained 100 ng of total DNA, 1× iTaq universal SYBR green supermix (Bio-Rad Laboratories, Hercules, CA, USA), and 300 nM late RT F (5′-TGTGTGCCCGTCTGTTGTGT-3′) and late RT R (5′-GAG TCCTGCGTCGAGAGAGC-3′) primers. The 2-LTR circles were quantified by utilizing a SYBR green-based qPCR which contained 100 ng of total DNA, 1× iTaq universal SYBR green supermix (Bio-Rad Laboratories, and 300 nM 2-LTR forward (5′-AACTAGGGAACCCACTGCTTAAG-3′) and 2-LTR reverse (5′-TCCACAGATCAAGGATATCTTGTC-3′) primers. The qPCR conditions consisted of an initial denaturation at 95°C for 3 min, followed by 39 cycles of amplification and acquisition at 94°C for 15 s, 58°C for 30 s, and 72°C for 30 s. To quantify the reverse transcription products, a standard curve was generated in parallel and under same conditions using 10-fold serial dilutions of known copy numbers (1 × 10 0 to 1 × 10 8 ) of the HIV-1 molecular clone plasmid. Similarly, to calculate the 2-LTR copies, a standard curve was generated using 1 × 10 0 to 1 × 10 8 of the p2-LTR plasmid. Copy numbers of reverse transcription and 2-LTR circles were determined by plotting the qPCR data against the respective standard curve. The qPCRs were performed in triplicates, and the data were analyzed using CFX Maestro software (Bio-Rad Laboratories).
To measure HIV-1 proviral DNA, a nested PCR method was used that consisted of a first-round endpoint PCR with primers designed to amplify only the integration junctions between human Alu repeats and HIV-1 viral DNA but not the unintegrated viral DNA, followed by a second-round qPCR with primers designed to specifically amplify only the viral LTR from the first-round PCR products (68, 80). The first-round PCR contained 100 ng of total DNA, 1× Bestaq reaction buffer (Applied Biological Materials Inc., Richmond, BC, Canada), deoxyribonucleotide triphosphate (dNTP) mix containing 200 μM concentrations of each nucleotide (Promega, Madison, WI, USA), 500 nM primers targeting Alu repeat sequence (5′-GCCTCCCAAAGTGCTGGGATTACAG-3′) and HIV-1 Gag sequence (5′-GTTCCTGCTATGTCACTTCC-3′), and 1.25 U of Bestaq DNA polymerase in a 50-μl final volume. The first-round PCR conditions consisted of an initial incubation at 95°C for 5 min, followed by 23 cycles of amplification at 94°C for 30 s, 50°C for 30 s, and 72°C for 4 min, and a final incubation at 72°C for 10 min. The second-round qPCR consisted of one-tenth of the product from the first-round PCR as the template DNA, 1× iTaq universal probe supermix (Bio-Rad Laboratories, Hercules, CA, USA), 300 nM (each) the viral LTR-specific primers that target the R region (5′-TCTGGCTAACTAGGGAACCCA-3′) and the U5 region (5′-CTGACTAAAAGGGTCTGAGG-3′), and 100 nM TaqMan probe (5′-6-carboxyfluorescein [FAM]-TTAAGCCTCAATAAAGCTTGCCTTGAGTGC-6-carboxytetramethylrhodamine [TAMRA]-3′). To quantify HIV-1 integration in the TREX1 knockdown cells, the LTR-specific primers and probes used were shTREX1 LTR forward (5′-CACAAGGCTACTTCCCTGATT-3′), shTREX1 LTR reverse (5′-CTCCTTCATTGGCCTCTTCTAC-3′), and probe (5′-FAM-TCCACTGACCTTTGGATGGTGCTT-TAMRA-3′). The qPCR conditions consisted of an initial incubation at 95°C for 3 min, followed by 39 cycles of amplification and acquisition at 94°C for 15 s, 58°C for 30 s, and 72°C for 30 s. During qPCR of the samples, a standard curve was generated in parallel and under same conditions using 10-fold serial dilutions of known copy numbers (1 × 10 0 to 1 × 10 8 ) of the HIV-1 molecular clone plasmid. The qPCR experiments were performed in triplicates, and the data were analyzed using CFX Maestro software (Bio-Rad Laboratories). The integrated viral DNA copy numbers were calculated by plotting the qPCR data against the standard curve.
HIV-1 PICs were isolated from infected T cells using a modified method previously published (49, 68, 84). Sup-T1 cells (4 × 10 8 ) distributed equally in 6-well plates were spinoculated at 480 × g with WT HIV-1 virions for 2 h at 25°C, and the cells were then cultured for 5 h at 37°C. The cells were pelleted by centrifugation for 10 min at 300 × g. The supernatant was then carefully aspirated, and the cell pellet was washed twice with 2 ml of K−/− buffer (20 mM HEPES [pH 7.6], 150 mM KCl, 5 mM MgCl2) at room temperature. The pellet was then gently lysed by resuspending in 2 ml of ice-cold K+/+ buffer (20 mM HEPES [pH 7.6], 150 mM KCl, 5 mM MgCl2, 1 mM dithiothreitol [DTT], 20 μg/ml aprotinin, 0.025% [wt/vol] digitonin) and rocking for 10 min at room temperature. The cytoplasmic extract containing the PICs (Cy-PICs) was then separated from other cellular components by differential centrifugation for 4 min at 1,500 × g at 4°C. The supernatant was transferred to a fresh microcentrifuge tube and centrifuged for 1 min at 16,000 × g at 4°C. The resulting supernatant was again transferred to a fresh microcentrifuge tube and treated with RNase A (Thermo Fisher, Waltham, MA, USA) for 30 min at room temperature to remove any cellular and or viral RNA. Finally, 60% sucrose (wt/vol) in K−/− buffer was added to a final concentration of 7% and gently mixed by pipetting. These PICs were then aliquoted, flash frozen in liquid nitrogen, and then stored in a −80°C freezer.
To determine the effect of TREX1 expression on HIV-1 PIC integration activity, PICs from control, TREX1-OE, and TREX1-KD cells were extracted. Activity of these PICs was measured by in vitro integration assays as previously described with slight modification (49, 68, 84). In brief, 300 ng of purified ФX174 target DNA was added to 100 μl of PICs and mixed well. The reaction mixture was incubated for 45 min at 37°C. The reactions were stopped and deproteinized by the addition of 0.5% SDS, 8 mM EDTA, and 0.5 mg/ml proteinase K followed by overnight incubation at 55°C. DNA was isolated the next day by phenol chloroform extraction followed by ethanol precipitation as per our published method (49, 68, 84).
To measure PIC integration activity, a nested PCR was employed. The first-round endpoint PCR amplified the integration junctions by utilizing primers designed to bind to the viral LTR and the target DNA. The first-round PCR consisted of 5 μl of DNA purified from the PIC assay reaction, 300 nM viral DNA-specific primer (5′-GTGCGCGCTTCAGCAAG-3′), 300 nM target DNA-specific primer (5′-CACTGACCCTCAGCAATCTTA-3′), 1× Bestaq reaction buffer, dNTP nucleotide mix containing 200 μM concentrations of each nucleotide, and 1.25 U of Bestaq DNA polymerase in a 50-μl final volume. The cycling conditions for the first-round PCR consisted of an initial denaturation step at 95°C for 5 min, followed by 23 cycles at 94°C for 30 s, 55°C for 30 s, and 72°C for 4 min, and a final extension at 72°C for 10 min.
The second-round qPCR utilized primers designed to specifically amplify the viral LTR from the first-round PCR products. The second-round qPCR contained one-tenth of the product from the 1 st -round PCR, 1× iTaq universal probe supermix, 300 nM viral LTR-specific primer that targets the R region (5′-TCTGGCTAACTAGGGAACCCA-3′), 300 nM U5 region-specific primer (5′-CTGACTAAAAGGGTCTGAGG-3′), and 100 nM TaqMan probe (5′-FAM-TTAAGCCTCAATAAAGCTTGCCTTGAGTGC-TAMRA-3′). The qPCR settings consisted of an initial incubation at 95°C for 3 min, followed by 39 cycles of amplification and acquisition at 94°C for 15 s and 58°C for 30 s, and a final incubation at 72°C for 30s. During qPCR of the samples, a standard curve was generated in parallel and under same conditions using 10-fold serial dilutions of known copy numbers (1 × 10 0 to 1 × 10 8 ) of the HIV-1 molecular clone plasmid. Integrated viral DNA copy numbers were determined by plotting the qPCR data against the standard curve. The qPCRs were performed in triplicates, and the data were analyzed using the CFX Maestro software (Bio-Rad Laboratories, Hercules, CA, USA).
To biochemically investigate the effect of TREX1 on HIV-1 PIC-associated DNA integration activity, we used purified recombinant human TREX1 protein. Purified TREX1 protein was prepared according to the published protocol (71). HIV-1 PICs were incubated with a range of concentrations of recombinant TREX1 and a circular target DNA (pGEM) instead of the linear ФX174 target DNA in the routine PIC assay, since circular DNA is inherently resistant to the exonuclease activity of TREX1 (51). The reaction contained 100 μl of PICs, recombinant TREX1 (0 nM tp 4 nM), and 300 ng of pGEM DNA. The reaction mixture was incubated for 1 h at 37°C and was stopped by the addition of 0.5% SDS, 8 mM EDTA, and 0.5 mg/ml proteinase K followed by incubation overnight at 55°C. DNA was isolated the next day by phenol chloroform extraction followed by ethanol precipitation. To quantify the PIC-associated DNA integration activity in the presence of TREX1, we used a nested PCR. The first-round endpoint PCR used primers designed to amplify the integration junctions between the viral DNA and the target pGEM plasmid DNA. The first-round PCR consisted of 5 μl of DNA purified from the PIC assay reaction, 300 nM viral DNA-specific primer (5′-CAATATCATACGCCGAGAGTGCGCGCTTCAGCAAG-3′), 300 nM pGEM-specific primer (5′-GTCACGACGTTGTAAAACGACG-3′), 1× Bestaq reaction buffer, dNTP nucleotide mix containing 200 μM concentrations of each nucleotide, and 1.25 U of Bestaq DNA polymerase in a 50-μl final volume. The cycling conditions for the first-round PCR consisted of an initial denaturation step at 95°C for 5 min, followed by 23 cycles at 94°C for 30 s, 55°C for 30 s, and 72°C for 4 min, and a final extension at 72°C for 10 min. The second-round qPCR was carried out as described in the previous section, and the integrated viral DNA copy numbers were determined by plotting the qPCR data against the standard curve.
To examine cellular distribution of TREX1, we analyzed cytoplasmic and nuclear fractions of uninfected and infected cells based on a published method with modifications (85). TZM-bl cells (3 × 10 6 ) were seeded in 10-cm cell culture plates. HIV-1 stocks were prepared in DMEM containing Polybrene (6 μg/ml) and were added to the cells. The plates were incubated at 37°C with 5% CO2. Cells were washed once with 1× PBS. Cells were then incubated with 0.25% trypsin (Thermo Fisher, Waltham, MA, USA) to detach them from the plate. Once the cells were completely detached, 8 ml of complete DMEM was added to the plate, and the mixture was transferred to a 15-ml Falcon tube on ice. The plate was washed once more to collect any residual cells with 5 ml of complete DMEM, which was then added to the 15-ml Falcon tube. Cells were centrifuged at 200 × g for 10 min at 4°C, and the medium was carefully removed. Cells were then washed twice with 10 ml of cold 1× PBS by centrifugation at 200 × g for 10 min at 4°C. The supernatant was removed following the final wash, and the pellet was resuspended in 1 ml of cold 1× PBS. The crude total protein fraction was collected by transferring 100 μl from the cell suspension to a 1.5-ml tube. The cells were pelleted by centrifugation at 200 × g for 7 min at 4°C. The supernatant was carefully removed, and the cells resuspended in 38 μl of whole-cell extraction buffer (50 mM Tris [pH 8], 280 mM NaCl, 10% glycerol, 0.5% IGEPAL, 5 mM MgCl2, RNase A, DNase I, 10 μg/ml ethidium bromide, protease inhibitor cocktail, and 5 μg/ml PMSF) for 1 h on ice. The samples were centrifuged for 1 h at 20,000 × g at 4°C. The supernatant was then transferred to a 1.5-ml tube and saved as the total protein fraction. The remaining 900 μl of the cell pellet was transferred to a 1.5-ml tube to process the cytosolic and nuclear fractions. The cells were centrifuged at 800 × g for 7 min at 4°C. The supernatant was carefully removed and the pellet was resuspended in 315 μl of lysis buffer (10 mM Tris [pH 6.8], 1 mM DTT, 1 mM MgCl2, 10% sucrose, 100 mM NaCl, 0.5% IGEPAL, protease inhibitor cocktail, and 5 μg/ml PMSF) and incubated on ice for 5 min. Following the incubation, the samples were centrifuged at 700 × g for 2 min at 4°C. The supernatant containing the cytosolic fraction was then carefully transferred to a fresh 1.5-ml tube. The pellet containing the crude nuclear fraction was washed twice in wash buffer (10 mM Tris [pH 6.8], 1 mM DTT, 1 mM MgCl2, 10% sucrose, 100 mM NaCl, protease inhibitor cocktail, and 5 μg/ml PMSF) by inverting the tube 3 times. The supernatant was removed each time by centrifugation at 700 × g for 2 min at 4°C. The washed crude pellet was then resuspended in 315 μl of extraction buffer (10 mM Tris [pH 6.8], 1 mM DTT, 1 mM MgCl2, 10% sucrose, 400 mM NaCl, protease inhibitor cocktail, and 5 μg/ml PMSF) and incubated on ice for 10 min. The nuclear fraction was collected by centrifuging the sample at 5,200 × g for 2 min at 4°C. The nuclear fraction was carefully removed and transferred to a fresh microcentrifuge tube. All samples were immediately used for protein analysis or stored at −80°C. The bicinchoninic acid method was used to determine protein concentrations. To probe TREX1 localization, equivalent amounts of protein were used for immunoblotting as previously described. Fraction integrity was determined by probing the nuclear and cytoplasmic fractions for lamin and GAPDH, respectively.
The HIV-1 intasomes used in our study were prepared by using published methods (56, 86, 87). Briefly, a double-stranded preprocessed viral DNA substrate was used as the donor DNA (Integrated DNA Technologies, Coralville, IA, USA). A 25-base oligonucleotide (5′-AGCGTGGGCGGGAAAATCTCTAGCA-3′) was synthesized with a complementary 27-base oligonucleotide (5′-ACTGCTAGAGATTTTCCCGCCCACGCT-3′) to provide the 3′ OH recessed end for the preprocessed CSC intasomes. Similarly, unprocessed SSC intasome donor DNA was formed from a 27-base oligonucleotide (5′-AGCGTGGGCGGGAAAATCTCTAGCAGT-3′) to complement the 27-base oligonucleotide. The recombinant Sso7d-integrase (Sso7d-IN) was expressed and purified in Escherichia coli BL21(DE3), and the cells were lysed in buffer containing 20 mM HEPES (pH 7.5), 10% glycerol, 2 mM β-mercaptoethanol (β-ME), 20 mM imidazole, and 1 M NaCl. The protein was purified by nickel affinity chromatography and subsequently by gel filtration on a Hiload 26/60 Superdex-200 column (GE Healthcare, Milwaukee, WI, USA) equilibrated with 20 mM HEPES (pH 7.5), 10% glycerol, 2 mM DTT, and 500 mM NaCl. The protein was concentrated using an Amicon centrifugal concentrator (EMD Millipore, Burlington, MA, USA) and then used for intasome assembly or flash frozen in liquid nitrogen and stored at −80°C. Intasomes were assembled by incubating 3 μM HIV-1 integrase and 1 μM 3′-processed or unprocessed viral DNA substrate in 20 mM HEPES (pH 7.5), 20% glycerol, 5 mM 2-mercaptoethanol, 5 mM CaCl2, 2 μM ZnCl2, 100 mM NaCl, and 50 mM 3-(benzyldimethylammonio)propanesulfonate (NDSB-256) at 30°C for 1 h. Assembled intasomes were purified as described previously (56, 86, 87), aliquoted, flash frozen in liquid nitrogen, and stored at −80°C.
Intasome assays were performed with 25 nM intasomes in TREX1 buffer (20 mM Tris-HCl [pH 7.5], 5 mM MgCl2, 10 mM DTT, and 100 μg/μl BSA), which was amenable for both intasome and TREX1 activity, in a 20-μl reaction volume. Purified TREX1 protein was added to the reaction mixture to final concentrations of 5, 10, or 25 nM where indicated. In vitro integration reactions with the intasomes were carried out with 300 ng of circular pGEM plasmid DNA and incubated at 37°C for 1 h. The reaction was terminated with 0.5% SDS and 8 mM EDTA together with 0.5 mg/ml protease K and deproteinized for 1 h at 55°C. The DNA was recovered by ethanol precipitation, as described above, and resuspended in the original reaction volume of nuclease-free water. The recovered DNA was used as the template DNA for a SYBR green-based quantitative PCR to amplify the junctions between viral DNA substrate and the target DNA. The qPCR was carried out in a 20-μl volume, containing 1 to 2 μl (5 to 10 ng) of purified DNA product from the INS integration assay, 300 nM concentration of each primer targeting the viral DNA substrate (5′-AGCGTGGGCGGGAAAATCTC-3′) and the pGEM DNA (5′-GTCACGACGTTGTAAAACGACG-3′), and 1× iTaq universal SYBR green supermix (Bio-Rad Laboratories, Hercules, CA, USA). The qPCR cycling conditions for quantifying INS activity included an initial incubation at 95°C for 3 min, followed by 39 cycles of amplification and acquisition at 94°C for 15 s, 55°C for 30 s, and 72°C for 30 s. The thermal profile for melting curve analysis was obtained by holding the reaction at 65°C for 30 s, followed by a linear ramp in temperature from 65 to 95°C at a ramp rate of 0.5°C/s and acquisition at 0.5°C intervals. The Maestro CFX program was used for data analysis (Bio-Rad Laboratories).
Exonuclease assays were carried out with either 18-mer or 20-mer fluorescein-labeled oligonucleotides annealed to unlabeled complementary 20-mer oligonucleotides. Unlabeled and fluorescein-labeled oligonucleotides were commercially synthesized (Thermo Fisher, Waltham, MA, USA). Unlabeled phosphorothioate(*)-modified oligonucleotides were synthesized by Integrated DNA Technology (Coralville, IA, USA). The sequences of the oligonucleotides are as follows: 20-mer U3 (unprocessed), 5′-fluorescein-AGTGAATTAGCCCTTCCAGT-3′; 18-mer U3 (processed), 5′-fluorescein-AGTGAATTAGCCCTTCCA-3′; 20-mer U3 (complement), 5′-ACTGGAAGGGCTAATTCACT-3′; 20-mer phosphorothioate U3 (complement), 5′-ACTGGAAGGGCTAATTCAC*T-3′; 20-mer U5 (unprocessed), 5′-fluorescein-TGTGGAAAATCTCTAGCAGT-3′; 18-mer U5 (processed), 5′-fluorescein-TGTGGAAAATCTCTAGCA-3′; 20-mer U5 (complement), 5′-ACTGCTAGAGATTTTCCACA-3′; 20-mer phosphorothioate U5 (complement), 5′-ACTGCTAGAGATTTTCCAC*A-3′; 20-mer random (unprocessed), 5′-fluorescein-TGTGTGGATAGCTCACGTGA-3′; 18-mer random (processed), 5′-fluorescein-TGTGTGGATAGCTCACGT-3′; 20-mer random (complement), 5′-TCACGTGAGCTATCCACACA-3′; and 20-mer phosphorothioate random (complement), 5′-TCACGTGAGCTATCCACAC*A-3′.
The double-stranded DNA substrates containing unprocessed U3 or U5 or random sequences were generated by annealing the appropriate fluorescein-labeled 20-mer oligonucleotide to the corresponding 20-mer unlabeled or 20-mer unlabeled and phosphorothioate-modified complementary oligonucleotide. The double-stranded DNA substrates containing processed U3 or U5 or random sequences were generated by annealing the appropriate fluorescein-labeled 18-mer oligonucleotide to the corresponding 20-mer unlabeled or 20-mer unlabeled and phosphorothioate-modified complementary oligonucleotide. Double-stranded oligonucleotides were generated by diluting 10 μM fluorescein-labeled oligonucleotide and 20 μM unlabeled oligonucleotide in annealing buffer containing 10 mM Tris (pH 7.5) and 10 mM NaCl and then heating in a 95°C dry bath for 5 min. The heating block was removed from the dry bath and allowed to slowly cool to room temperature, and a sample aliquot was resolved on a 20% native PAGE gel to verify effective annealing of the oligonucleotides.
Exonuclease assays were assembled in a 50-μl final volume and contained 20 mM Tris-HCl (pH 7.5), 5 mM MgCl2, 2 mM DTT, 100 μg/ml bovine serum albumin, double-stranded DNA substrates (250 nM to 1 μM), and TREX1 protein (200 pM to 500 nM). The reaction mixtures were assembled on ice and then transferred to a 37°C dry bath for a designated time. Reactions were stopped by the addition of TREX1 assay dye (95% formamide and 0.05% xylene cyanol) followed by heating at 95°C for 1 min. Exonuclease reaction products were resolved on a 20% 19:1 acrylamide/bis-acrylamide 7 M urea PAGE gel. The gel-resolved reaction products were visualized using a PharosFX plus molecular imager system (Bio-Rad Laboratories, Hercules, CA, USA), and the band intensities were calculated using Quantity One 1-D analysis software (Bio-Rad Laboratories).
Data were expressed as means ± SEMs obtained from three independent experiments. Significance of differences between control and treated samples was determined by Student’s t test. P values of
This work was supported by the National Institutes of Health grants R01 AI136740, R01 DA 042348, R56 AI122960, R24 DA036420, and U54 MD007586 to C.D. and a Research Centers in Minority Institutions (RCMI) grant U54MD007586 to J.P. This work is also supported in part by the Meharry Translational Research Center (MeTRC) grant U54MD007593 and Tennessee CFAR grant P30 AI110527 from the National Institutes of Health to C.D. M.L. and R.C. are supported by the intramural research program of the National Institutes of Health.
We declare that we have no conflict of interest with the content of this article.
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