Regulated transgene expression may reduce transgene-specific and genotoxic risks associated with gene therapy. To prove this concept, we have investigated the suitability of doxycycline (Dox)-inducible human cytidine deaminase (hCDD) overexpression from lentiviral vectors to mediate effective myeloprotection while circumventing the lymphotoxicity observed with constitutive CDD activity. Rapid Dox-mediated transgene induction associated with a 6–17-fold increase in drug resistance was observed in 32D and primary murine bone marrow (BM) cells. Moreover, robust Dox-regulated transgene expression in the entire haematopoietic system was demonstrated for primary and secondary recipients of hCDD-transduced R26-M2rtTA transgenic BM cells. Furthermore, mice were significantly protected from myelosuppressive chemotherapy as evidenced by accelerated recovery of granulocytes (1.9±0.6 vs 1.3±0.3, P=0.034) and platelets (883±194 vs 584±160 103 per μl, P=0.011). Minimal transgene expression in the non-induced state and no overt cellular toxicities including lymphotoxicity were detected. Thus, using a relevant murine transplant model our data provide conclusive evidence that drug-resistance transgenes can be expressed in a regulated fashion in the lymphohaematopoietic system, and that Dox-inducible systems may be used to reduce myelotoxic side effect of anticancer chemotherapy or to avoid side effects of high constitutive transgene expression.
While haematopoietic gene therapy has been shown to represent a promising and successful therapeutic option for a number of congenital diseases and in particular immunodeficiency disorders,1, 2, 3, 4 these studies also have highlighted the problems still associated with this approach. Insertional mutagenesis leading to the clinical manifestation of leukaemias represents the most noticeable and serious side effect,4, 5 but also specific transgene-induced toxicities6, 7, 8 or transgene-directed immune reactions can pose serious risks. To circumvent or at least reduce these problems, the use of regulated gene transfer systems, allowing for temporary-controlled transgene expression has been suggested and this strategy appears particularly feasible for gene therapy approaches in which expression of the therapeutic transgene only is required for defined time periods.
A number of regulated gene expression systems incorporating different promoter-regulatory elements have been explored. Although some of these systems exploit promoters responsive to specific chemical or physical stimuli, such as metal iones,9 temperature10 or O2 saturation,11 regulatable systems based on the application of drugs, such rapamycin12 or the steroid hormones mifepristone or tamoxifen,13, 14 currently offer powerful alternatives. In particular, tetracycline-regulated systems (Tet-systems), as introduced by Gossen and Bujard15 in 1992, offer a reliable and robust approach to regulated transgene expression and currently represent the most frequently used inducible gene expression system. The Tet-system is dependent on two genetic elements: (i) the Escherichia coli Tet repressor protein (TetR) that exerts its transcriptional repression by binding to the Tet operator (TetO) and (ii) the presence of tetracycline or its potent analogue doxycycline (Dox) induces a conformational change within TetR and thereby abolishes the interaction with TetO and releases the transcriptional blockade.15, 16 For eukaryotes, a fusion protein of the activation domain of the herpes simplex virus VP16 protein and the TetR, called transcriptional activator (tTA), has been introduced that induces gene expression by binding to a promoter element (Ptet) placed downstream of TetO and this interaction is disrupted in the presence of Dox (Tet-OFF system).17 However, for in vivo usage, OFF-systems suffer from two major drawbacks: continuous presence of the activator is required in the quiescent state and the induction kinetic is mainly determined by the rate of drug clearance. Therefore, a modification of four amino acids within the activator was introduced, resulting in a reverse tTA (rtTA) that only binds to Ptet in the presence of Dox (Tet-ON system),16 and subsequently a number of modifications have been introduced into the rtTA and Ptet elements to reduce the background levels of transgene expression in the Tet-ON system.18, 19 One area that appears particularly suited for the use of inducible gene expression systems is the transfer of chemotherapy-resistance (CTX-R) genes into haematopoietic stem cells (HSCs) to protect the haematopoietic system from the toxicity of defined anti-neoplastic agents, as for this application transgene expression only is required for the relatively short periods of cytotoxic drugs administration. A number of such CTX-R genes have been identified and for several of them protection of haematopoiesis from the associated cytotoxic agents has been established in murine as well as large animal models.20, 21 This includes mutant forms of dihydrofolate reductase (mutDHFR),22, 23, 24 the multidrug-resistance type 1 (MDR1) gene coding for the cellular efflux pump p-glycoprotein25, 26, 27 or the gene coding for the DNA repair protein O6-methylguanine methyltransferase.28, 29, 30, 31 Cytidine deaminase represents another interesting CTX-R gene that protects lymphohaematopoietic cells from the cytotoxic deoxycytidine analogues cytosine-arabinoside (1-β-D-arabinofuranosylcytosine, Ara-C), gemcitabine (2′,2′-difluorodeoxycytidine) and 5-azacitabine (5-aza-2′deoxycytidine). In particular, Ara-C has been utilized clinically for extended time periods and has established itself as the most effective single agent in the treatment of acute leukaemias. Importantly, a protective effect of human cytidine deaminase (hCDD) gene transfer in the context of Ara-C and gemcitabine application has been established in murine and human clonogenic progenitor cells32, 33, 34 as well as in a murine in vivo bone marrow transplant model.35 However, these studies also revealed considerable lymphotoxicity induced by high constitutive hCDD expression in vivo.35
Thus, in a proof-of-principle study, we have investigated the suitability of Dox-regulated vector constructs to avoid transgene-specific side effects of gene therapy such as the lymphotoxicity associated with hCDD overexpression in the haematopoietic system. We here describe rapid and robust transgene expression upon Dox application with moderate background levels in the Dox-OFF state. This allowed for protection of the haematopoietic system from Ara-C-induced toxicity and at least at the expression level obtained here with the inducible system completely abrogated the lymphotoxicity observed previously in the context of high constitutive hCDD expression.
Efficient transgene induction by Dox in 32D myeloid cells
To investigate inducible hCDD expression, we have generated Dox-regulated third generation self-inactivating (SIN) lentiviral vectors expressing the hCDD-cDNA in combination with an enhanced green fluorescence protein (eGFP)-reporter gene (SIN.Tet.CDD; Figure 1a) and a control vector expressing only eGFP (SIN.Tet.GFP; Figure 1b). For both vectors, transgenes are expressed in the presence of Dox (Tet-ON system), and the SIN configuration increases the safety of the vectors and reduces the risk of vector mobilization.36 To investigate regulated expression of hCDD and eGFP from these constructs, myeloid 32D cells were transduced with either SIN.Tet.CDD or SIN.Tet.GFP in combination with SIN.PGK.rtTA3 (Figure 1c). Exposure to 2 μg ml−1 of Dox revealed a transduction efficiency of 30–50% for co-transduced eGFP+ cells and purity of the eGFP+ population subsequently was increased to >97% by fluorescence-activated cell sorting (data not shown).
The dose-response correlation of Dox-induced transgene expression in SIN.Tet.CDD-transduced 32D cells was investigated by exposing these cells for various time-intervals to Dox concentration ranging from 0 to 2.0 μg ml−1. As depicted in Figure 2a, administration of 0.2–2.0 μg ml−1 Dox induced eGFP expression to plateau levels within 24 h (for 2 μg ml−1; see also Supplementary Video 1). Rapid induction of eGFP also was observed at lower Dox concentrations, although exposure to 0.008 or 0.04 μg ml−1 led to notably reduced mean fluorescence intensities plateau levels. Mean fluorescence intensity levels measured in the presence of 0.002 and 0 μg ml−1 Dox were indistinguishable, albeit significantly higher than those for non-transduced cells. A similar dependency on Dox dosage, but a clearly delayed induction kinetic, was observed for hCDD expression (Figure 2b). In addition, optimal hCDD expression was observed for 0.2–2.0 μg ml−1 Dox; however, in contrast to eGFP, maximal hCDD expression was observed until days 4–5. Also for hCDD, low background expression in the absence of Dox was detectable, as demonstrated by a faint protein band on day 5.
Inducible CDD expression mediates profound protection against Ara-C in 32D cells
Functionality of Dox-inducible hCDD expression was evaluated by exposing SIN.Tet.CDD- or SIN.Tet.GFP-transduced 32D cells to Ara-C after 1 week of preculture in the presence or absence of Dox. In these experiments, SIN.Tet.CDD-transduced cells cultured in the presence of 2.0 μg ml−1 Dox proved completely resistant to Ara-C concentrations of up to 5000 nM, whereas non-Dox treated control cells or SIN.Tet.GFP-transduced cells were susceptible to Ara-C exposure from 50 nM onwards (Figure 2c). This correlates to a more than 17-fold increased LD50 concentration (from 300 to 350 to >5000 nM) even in cells transduced at low MOI and carrying only one copy of the SIN.Tet.CDD provirus. Moreover, as depicted in Figure 2d, hCDD-mediated drug resistance was dependent on the Dox dosage. When SIN.Tet.CDD-transduced 32D cells were exposed to Dox for 72 h, maximal protection from Ara-C was shown for Dox concentration of 0.2 μg ml−1 (LD50 >2.000 nM Ara-C), and this dose-response curve recapitulated the hCDD induction observed at the protein level. Of note, background hCDD expression in the absence of Dox did not induce relevant protection against Ara-C as LD50 values were unaffected by administration of 0–0.08 μg ml−1 of Dox.
Rapid downregulation of transgene activity upon discontinuation of Dox exposure
Cessation of Dox treatment resulted in rapid downregulation of transgene expression as evidenced by an effective shut-down of eGFP expression in SIN.Tet.GFP-transduced 32D cells within 48–72 h (Figure 2e; Supplementary Video 1). In addition, changes in hCDD protein expression were slower and markedly reduced levels of hCDD were not detected before 72 h of Dox withdrawal (Figure 2f). Of note, again moderate levels of hCDD protein were detectable in SIN.Tet.CDD-transduced control cells in the absence of Dox.
Efficient Dox-induced transgene expression in SIN.Tet.CDD-transduced primary haematopoietic cells of R26-M2rtTA mice
Next, we evaluated our vectors in primary haematopoietic cells. To this end, lineage-negative (Lin−) BM cells from B6.Cg-Gt(ROSA)26Sortm1(rtTA*M2)Jae/J (R26-M2rtTA) mice, constitutively expressing a rtTA protein (M2rtTA), were transduced with SIN.Tet.CDD or SIN.Tet.GFP. Similar to 32D cells, a dose-dependent increase in transgenic eGFP expression was measured in SIN.Tet.GFP-transduced cells, reaching a plateau within 24 h for Dox concentrations beyond 0.2 μg ml−1 (Figure 3a). Low background activity was detectable in cells cultured in the absence of Dox.
Expression of hCDD in Lin− cells was evaluated by western blot analysis of SIN.Tet.CDD- and SIN.Tet.GFP-transduced cells, which demonstrated robust hCDD induction within 96 h of exposure to 2 μg ml−1 Dox (Figure 3b). Furthermore, functionality of hCDD expression was demonstrated by marked drug resistance of clonogenic progenitor cell-derived colonies in the presence of Dox. Survival of SIN.Tet.CDD-transduced colonies in the presence of 300 and 600 nM Ara-C was >50% and >30%, respectively, whereas colony-forming capacity of SIN.Tet.GFP-transduced control cells was markedly reduced for all Ara-C concentrations exceeding 30 nM (Figure 3c). LD50 doses were increased ∼6-fold, from 50 to 300 nM Ara-C.
Dox-regulated transgene expression in peripheral blood T, B and myeloid cells in murine bone marrow (BM) chimeras
Next, we investigated the potential of Dox-induced hCDD expression in a murine bone marrow transplant model employing Lin− BM cells from R26-M2rtTA mice transduced with SIN.Tet.CDD or SIN.Tet.GFP as donor material. Primary transduction efficacy in these studies ranged from 30 to 50% and was documented by flow cytometric analysis of eGFP expression in an aliquot of donor cells exposed to Dox ex vivo. Figures 4a and b show induction of eGFP expression in peripheral blood B, CD4+ and CD8+ T, and myeloid cells upon the administration of Dox by drinking water. In all, haematopoietic cell compartments induction of eGFP was detected within 2 days and plateau levels were reached within 13 days. Thereafter, mean fluorescence intensity values remained constant until Dox administration was discontinued. Upon Dox, withdrawal eGFP mean fluorescence intensity values for B, CD4+ and CD8+ T, as well as myeloid cells, decreased substantially the first 20 days and reached almost background levels after 45 days (Figure 4c).
Dox-induced hCDD expression protects the haematopoietic system from Ara-C toxicity in vivo
To investigate the protective effect conveyed to SIN.Tet.CDD-transduced lymphohaematopoietic cells and their progeny by Dox application, mice receiving Dox for at least 1 month were treated with Ara-C utilizing, a previously described application schedule (500 mg kg−1, 1 × daily intraperitoneally, d1–4).35 At the time of treatment, mice had received Dox for at least 1 month. Of note, eGFP transgene expression rates at the time of treatment only were 15–25% for the individual cell compartments. Peripheral blood analysis was performed 3 days in advance and on day 2, 4 and 7 after the last Ara-C treatment. As shown in Table 1, Dox-induced expression of hCDD led to significantly improved recovery of granulocytes on day 4 (0.6±0.1 versus 0.4±0.1 × 103 per μl; P<0.05) and day 7 (1.9±0.6 versus 1.3±0.3 × 103 per μl; P<0.05) and platelets on day 7 (883±194 versus 584±160 × 103 per μl; P<0.05) post-treatment when compared with control animals transplanted with SIN.Tet.GFP-transduced cells. Accelerated recovery in the SIN.Tet.CDD group was also observed for lymphocytes (day 7 post-treatment), but here differences failed to reach statistical significance (P=0.08). As with constitutive hCDD expression,35 no long-term enrichment of vector-transduced cells was observed, when the relative contribution of gene-modified cells to peripheral blood cell compartments before and after Ara-C application was compared (Supplementary Table 1). These results were recapitulated in a second independent experiment (data not shown).
Dox-inducible eGFP and hCDD transgene expression in BM, spleen and thymus
After 24–28 weeks, primary recipients were killed. Vector copy numbers were determined from BM cells and revealed 3.5±0.8 vector copies in the SIN.Tet.CDD (n=6) versus 4.0±3.4 copies in the Sin.Tet.GFP (n=6) group. Dox-mediated gene expression was investigated in BM, spleen and thymus of animals transplanted with SIN.Tet.CDD-transduced cells. At this time point animals had received Dox for a minimum of 8 weeks. As evident from Figure 5a, efficient expression of the eGFP transgene was observed in all BM-derived stem- and progenitor compartments. EGFP expression also was detected in BM-derived mature and immature B cells, all thymic T-cell fractions ranging from primitive double-negative via double-positive to more differentiated single-positive CD4+ or CD8+ T cells (Figure 5b) as well as splenic mature B and T cells (Figure 5c). Furthermore, eGFP expression was observed in spleen-derived myeloid cells.
Expression of the hCDD transgene in haematopoietic organs of SIN.Tet.CDD animals was evaluated by western blot analysis performed on total organ cell lysates. To compensate for differences in loaded amounts of protein, densitometric analysis assessing hCDD protein levels relative to the levels of vinculin was utilized. Although hCDD expression was undetectable in haematopoietic organs of wild-type animals, these studies clearly demonstrated hCDD expression in the BM, spleen and thymus of mice transplanted with SIN.Tet.CDD-transduced cells (Figure 5d).
Not surprisingly, effective hCDD expression also was observed on a functional level when Ara-C resistance of progenitor-derived colonies was assessed in clonogenic assays. When total BM cells from two mice transplanted with SIN.Tet.CDD-transduced cells and wild-type C57BL/6 mice were compared, pronounced differences were noted. In particular, virtually all colonies derived from control C57BL/6 mice died at 400 nM Ara-C (∼1% surviving colonies), whereas, 32 and 50% of colonies from SIN.Tet.CDD animals survived this treatment (Figure 5e).
Efficient transgene expression upon Dox exposure in secondary recipients
The long-term reconstitution potential of SIN.Tet.CDD- or SIN.Tet.GFP- transduced HSCs was further documented by secondary transplantation. Robust transgenic eGFP expression was observed in peripheral blood B, CD4+ and CD8+ T, as well as myeloid cells of secondary recipients when Dox was administered by drinking water 6 weeks after the secondary transplantation (Supplementary Figure 1).
No evidence of cellular toxicity including lymphotoxicity following Dox-regulated hCDD expression
A major aspect of our study was the prevention of CDD-mediated lymphotoxicity observed previously with constitutive hCDD (over)expression.35 In these studies, lymphotoxicity manifested primarily as a significantly reduced contribution of gene modified B and T cells to lymphopoiesis. Therefore, we have systematically investigated the contribution of gene-modified (that is, eGFP+) cells to the various lymphoid cell compartments comparing animals that received either SIN.Tet.CDD- or SIN.Tet.GFP-transduced transplants. Even after Dox administration for up to 8 weeks, no alterations in peripheral blood cell counts including the lymphocyte count were observed (also see Table 1, day −3 data). In addition, the relative contribution of gene-modified cells to peripheral blood B, CD4+ or CD8+ T and myeloid cells remained fairly constant during the 8 weeks of Dox administration (Figures 6a and b) and, even more important, similar contribution of transduced cells were observed for the myeloid and the lymphoid cell compartment at least at the CDD expression levels achieved with the Dox-inducible system.
The study presented here demonstrates efficient Dox-regulated transgene expression in myeloid 32D and primary BM cells in vitro as well as in a murine bone marrow transplant model. In the transplant model, Dox-induced CDD expression not only significantly reduced Ara-C-induced myelotoxicity but at least at the expression level obtained with the Dox inducible system also abrogated the lymphotoxicity observed previously with constitutive hCDD expression.35 Clearly, high transgene induction in the presence and tight regulation in the absence of Dox, that is, an adequate therapeutic window, constitutes a major prerequisite for the successful use of Dox-regulated vector systems. To this point we not only have shown robust and rapid transgene expression upon Dox application but also moderate background levels in the Dox-OFF state. Importantly, the minor hCDD background levels observed in the OFF state had no unwanted side effect in vitro or in vivo, and in the in vitro situation not even correlated to an increased protection capacity. Furthermore, effective shut-down of eGFP as well as hCDD transgene expression was observed upon Dox cessation. The prolonged shut-off kinetic for hCDD most likely reflects differences in protein half life, whereas the more rapid detection of GFP upon Dox administration easily can be explained by the higher sensitivity of flow-cytometric versus western blot analysis.
In our study, lack of cell type-specific toxicity and particularly lymphotoxicity was indicated by normal cell counts and, more importantly, similar contribution of gene-modified cells to all lymphohaematopoietic cell compartments. This is important as in previous studies, using conventional LTR-driven γ-retroviral vectors CDD-induced lymphotocity was identified by a significantly reduced relative contribution of gene marked cells to lymphoid as compared to myeloid compartments.35 A potential biochemical basis for this observation has been identified recently, when selective toxicity within the lymphoid compartment was observed in mice lacking the deoxycytidine kinase gene.37 In the deoxyribonucleotide salvage pathway, deoxycytidine kinase catalyses the phosphorylation of deoxycytidine to the monophosphate state and thereby competes with cytidine deaminase for this substrate. Thus, lack of deoxycytidine kinase as well as overexpression of cytidine deaminase both can be expected to diminish production of deoxycytidine triphosphate, the final product of the nucleotide salvage pathway. Although the deoxyribonucleotide salvage pathway is dispensable for most developmental processes, including embryo- and organogenesis, it appears to have a critical role during early T- and B-cell development when T-cell receptor- or VDJ-recombination is followed by massive cellular proliferation.37 Nevertheless, the evidence gathered in the present study does not necessarily establish a causal relationship between the lack of lymphotoxicity and the Dox-regulation of CDD expression, in particular as no control group expressing hCDD constitutively in the haematopoietic system was employed. Thus, other explanations and foremost lower levels of CDD expression from the Dox-induced pTet as compared to the SFFV-LTR promoter have to be considered. At least in 32D cells, indeed lower CDD expression in SIN.Tet.CDD- versus RSF91.CDD- transduced cells have been observed (own unpublished date). Thus, the CDD levels achieved by our Dox-regulated system, while allowing for myeloprotection, may be too low to cause lymphoid toxicity. This notion is supported by the fact that in the present study even prolonged Dox application for up to 90 days was not associated with lymphotoxic side effects.
No enrichment of hCDD-positive cells was observed in our model following Ara-C application. This is in line with previous studies also failing to show consistent enrichment of hCDD-positive cells by short-term high-dose Ara-C application in an in vivo model35 and probably reflects the relative quiescence of primitive stem cell populations in steady-state haematopoiesis and the S-phase-specific cytotoxic activity of Ara-C.38, 39 Indeed, we recently demonstrated that toxicity of short-term high-dose Ara-C application primarily effect haematopoietic precursor and late progenitor populations, while stem cells or long-lived progenitors remained relatively unaffected.40 Ara-C toxicity for primitive haematopopietic cells can be augmented, however, by prolonged low-dose Ara-C application (for example, 30–60 mg kg−1 daily for 10–20 days). In this setting at least transient enrichment of CDD-transduced haematopoietic cells has been achieved.40
Dox-regulated transgene expression also offers a way to reduce the genotoxic side effects of integrating vector-based gene therapy approaches and particularly the risk of insertional mutagenesis. As the activation of neighbouring oncogenes by vector integrations clearly is linked to vector-associated enhancer activity,41 the temporary nature of the transgene expression from inducible system should substantially reduce this risk. In addition, state-of-the-art third generation SIN lentiviral constructs were employed in our study. In comparison to conventional LTR-driven γ-retroviral vectors, which preferentially integrate into gene-regulatory regions,42, 43 these constructs offer improved safety features as lentiviral vectors not only are less likely to integrate in the vicinity of promoter elements and CpG islands but also allow for reduced ex vivo cell manipulation times. Moreover, the SIN configuration permits the usage of alternative regulatory sequences, such as physiological endogenous promoters, that further reduce the genotoxic risk by limiting the range of influence of enhancer activity or miRNA targets to increase cell-type specificity.41, 44, 45
Clearly, immunogenicity of the continuously expressed transactivator protein represents a concern for a potential clinical application of Dox-regulated vector system, and studies in non-human primates indicate that in certain situations, the Tet-activator may elicit a cellular and humoral immune response.46, 47 Given the therapeutic efficacy of Ara-C, particularly in haematopoietic malignancies, this setting also represents the most likely scenario for the clinical application of Dox-regulated CDD expression. This is probably less problematic in myeloablative and in particular allogeneic HSC-transplant settings, where de novo formation of the immune system including tolerance induction to novel antigens should prevent immune reactions to components of the Tet-system.48 Immunogenicity may become more problematic, however, with strategies aiming at the delivery of Tet-regulated transgenes to non or only partially myeloablated hosts. In this situation, immunomodulatory approaches based on regulatory T cells or tolerogenic dendritic cells may be required.49
Another hurdle for the clinical application of Dox-inducible transgene expression systems is the transfer of two expression cassettes to HSCs. This appears problematic not only in terms of efficacy but also with regard to safety considerations, as more viral integrations sites could translate into an increased risk of insertional mutagenesis. Currently, the most promising solution to this problem are all-in-one vector constructs, which allow for simultaneous expression of the TetR as well as an Tet-regulated transgene of interest from two different transcription units incorporated into the same vector construct.18, 50, 51
Thus, a number of issues still have to be addressed before the clinical applications of Dox-regulated gene expression systems in HSC gene therapy. However, once these problems are solved, this strategy holds considerable promise to reduce insertion-triggered genotoxic as well as transgene-specific side effects of gene therapy for approaches in which transgene expression is only required temporarily, such as the transgenic expression of drug-resistance genes in the haematopoietic system.
Materials and methods
C57BL6/J mice were obtained from Charles River Laboratories (Sulzfeld, Germany) and B6.Cg-Gt(ROSA)26Sortm1(rtTA*M2)Jae/J (R26-M2rtTA) transgenic mice from the central MHH animal facility (Hannover, Germany). All mice were kept in IVC racks (Allentown Inc., Mömbris, Germany) in pathogen-free conditions. All animal experiments were approved by the local animal welfare committee and performed according to their guidelines.
Lentiviral constructs were based on third generation SIN lentiviral vectors modified by insertion of a woodchuck hepatitis virus-derived post-transcriptional-regulatory element (wPRE).52 SIN.Tet.GFP (also called pRRL.PPT.pTet.eGFP.pre*) was constructed by cloning the Tet promoter15 as an XhoI/AgeI fragment (amplified by primers 5′ Tet xho 5′-IndexTermGCCTCGAGCTAGACGAGTTTACTCCCTATCAGTGATAGAGAACGATGTC-3′ and 3′ Tet age 5′-IndexTermCGACCGGTGCGATCTGACGGTTCACTAAACGAG-3′; restriction sites underlined) into pRRL.PPT.SF.GFP.pre*53 substituting for the SFFV U3 promoter. To generate SIN.Tet.CDD, the cDNA of hCDD (Open Biosystem, IHS1380-OB-97652440, Epsom, UK) was cloned via AgeI/SalI into the SIN.Tet.GFP backbone followed by an exchange of eGFP for IRES.eGFP using SalI restriction. For generation of SIN.PGK.rtTA3, the reverse transactivator rtTA3(ref. 17) was amplified by PCR from a pTRIPZ vector (Open Biosystems), using primers 5′ rtTA3 BamHi 5′-IndexTermGTGGATCCGCCACCATGTCTAGGCTGGACAAGAGCAAA-3′ and 3′ rtTA3 SalI 5′-IndexTermTTGTCGACTTACCCGGGGAGCATGTCAAGGT-3′ and cloned into pRRL.PPT.PGK.GFPpre52 (kindly provided by Luigi Naldini, Milano). All PCR products were verified by sequencing.
Production of viral supernatants and titration
Viral supernatants were generated by transfection of 293T cells as described previously.41, 54 In brief, 7 × 106 293T cells cultured in high-glucose Dulbecco’s modified Eagle’s Medium (PAA, Cölbe, Germany) supplemented with 10% fetal calf serum, 100 U ml−1 penicillin/streptomycin and 2 mmol l−1 glutamine (all PAA) were seeded 24 h before transfection in 10 cm dishes. Cells were transfected with 5 μg lentiviral vector, 8 μg pcDNA3.GP.4xCTE (expressing HIV-1 gag/pol), 5 μg pRSV-Rev and 2 μg pMD.G (encoding the VSV glycoprotein) using calcium phosphate precipitation in the presence of 20 mmol l−1 HEPES (PAA) and 25 μmol l−1 Chloroquine (Sigma-Aldrich, Munich, Germany). Viral supernatant was harvested 36 and 48 h post-transfection and concentrated using ultracentrifugation (Becton Dickinson, Heidelberg, Germany) for 3 h at 30 000 g and 4 °C. Titres were determined by transduction of transgenic murine SC1 fibroblasts, expressing either the rtTA protein rtTA3 (kindly provided by Olga Kustikova, MHH Hannover, Hannover, Germany) or the RRL.PPT.pTet.eGFP.pre* vector with serial dilution of viral supernatant in the presence of 2 μg ml−1 Doxycycline-Hyclate (Sigma-Aldrich). Percentage of transduced cells was determined 5 days post-transduction by flow cytometric assessment of eGFP-expressing cells.
In vitro experiments
Murine 32D cells were cultured in RPMI-1640 supplemented with 10% fetal calf serum, 100 U ml−1 penicillin/streptomycin (Pen/Strep), 2 mmol l−1 glutamine (all PAA) and 2 ng ml−1 rmIL-3 (Peprotech, Hamburg, Germany). Cells were co-transduced with VSVg pseudotyped lentiviral particles (SIN.Tet.GFP/SIN.PGK.rtTA3 or SIN.Tet.CDD/SIN.PGK.rtTA3) in the presence of 10 μg ml−1 protaminsulfate (Roth, Karlsruhe, Germany) at 37 °C. Twenty-four hours after transduction, cells were exposed to 2 μg ml−1 of Dox for 48 h and subsequently sorted for eGFP expression (FACS AriaIIu, Becton Dickinson) to establish transgenic 32D cells of purity >97%. Following Dox-starvation for 2–4 weeks, transgenic 32D cells were stimulated with different Dox concentrations and transgene expression was analysed by flow cytometry using FACSCalibur (Becton Dickinson) or by western blot analysis at different time points. Dox withdrawal studies were performed using transgenic 32D cells stimulated with Dox for 2–4 weeks. Transgene expression was analysed by flow cytometry using FACSCalibur or by western blot analysis. For in vitro protection as well as LD50 analysis, transgenic 32D cells were cultured for 3 days in the presence of different concentrations of Ara-C (Alexan, Neocorp AG, Weilheim, Germany) and either 2 μg ml−1 or different concentrations of Dox, respectively. Cell survival was assessed by flow cytometry analysis using propidium iodide staining (Sigma Aldrich).
Lineage negative BM cells
BM cells were isolated from femura and tibiae of 12–24 week-old B6.Cg-Gt(ROSA)26Sortm1(rtTA*M2)Jae/J (R26-M2rtTA) transgenic mice. Lin− cells were purified using MACS separation (Lineage Cell depletion kit, Miltenyi, Bergisch Gladbach, Germany) and prestimuled for 24 h in StemSpan medium (StemCell Technologies, Cologne, Germany) supplemented with 10 ng ml−1 rmSCF, 20 ng ml−1 rmTPO, 20 ng ml−1 rmIGF and 10 ng ml−1 rhFGF (all PeproTech).55 Subsequently, cells were transduced once (MOI 20) with SIN.Tet.CDD or SIN.Tet.GFP on retronection (10 μg cm−2; Takara, Otsu, Japan) -coated dishes as recommended by the manufacturer. Subsequently, transduced cells were stimulated for 48 h with 2 μg ml−1 Dox, sorted for eGFP expression and subjected either to clonogenic progenitor assays or Dox kinetic studies. For Dox-ON kinetics, transduced Lin− cells were cultured in the presence of different concentrations of Dox and analysed using flow cytometry at different time points.
Clonogenic progenitor assays
Clonogenic progenitor cells were assessed by incubating 1.500 SIN.Tet.GFP or SIN.Tet.CDD transduced Lin− cells from R26-M2rtTA transgenic mice, previously sorted for eGFP expression. Alternatively, 5 × 105 total BM cells harvested from primary SIN.Tet.CDD-transplanted recipients were utilized. Clonogenic cultures were performed in 1 ml IMDM/1.3% methylcellulose supplemented with 15% fetal calf serum, 2% bovine serum albumin, 2 mM L-glutamine, 50 μM 2-Mercaptoethanol, 10 μg ml−1 rh-insulin, 200 μg ml−1 human transferrin, 50 ng ml−1 rm-SCF, 10 ng ml−1 rm-IL3, 10 ng ml−1 rm-IL6 and 5 IU ml−1 rh-EPO (HSC007, R&D Systems, Wiesbaden-Nordenstadt, Germany) in the presence of 2 μg ml−1 Dox and different concentrations of Ara-C. Colonies of more than 50 cells were counted after 7 days.
Murine in vivo bone marrow transplant model
Donor cell isolation, transduction and transplantation
Lin− BM cells of 12–24 week-old R26-M2rtTA transgenic mice were isolated and transduced as described above. Forty-eight hours after transduction, cells were transplanted into tail veins of lethally (10 Gy) irradiated 10–12 weeks-old female C57BL6/J mice (Charles River Laboratories). Haematologic reconstitution was confirmed by peripheral blood counts 24–30 days after transplantation.
Peripheral blood sampling and analysis
For peripheral blood analysis, 10 μl (for peripheral blood counts) or 10–20 μl (for flow cytometric analysis of eGFP expression) of blood was taken from the retroorbital plexus. Blood counts were evaluated on veterinary blood cell counter (VetABC, Scil Animal Care, Viernheim, Germany). Peripheral blood eGFP+ cells were analysed by flow cytometry using LSRII (Becton Dickinson, antibodies given in Supplementary Table 2) following red cell lysis.
After haematologic reconstitution, animals received drinking water supplemented with 2 mg ml−1 Dox and 25% sucrose (Roth) for 4–12 weeks, to induce transgene expression and to perform in vivo protection analysis. Thereafter, Dox was withdrawn for 4–8 weeks before, mice were treated again with Dox by drinking water for at least 4 weeks before final analysis.
After 4 weeks of Dox administration, animals were treated intraperitoneally with 500 mg kg−1 Ara-C for 4 consecutive days. To investigate haematopoietic recovery, peripheral blood analysis was performed on day 2, 4 and 7 after the last Ara-C dose.
Analysis of haematopoietic subcompartments from BM, spleen and thymus
At 16–24 weeks after transplantation, animals were killed and eGFP+ cells from phenotypically defined cell subcompartments of BM, spleen and thymus were analysed in addition to peripheral blood, using flow cytometry. In brief, cells were isolated from different haematopoietic organs using cell strainer (Becton Dickinson), washed with PBS supplemented with 2 mM EDTA and 2% fetal calf serum followed by a 1-min red blood cell depletion step utilizing lysis buffer (0.83% NH4Cl, 0.5% KHCO3, 0.5 mM EDTA) and stained for 45 min with conjugated antibodies. All antibodies were used as recommended by the manufacturer. Data were analysed using LSR II (Becton Dickinson) and FlowJo software (TreeStar, Ashland, OR, USA)
A total of 20–30 × 106 BM cells from primary recipients were transplanted by tail vein into lethally irradiated (10 Gy) 10–12 weeks-old female C57BL6/J mice. Transgenic eGFP expression in peripheral blood cells was analysed 6–8 weeks days after transplantation and including 2 weeks of Dox treatment as described above.
Western blot analysis of hCDD expression
Protein samples from SIN.Tet.CDD- or SIN.Tet.GFP-transduced 32D and Lin− BM cells or BM, thymus and spleen cells of animals in the SIN.Tet.CDD group 24–32 weeks post-transplantation were prepared using RIPA-Buffer (Sigma-Aldrich) and protease inhibitor (Roche Diagnostics, Manheim, Germany). Subsequently, 10 μg of protein lysates were loaded on a 12% SDS gel electrophoresis. Protein samples from SIN.Tet.CDD-transplanted animals were prepared using RIPA buffer according to the manufacturer’s suggestion. Subsequently, blots were incubated over night at 4 °C with either rabbit polyclonal antibody anti-hCDD (Ab56053-100, AbCam, Cambridge, UK) according to manufacturer’s description or a mouse IgG monoclonal anti-vinculin antibody (Sigma Aldrich) 1:1000, respectively. Secondary staining was performed at room temperature for 1 h using peroxidase-conjugated donkey anti-rabbit or donkey anti-mouse antibodies (both Jackson ImmunoResearch, Newmarket, Suffolk, UK). To detect peroxidase activity, SuperSignal West Femto Substrate Kit (Thermo Fisher Scientific, Schwerte, Germany) and universal hood II (BioRad, Munich, Germany) were used.
Continuous live cell imaging of Dox-dependent eGFP reporter expression was performed on SIN.Tet.GFP-transduced 32D cells. For Dox-ON kinetic, transduced cells starved of Dox for 4 or more weeks were exposed to 2 μg ml−1 Dox on day 0. For Dox-OFF kinetic, transduced cells were cultured in the presence of 2 μg ml−1 of Dox for 4 weeks or more before Dox was withdrawn. To avoid floating of suspension cells during the imaging process, 5 × 104 cells were immobilized in non-adhesive triangular microcavities. Brightfield and fluorescence images were captured in 15 min intervals for at least 65 h for Dox-ON or 137 h for Dox-OFF using the AxioObserver Z1 fluorescence microscope (Zeiss, Jena, Germany) with a 37-°C humidity chamber and 5% CO2 levels. Video analysis was performed with Axiovision Software 4.70 (Zeiss).
Genomic DNA from transduced or non-transduced cells was isolated using GenElute Mammalian Genomic DNA Miniprep Kit (Sigma-Aldrich, Steinheim, Germany). Mean copy number was determined by quantitative PCR using primers detecting woodchuck post-transcriptional element (wPRE) and polypyrimidine tract binding protein 2 (PTBP2) as internal reference. Quantitative PCR was performed using FAST SYBR Green (Stratagene, Santa Clara, CA, USA) on a StepOnePlus (Applied Biosystems, Carlsbad, CA, USA). A plasmid standard containing the sequences for wPRE and PTBP2 was used for quantification.
Statistical analysis was performed using Prism 5 software (GraphPad, La Jolla, CA, USA). Unless otherwise noted Student‘s t-test (unpaired, one-tailed) was used.
Aiuti A, Slavin S, Aker M, Ficara F, Deola S, Mortellaro A et alCorrection of ADA-SCID by stem cell gene therapy combined with nonmyeloablative conditioning. Science 2002; 296: 2410–2413.
Boztug K, Schmidt M, Schwarzer A, Banerjee PP, Diez IA, Dewey RA et al. Stem-cell gene therapy for the Wiskott-Aldrich syndrome. N Engl J Med 363: 1918–1927.
Cavazzana-Calvo M, Hacein-Bey S, de Saint Basile G, Gross F, Yvon E, Nusbaum P et al. Gene therapy of human severe combined immunodeficiency (SCID)-X1 disease. Science 2000; 288: 669–672.
Ott MG, Schmidt M, Schwarzwaelder K, Stein S, Siler U, Koehl U et al. Correction of X-linked chronic granulomatous disease by gene therapy, augmented by insertional activation of MDS1-EVI1, PRDM16 or SETBP1. Nat Med 2006; 12: 401–409.
Hacein-Bey-Abina S, Garrigue A, Wang GP, Soulier J, Lim A, Morillon E et al. Insertional oncogenesis in 4 patients after retrovirus-mediated gene therapy of SCID-X1. J Clin Invest 2008; 118: 3132–3142.
Woods NB, Bottero V, Schmidt M, von Kalle C, Verma IM . Gene therapy: therapeutic gene causing lymphoma. Nature 2006; 440: 1123.
Wicke DC, Meyer J, Buesche G, Heckl D, Kreipe H, Li Z et al. Gene therapy of MPL deficiency: challenging balance between leukemia and pancytopenia. Mol Ther 2010; 18: 343–352.
Gentner B, Visigalli I, Hiramatsu H, Lechman E, Ungari S, Giustacchini A et al. Identification of hematopoietic stem cell-specific miRNAs enables gene therapy of globoid cell leukodystrophy. Sci Transl Med 2010; 2: 58ra84.
Mayo KE, Warren R, Palmiter RD . The mouse metallothionein-I gene is transcriptionally regulated by cadmium following transfection into human or mouse cells. Cell 1982; 29: 99–108.
Wurm FM, Gwinn KA, Kingston RE . Inducible overproduction of the mouse c-myc protein in mammalian cells. Proc Natl Acad Sci USA 1986; 83: 5414–5418.
Kenneth NS, Rocha S . Regulation of gene expression by hypoxia. Biochem J 2008; 414: 19–29.
Rivera VM, Clackson T, Natesan S, Pollock R, Amara JF, Keenan T et al. A humanized system for pharmacologic control of gene expression. Nat Med 1996; 2: 1028–1032.
Roscilli G, Rinaudo CD, Cimino M, Sporeno E, Lamartina S, Ciliberto G et al. Long-term and tight control of gene expression in mouse skeletal muscle by a new hybrid human transcription factor. Mol Ther 2002; 6: 653–663.
Vegeto E, Allan GF, Schrader WT, Tsai MJ, McDonnell DP, O'Malley BW . The mechanism of RU486 antagonism is dependent on the conformation of the carboxy-terminal tail of the human progesterone receptor. Cell 1992; 69: 703–713.
Gossen M, Bujard H . Tight control of gene expression in mammalian cells by tetracycline-responsive promoters. Proc Natl Acad Sci USA 1992; 89: 5547–5551.
Gossen M, Freundlieb S, Bender G, Muller G, Hillen W, Bujard H . Transcriptional activation by tetracyclines in mammalian cells. Science 1995; 268: 1766–1769.
Das AT, Zhou X, Vink M, Klaver B, Verhoef K, Marzio G et al. Viral evolution as a tool to improve the tetracycline-regulated gene expression system. J Biol Chem 2004; 279: 18776–18782.
Heinz N, Schambach A, Galla M, Maetzig T, Baum C, Loew R et al. Retroviral and transposon-based tet-regulated all-in-one vectors with reduced background expression and improved dynamic range. Hum Gene Ther 22: 166–176.
Urlinger S, Baron U, Thellmann M, Hasan MT, Bujard H, Hillen W . Exploring the sequence space for tetracycline-dependent transcriptional activators: novel mutations yield expanded range and sensitivity. Proc Natl Acad Sci USA 2000; 97: 7963–7968.
Trobridge GD, Kiem HP . Large animal models of hematopoietic stem cell gene therapy. Gene Therapy 2010; 17: 939–948.
Moritz T, Williams DA . Marrow protection—transduction of hematopoietic cells with drug resistance genes. Cytotherapy 2001; 3: 67–84.
Allay JA, Persons DA, Galipeau J, Riberdy JM, Ashmun RA, Blakley RL et al. In vivo selection of retrovirally transduced hematopoietic stem cells. Nat Med 1998; 4: 1136–1143.
Capiaux GM, Budak-Alpdogan T, Alpdogan O, Bornmann W, Takebe N, Banerjee D et al. Protection of hematopoietic stem cells from pemetrexed toxicity by retroviral gene transfer with a mutant dihydrofolate reductase-mutant thymidylate synthase fusion gene. Cancer Gene Ther 2004; 11: 767–773.
Meisel R, Bardenheuer W, Strehblow C, Sorg UR, Elmaagacli A, Seeber S et al. Efficient protection from methotrexate toxicity and selection of transduced human hematopoietic cells following gene transfer of dihydrofolate reductase mutants. Exp Hematol 2003; 31: 1215–1222.
Sorrentino BP, Brandt SJ, Bodine D, Gottesman M, Pastan I, Cline A et al. Selection of drug-resistant bone marrow cells in vivo after retroviral transfer of human MDR1. Science 1992; 257: 99–103.
Hildinger M, Schilz A, Eckert HG, Bohn W, Fehse B, Zander A et al. Bicistronic retroviral vectors for combining myeloprotection with cell-surface marking. Gene Therapy 1999; 6: 1222–1230.
Schiedlmeier B, Schilz AJ, Kuhlcke K, Laufs S, Baum C, Zeller WJ et al. Multidrug resistance 1 gene transfer can confer chemoprotection to human peripheral blood progenitor cells engrafted in immunodeficient mice. Hum Gene Ther 2002; 13: 233–242.
Jansen M, Sorg UR, Ragg S, Flasshove M, Seeber S, Williams DA et al. Hematoprotection and enrichment of transduced cells in vivo after gene transfer of MGMT(P140K) into hematopoietic stem cells. Cancer Gene Ther 2002; 9: 737–746.
Milsom MD, Jerabek-Willemsen M, Harris CE, Schambach A, Broun E, Bailey J et al. Reciprocal relationship between O6-methylguanine-DNA methyltransferase P140K expression level and chemoprotection of hematopoietic stem cells. Cancer Res 2008; 68: 6171–6180.
Neff T, Beard BC, Peterson LJ, Anandakumar P, Thompson J, Kiem HP . Polyclonal chemoprotection against temozolomide in a large-animal model of drug resistance gene therapy. Blood 2005; 105: 997–1002.
Zielske SP, Reese JS, Lingas KT, Donze JR, Gerson SL . In vivo selection of MGMT(P140K) lentivirus-transduced human NOD/SCID repopulating cells without pretransplant irradiation conditioning. J Clin Invest 2003; 112: 1561–1570.
Bardenheuer W, Lehmberg K, Rattmann I, Brueckner A, Schneider A, Sorg UR et al. Resistance to cytarabine and gemcitabine and in vitro selection of transduced cells after retroviral expression of cytidine deaminase in human hematopoietic progenitor cells. Leukemia 2005; 19: 2281–2288.
Momparler RL, Eliopoulos N, Bovenzi V, Letourneau S, Greenbaum M, Cournoyer D . Resistance to cytosine arabinoside by retrovirally mediated gene transfer of human cytidine deaminase into murine fibroblast and hematopoietic cells. Cancer Gene Ther 1996; 3: 331–338.
Neff T, Blau CA . Forced expression of cytidine deaminase confers resistance to cytosine arabinoside and gemcitabine. Exp Hematol 1996; 24: 1340–1346.
Rattmann I, Kleff V, Sorg UR, Bardenheuer W, Brueckner A, Hilger RA et al. Gene transfer of cytidine deaminase protects myelopoiesis from cytidine analogs in an in vivo murine transplant model. Blood 2006; 108: 2965–2971.
Zufferey R, Dull T, Mandel RJ, Bukovsky A, Quiroz D, Naldini L et al. Self-inactivating lentivirus vector for safe and efficient in vivo gene delivery. J Virol 1998; 72: 9873–9880.
Toy G, Austin WR, Liao HI, Cheng D, Singh A, Campbell DO et al. Requirement for deoxycytidine kinase in T and B lymphocyte development. Proc Natl Acad Sci USA 107: 5551–5556.
Stentoft J . The toxicity of cytarabine. Drug Saf 1990; 5: 7–27.
van Pelt K, de Haan G, Vellenga E, Daenen SM . Administration of low-dose cytarabine results in immediate S-phase arrest and subsequent activation of cell cycling in murine stem cells. Exp Hematol 2005; 33: 226–231.
Brennig S, Rattmann I, Lachmann N, Schambach A, Williams DA, Moritz T . In vivo enrichment of cytidine deaminase gene-modified hematopoietic cells by prolonged cytosine-arabinoside application. Cytotherapy 2012; 14: 451–460.
Modlich U, Navarro S, Zychlinski D, Maetzig T, Knoess S, Brugman MH et al. Insertional transformation of hematopoietic cells by self-inactivating lentiviral and gammaretroviral vectors. Mol Ther 2009; 17: 1919–1928.
Cattoglio C, Facchini G, Sartori D, Antonelli A, Miccio A, Cassani B et al. Hot spots of retroviral integration in human CD34+ hematopoietic cells. Blood 2007; 110: 1770–1778.
Schroder AR, Shinn P, Chen H, Berry C, Ecker JR, Bushman F . HIV-1 integration in the human genome favors active genes and local hotspots. Cell 2002; 110: 521–529.
Montini E, Cesana D, Schmidt M, Sanvito F, Ponzoni M, Bartholomae C et al. Hematopoietic stem cell gene transfer in a tumor-prone mouse model uncovers low genotoxicity of lentiviral vector integration. Nat Biotechnol 2006; 24: 687–696.
Lachmann N, Jagielska J, Heckl D, Brennig S, Pfaff N, Maetzig T et al. MicroRNA-150-regulated vectors allow lymphocyte-sparing transgene expression in hematopoietic gene therapy. Gene Therapy 2011, e-pub ahead of print 6 October 2011; doi: 10.1038/gt.2011.148.
Favre D, Blouin V, Provost N, Spisek R, Porrot F, Bohl D et al. Lack of an immune response against the tetracycline-dependent transactivator correlates with long-term doxycycline-regulated transgene expression in nonhuman primates after intramuscular injection of recombinant adeno-associated virus. J Virol 2002; 76: 11605–11611.
Latta-Mahieu M, Rolland M, Caillet C, Wang M, Kennel P, Mahfouz I et al. Gene transfer of a chimeric trans-activator is immunogenic and results in short-lived transgene expression. Hum Gene Ther 2002; 13: 1611–1620.
Mackall CL, Fleisher TA, Brown MR, Andrich MP, Chen CC, Feuerstein IM et al. Age, thymopoiesis, and CD4+ T-lymphocyte regeneration after intensive chemotherapy. N Engl J Med 1995; 332: 143–149.
Gregori S, Tomasoni D, Pacciani V, Scirpoli M, Battaglia M, Magnani CF et al. Differentiation of type 1 T regulatory cells (Tr1) by tolerogenic DC-10 requires the IL-10-dependent ILT4/HLA-G pathway. Blood 116: 935–944.
Benabdellah K, Cobo M, Munoz P, Toscano MG, Martin F . Development of an all-in-one lentiviral vector system based on the original TetR for the easy generation of Tet-ON cell lines. PLoS One 6: e23734.
Szulc J, Wiznerowicz M, Sauvain MO, Trono D, Aebischer P . A versatile tool for conditional gene expression and knockdown. Nat Methods 2006; 3: 109–116.
Dull T, Zufferey R, Kelly M, Mandel RJ, Nguyen M, Trono D et al. A third-generation lentivirus vector with a conditional packaging system. J Virol 1998; 72: 8463–8471.
Maetzig T, Brugman MH, Bartels S, Heinz N, Kustikova OS, Modlich U et al. Polyclonal fluctuation of lentiviral vector-transduced and expanded murine hematopoietic stem cells. Blood 117: 3053–3064.
Heckl D, Wicke DC, Brugman MH, Meyer J, Schambach A, Busche G et al. Lentiviral gene transfer regenerates hematopoietic stem cells in a mouse model for Mpl-deficient aplastic anemia. Blood 117: 3737–3747.
Zhang CC, Lodish HF . Murine hematopoietic stem cells change their surface phenotype during ex vivo expansion. Blood 2005; 105: 4314–4320.
We thank Matthias Ballmaier (PhD) and his team from the Core-Facility Cell-Sorting of Hannover Medical School for cell sorting, and Doreen Lüttge (Hannover Medical School) for excellent technical assistance. Furthermore, we thank Georg Kensah (Hannover Medical School) for the help in performing the timelapse video studies. This work was supported by grants from the Deutsche Forschungsgemeinschaft: Cluster of Excellence REBIRTH (Exc 62/1), SPP1230 Grant MO 886/3-1 (UM and TM) and by the EU framework programme grant PERSIST (CHB).
The authors declare no conflict of interest.
Supplementary Information accompanies the paper on Gene Therapy website
About this article
Cite this article
Lachmann, N., Brennig, S., Pfaff, N. et al. Efficient in vivo regulation of cytidine deaminase expression in the haematopoietic system using a doxycycline-inducible lentiviral vector system. Gene Ther 20, 298–307 (2013). https://doi.org/10.1038/gt.2012.40
- cytidine deaminase
- drug resistance
- inducible expression
Molecular Therapy (2019)
The CpG-sites of the CBX3 ubiquitous chromatin opening element are critical structural determinants for the anti-silencing function
Scientific Reports (2017)
Induced neural stem/precursor cells for fundamental studies and potential application in neurodegenerative diseases
Neuroscience Bulletin (2015)
Deoxycytidine-kinase knockdown as a novel myeloprotective strategy in the context of fludarabine, cytarabine or cladribine therapy