VSV-G-pseudotyped third generation bidirectional Lentiviral Vectors (bdLV) were produced by calcium phosphate transfection into 293T cells and concentrated by ultracentrifugation as described previously (27). Titer was estimated by limiting dilution: vector particles were measured by HIV-1 Gag p24 Ag immune capture (NEN Life Science Products, MA, USA), and vector infectivity was calculated as the ratio between titer and total particles. Titers ranged between 5 × 10⁸ and 6 × 10⁹ transducing units/mL, while infectivity between 5 × 10⁴ and 10⁵ transducing units/ng p24. To produce concentrated Vpx-incorporating viral-like particles (VLPs), 293T cells were co-transfected with a VSV-g expressing plasmid and the Simian Immunodeficiency Virus-derived packaging plasmid SIV3+, as previously described (26). For bioluminescence imaging (BLI), luciferase-encoding cDNA was cloned into in LV-GFP instead of the GFP gene and into LV-IL10 instead of ΔNGFR gene to allow in vivo tracking of transduced murine DC (DC^NGFR and DC^IL−10, respectively).

Human peripheral blood was obtained from healthy donors in accordance with local committee approval (TIGET09), and with the Declaration of Helsinki. Peripheral blood mononuclear cells were isolated by density gradient centrifugation over Lymphoprep™ (Axis-Shield PoC AS, Norway).

CD14⁺ cells were isolated from PBMC by positive selection using CD14 MicroBeads (Miltenyi Biotech, Germany) according to the manufacturer's instructions. Cells were cultured in RPMI 1640 (Lonza, Switzerland) with 10% fetal bovine serum (FBS) (Euroclone, Italy), 100 U/ml penicillin/streptomycin (Lonza, Switzerland), 2 mM L-glutamine (Lonza, Switzerland), at 10⁶ cells/ml in a 1 ml volume in a 24-well culture plate, supplemented with rhGM-CSF (Miltenyi Biotech, Germany) at 100 ng/ml and rhIL-4 (Miltenyi Biotech, Germany) at 10 ng/ml for 7 days at 37°C with 5% CO₂. One ml per well of fresh pre-warmed medium with cytokines, at final concentration as above, was added on day 3. To obtain mature DC (mDC), un-transduced DC were activated at day 5 with 1 μg/ml of LPS (Sigma Aldrich, CA, USA). For DC transduction, monocytes were exposed for 6 h to Vpx-VLP and then were transduced with the indicated vectors at Multiplicity of Infection (MOI) of 5 at day 0, 2, or 5. After overnight incubation, half of the medium was replaced with fresh medium supplemented with cytokines to dilute the vector concentration. For DC^IL−10 generation, 10 ng/ml of rhIL-10 (CellGenix, Germany) was added at day 0. In some experiments, DC^IL−10 were activated at day 6 with 1 μg/ml of LPS (Sigma Aldrich, CA, USA) or with 10 μg/ml of Poli (I:C) (InvivoGen, CA, USA). DC were harvested on day 7 for phenotypical, molecular, and functional analyses.

In some experiments, 10⁵ DC were plated in 200 μl of final volume, alone or in the presence of the following stimulation: 1 μg/ml of LPS (Sigma Aldrich, CA, USA), 10⁸ cells/ml of Heat Killed Listeria Monocytogenes, 1 μg/ml of Flagellin S. typhimurium, 10 μg/ml of Poli (I:C), 5 μM of ODN2006 (CpG) (InvivoGen, CA, USA) or a mix of 10 ng/ml for each cytokine of IL-1β, TNF-α, and IL-6 (R&D Systems, MN, USA). After 24 h, supernatants were collected to evaluate the cytokine secretion profile by ELISA, and cells were analyzed by flow cytometry.

CD3⁺, CD4⁺, and CD8⁺ T cells were purified from PBMC by negative selection using their respective human T cell Isolation kit (Miltenyi Biotech, Germany) according to the manufacturer's instructions. All T cell cultures were performed in X-VIVO 15 medium (Lonza, Switzerland), supplemented with 5% human serum (Sigma Aldrich, CA, USA), and 100 U/ml penicillin/streptomycin (Lonza, Switzerland). T cells were labeled with Cell Proliferation Dye eFluor^® 670 (eBioscience, CA, USA) according to manufacturer's instructions and stimulated with 10⁴ allogeneic DC (10:1, T:DC). After 5 days, T cells were collected, washed, and their phenotype and proliferation were analyzed by flow cytometry.

For T cell differentiation, 10⁶ CD4⁺ T cells were cultured with 10⁵ allogeneic DC (10:1, T:DC). After 10 days, primed T cells were collected and purified using CD4 Microbeads (Miltenyi Biotech, Germany). T cells stimulated with DC^UT are referred to as T(DC^UT) cells, while those stimulated with DC^GFP as T(DC^GFP) cells. T cells cultured with unstimulated DC^IL−10 are referred to as T(DC^IL−10) cells, while those cultured with LPS- or Poli I:C-stimulated DC^IL−10 are referred to as T(DC^{IL−10−LPS}) or T(DC^{IL−10−POLI}) cells, respectively.

For recall response proliferation, primed CD4⁺ T cells were stained with Cell Proliferation Dye eFluor^® 670 (eBioscience, CA, USA) and plated with DC^UT from the same donor used for priming (10:1, T:DC). After 3 days of stimulation, T cells were collected, washed, and proliferation was evaluated by flow cytometry.

To evaluate the suppressive activity of T(DC^IL−10), T(DC^{IL−10−LPS}), or T(DC^{IL−10−POLI}) cells, we stained total CD4⁺ T cells (responder cells) autologous to T cells used in priming with Cell Proliferation Dye eFluor^® 450 (eBioscience, CA, USA), and activated them with mDC from the same donor used for priming. T(DC^IL−10), T(DC^{IL−10−LPS}), or T(DC^{IL−10−POLI}) cells stained with Cell Proliferation Dye eFluor^® 670 (eBioscience, CA, USA), were added at a 1:1 ratio with responder cells (total T:DC ratio is 10:1). After 4 days, the percentages of divided responder T cells were calculated by proliferation dye dilution by flow cytometer.

For DC, the indicated number of cells were plated in 200 μl of final volume and left unstimulated or activated with 200 ng/ml of LPS (Sigma, CA, USA) and 50 ng/ml of IFN-γ (R&D System, MN, USA). Supernatants were collected after 48 h and levels of IL-6, IL-10, IL-12, and TNF-α were tested.

For CD4⁺ T cells, IL-10, and IFN-γ production was quantified in co-culture supernatants. Cytokine concentration was evaluated by standard sandwich ELISA, with purified and biotinylated antibody couples (Becton Dickinson, CA, USA).

NSG, Balb/c and C57Bl/6 female mice were purchased from Charles-River Italia. All mice were fed standard laboratory diet and maintained under standard laboratory conditions free of specific pathogens. All animal care procedures were performed according to protocols approved by the OSR Institutional Animal Care and Use Committee (IACUC protocol #488, #632, and #748), following the 3R principles (replacement, reduction, and refinement) and the Decreto Legislativo #116 dated January 27th, 1992, from the Italian Parliament.

Two–five days old NSG (NOD.Cg-PrkDC^cid Il2rg^tm1WjI/SzJ, JAX mouse strain) mice were sub-lethally irradiated (1.5cGy) and injected intra-hepatically 5–7 h later with 10⁵ CD34⁺ (purity ≥ 95%, Lonza), as previously described (28). Percentages of human cells in peripheral blood were monitored by flow cytometry starting from 8 weeks post-transplant. Once human engraftment was stable and T cell repopulation detectable (usually around 11–13 weeks post-transplant), humanized mice (huMice) were immunized by intravenous injection (i.v.) of 5 × 10⁶ irradiated allogeneic CD3⁻ cells (6,000 rad), magnetically isolated with Dynabeads CD3 (Thermo Fisher Scientific, MA, USA) from human PBMC. One week later, human T cell percentages were assessed by flow cytometry, huMice were randomly assigned to experimental groups and injected with 3 × 10⁵ un-transduced DC, or 3 × 10⁵ un-transduced plus 3 × 10⁵ transduced DC, differentiated from CD14⁺ monocytes isolated from the same donor used for CD3⁻ purification. After peripheral blood phenotyping at day 10 and 13, huMice were suppressed at day 13 and peripheral blood harvested.

Female Balb/c mice were sacrificed and bone marrow (BM) was harvested. BM cells were cultured in IMDM (Lonza, Switzerland) with 10% fetal bovine serum (FBS) (Euroclone, Italy), 100 U/ml penicillin/streptomycin (Lonza, Switzerland), 2 mM L-glutamine (Lonza, Switzerland), at 10⁶ cells/ml in a 1 ml volume in a 24-well culture plate, supplemented with rmGM-CSF (R&D Systems, MN, USA) at 25 ng/ml for 9 days at 37°C with 5% CO₂. One ml per well of fresh pre-warmed medium with rmGM-CSF, at final concentration as above, was added on day 3 and, after removal of 1 ml of supernatant from the culture, at day 5. DC were activated at day 7 with 200 ng/ml of LPS (Sigma Aldrich, CA, USA). For DC transduction, cells were transduced with indicated vectors at MOI of 10 at day 2.

Spleen mononuclear cells were isolated from female C57Bl/6 (H-2b) mice and stained with Proliferation Dye eFluor^® 670 (eBioscience, CA, USA). 10⁵ splenocytes were plated with 10⁴ DC^GFP or DC^IL−10 differentiated from BM of female Balb/c mice and collected after 5 days to assess the proliferation by dye dilution.

Female Balb/c (H-2d) mice were lethally irradiated (10 Gy) and intravenously injected with 2 × 10⁶ DC^NGFR or DC^IL−10, with or without the addition of 10⁷ Balb/c (H-2d) BM cells. Small-animal bioluminescence imaging (BLI) was performed using the IVIS Spectrum CT System (Perkin Elmer). The system is composed of a low-noise, back-thinned, back-illuminated charge-coupled device (CCD) camera cooled at −90°C with a quantum efficiency in the visible range above 85%. Each mouse received an intravenous injection of 150 mg luciferin/kg body weight 10 min before BLI. During image acquisition, the animals were kept at 37°C and under gaseous anesthesia (2–3% isoflurane and 1 L/min oxygen). Dynamic BLI was performed by acquiring a set of images every 2 min from 10 to 20 min after luciferin injection to detect the highest BLI signal. The images were obtained using the following settings: exposure time = auto, binning = 8, and field of view = 23.4 cm. Dark images were acquired before and then subtracted to bioluminescence images; no emission filters were used during BLI acquisitions. BLI image analysis was performed by placing a region of interest (ROI) over the body of the mouse (tail excluded) and by measuring the total flux (photons/seconds) within the ROI. Images were acquired and analyzed using Living Image 4.5 (Perkin Elmer).

Acute graft vs. host disease (GvHD) was induced by a single intravenous injection of BM cells (10 × 10⁶) supplemented with 5 × 10⁶ spleen mononuclear cells isolated from female C57Bl/6 (H-2b) mice into recipient female Balb/c (H-2d) mice lethally irradiated (10 Gy total body irradiation). Recipients received single intravenous injections of host-matched DC^GFP or DC^IL−10 (2 × 10⁶) 3 days after transplantation. Recipients were monitored once every other day from the day of transplantation to determine survival time, body weight and score (fur, hunch, skin lesion, mobility). Moribund mice were euthanized for ethical reason (more than 25% of weight loss or score higher than six).

Fluorochrome-conjugated antibodies against the following antigens were used for human DC staining: NGFR, CD1a, CD14, CD83, CD86, HLA-DR, CD16, CD163 (Becton Dickinson, CA, USA), and CD141 (Miltenyi Biotech, Germany), HLA-G (Exbio, Czech Republic), ILT4 (Beckman Counter, NJ, USA). The following fluorochrome-conjugated antibodies were used for human CD3⁺ T cell staining: anti-CD3, anti-CD4, anti-CD8, and anti-CD45RA (Becton Dickinson, CA, USA), anti-CD49b and anti-LAG-3 (Miltenyi Biotec, Germany). For Tr1 cell detection, CD4⁺ T cells were stained as previously described (29).

Fluorochrome-conjugated antibodies against the following antigens were used for murine DC staining: NGFR, CD11c, CD80, CD83, CD86, I-A/I-E (Becton Dickinson, CA, USA). The following fluorochrome-conjugated antibodies were used for murine splenocyte staining: anti-CD3, anti-CD4, and anti-CD8 (Becton Dickinson, CA, USA).

FcR Blocking Reagent (Miltenyi Biotech, Germany for human sample and Becton Dickinson, CA, USA for mice samples) was used in all preparations to avoid non-specific staining. Briefly, cells were centrifuged and re-suspended in Dulbecco's Phosphate-Buffered Saline (DPBS, Corning) supplemented with 2% FBS (Lonza, Switzerland). Cells were incubated at room temperature for 15 min, centrifuged and fixed with 1% formaldehyde solution methanol-free (Thermo Fisher Scientific, MA, USA). For cultured cells, the described passages where preceded by staining with LIVE/DEAD™ Fixable Dead Cell Stain Kit (Invitrogen, CA, USA) following manufacturer's instructions.

For huMice blood staining, 100 μl of whole blood was stained with antibodies against surface markers for 15 min at room temperature. Cells were then fixed, permeabilized and stained with anti-KI67 (Becton Dickinson, CA, USA) using Foxp3/Transcription Factor Staining Buffer Set (eBioscience, USA). Samples were acquired using the FACSCanto II or Fortessa Flow Cytometers (Becton Dickinson, CA, USA) and data were analyzed with FlowJo software (FlowJo LLC, USA).

Wilcoxon matched pairs test (two-tailed) were used for statistical analysis. All results are presented as mean values ± standard deviation, unless differently specified in the figure legend. Differences were regarded as significant at ^*P ≤ 0.05, ^**P ≤ 0.01, and ^***P ≤ 0.001. Results were analyzed using GraphPad Prism 5.0 (GraphPad Software, CA, USA).

Lentiviral vector (LV) transduction of monocyte-derived DC has been described; however, transduction efficiency is generally low due to the inhibition of reverse transcription in human myeloid cells mediated by SAMHD1 (19, 20, 23). The use of polybrene in combination with simian immunodeficiency virus (SIV)-derived accessory protein Vpx during LV transduction has been proposed to overcome this limitation (26, 30). We optimized the transduction protocol of human monocyte-derived DC with LV encoding large size plasmid (~10 kb), by pre-treating CD14⁺ cells with viral-like particles containing Vpx (Vpx-VLP) in the absence of polybrene before exposure to a bdLV co-encoding for GFP and ΔNGFR (LV-GFP). CD14⁺ cells were pre-treated or not with Vpx-VLP for 6 h on day 0, 2, or 5 before LV-GFP exposure during monocyte-derived DC differentiation, and transduction efficiency was evaluated at the end of differentiation (Figure S1A). Pre-treatment with Vpx-VLP at all time points analyzed improved transduction, reaching the highest efficiency when cells were pre-treated with Vpx-VLP at day 0 (up to 95% of ΔNGFR⁺ DC, Figure S1B). As expected, in the absence of Vpx-VLP, DC were transduced at very low levels (<6% of ΔNGFR⁺ DC), irrespectively of the day of transduction (Figure S1B). We then applied the above protocol to transduce CD14⁺ cells with a bdLV co-encoding for IL-10 and ΔNGFR (LV-IL-10) (31). Although the transduction efficiency of DC^IL−10 was similar to that of DC^GFP (not shown), the differentiated population contained two distinct cell subtypes (CD14⁻CD16⁻ and CD14⁺CD16⁺; Figure S1C). Since the presence of IL-10 from day 0 of differentiation results in high CD14 and CD16 expression in DC-10 (32), we hypothesized that the observed heterogenicity in DC^IL−10 was due to a later exposure to IL-10. For this reason, we added exogenous IL-10 at day 0 during CD14⁺ cell pre-treatment with Vpx-VLP and exposure to LV-IL-10 (Figure 1A) and we obtained a homogenous population of CD14⁺CD16⁺ DC^IL−10 (Figure S1C), with an average transduction efficiency of 90% for DC^IL−10, which was superimposable to that of DC^GFP (Figure 1B). In conclusion, we established an efficient protocol to transduce human monocytes during DC differentiation with large-size LV.

We previously described a subset of human DC, called DC-10, that are differentiated in vitro through exposure of monocytes to IL-10 during DC differentiation (32) and, recently, we identified DC-10-specific biomarkers (33). Since DC^IL−10 are differentiated in the presence of IL-10, it was not surprising that they expressed the DC-10 markers CD14, CD16, CD141, and CD163 at significantly higher levels compared to un-transduced (DC^UT) and LV-GFP transduced (DC^GFP) cells, while the levels of CD1a, marker of the classical monocyte-derived DC, were significantly lower (Figure 2A). We next evaluated the expression of MHC class II and costimulatory molecules, important for appropriate antigen presentation to and activation of T cells. In all donors tested, HLA-DR expression in DC^IL−10 was significantly higher than in DC^UT and DC^GFP, while the expression of CD83 and CD86 on DC^IL−10 was variable among donors tested, with some donors showing higher and others similar expression levels compare to control DC (Figure 2B). The levels of the tolerogenic molecules HLA-G and ILT4, known to be required for T regulatory type 1 (Tr1) cell induction via DC-10 (32), were significantly higher on DC^IL−10 compared to control DC (Figure 2C).

As expected, DC^IL−10 secreted high amounts of IL-10 at steady state and upon LPS/IFN-γ stimulation, in the absence of IL-12. Conversely, stimulated DC^UT and DC^GFP produced high levels of IL-12 and variable levels of IL-10, always lower compared to that of DC^IL−10. There were no differences in the secretion of IL-6 upon stimulation, while higher IL-6 secretion in DC^IL−10 compared to DC^UT at steady state was detected. TNF-α release was not observed in the absence of stimulation, whereas the lowest levels of TNF-α were measured in stimulated DC^IL−10 compared to controls (Figure 3). Overall, DC^IL−10 showed a high IL-10/TNF-α and IL-10/IL-12 ratio indicative of their skewing toward tolerance, opposite to the pro-inflammatory cytokine profile displayed by DC^UT and DC^GFP. Altogether, our results show that stable and enforced expression of IL-10 in human monocyte-derived DC through bdLV-mediated gene transfer promotes the differentiation of mature tolerogenic DC-10-like cells, which express HLA-G and ILT4.

We then proceeded to the functional characterization of DC^IL−10 and found that they induced significantly lower proliferative responses in allogeneic CD3⁺ T cells when compared to that elicited by control DC^UT and DC^GFP, in both CD4⁺ and CD8⁺ T cells (Figure 4A). According to their similarity to DC-10 and their ability to secrete high levels of IL-10, stimulation of allogeneic CD4⁺ T cells with DC^IL−10 allowed the differentiation of cells—T(DC^IL−10) cells—that contained a significantly higher proportion of Tr1 cells compared to T cells primed with DC^UT and DC^GFP—T(DC^UT) and T(DC^GFP) cells, respectively—(Figure 4B). In line with the presence of Tr1 cells, T(DC^IL−10) cells re-stimulated with mature DC (mDC), autologous to DC used for priming, were hypo-responsive (Figure 4C), and produced significantly higher level of IL-10, but similar levels of IFN-γ compared to both T(DC^UT) and T(DC^GFP) cells (Figure 4D). Notably, anergy was alloantigen-specific, since the stimulation with a third party mDC induced comparable levels of proliferation in all primed T cells (Figure S2). Moreover, T(DC^IL−10) cells suppressed the proliferation of autologous CD4⁺ T cells stimulated with mDC from the same donor used for priming, with an average of 67% of suppression (Figure 4E). Overall, these findings indicate that bdLV-mediated IL-10 gene transfer in DC induces a cell population endowed with the ability to modulate allogeneic T cell responses and promote the differentiation of alloantigen-specific Tr1 cells in vitro.

One of the major hurdles in the use of tolDC as cell product is their stability, thus we investigated the impact of a pro-inflammatory milieu on the phenotype and functions of DC^IL−10. To this end, we assessed phenotype and function of DC^IL−10 stimulated in vitro with different toll-like receptor (TLR) agonists (LPS, Listeria, Flagellin, Poli I:C, and CpG) or with a mix of pro-inflammatory cytokines (IL-1β, TNF-α, and IL-6). No major changes in DC^IL−10 phenotype were observed upon activation, with the exception of a significant down-regulation of CD16 upon Listeria and CpG stimulation, and a significant upregulation of CD86 upon Listeria- and LPS-mediated activation (Figures 5A,B). In line with results obtained in DC-10 (33), DC^IL−10 stably expressed CD141, and CD163 independently from the stimuli used (Figure 5A). Furthermore, DC^IL−10 maintained their ability to secrete high amounts of IL-10 in the absence of IL-12, independently from the stimulation used. Activation with LPS, Listeria, and Flagellin of DC^IL−10 promoted significantly higher levels of IL-6 and TNF-α compared to unstimulated conditions (Figure S3). Thus, DC^IL−10 are phenotypically stable cells and activation with some TLR agonists further promote their activation, as demonstrated by the up-regulation of CD86 and the increase in IL-6 and TNF-α secretion.

Analysis of the expression of the tolerogenic molecules HLA-G and ILT4 on DC^IL−10 showed that, among the different stimuli used, TLR3- and TLR4-mediated activation had opposite effects: Poli I:C exposure increased HLA-G and decreased ILT4, while LPS stimulation down-regulated HLA-G and up-regulated ILT4 (Figure 6A, Figure S4). Since HLA-G and ILT4 are involved in DC-10-mediated induction of Tr1 cells (32), we investigated how DC^IL−10 tolerogenic activity varied upon stimulation with LPS (DC^{IL−10−LPS}) and Poli I:C (DC^{IL−10−POLI}). In line with the changes in HLA-G levels, the percentage of Tr1 cells induced at the end of T cell differentiation was higher in T cells induced by DC^{IL−10−POLI} and lower in T cells induced by DC^{IL−10−LPS}. Despite these differences, all the three DC^IL−10 populations induced Tr1 cells at a higher efficiency when compared to control DC^GFP (Figure 6B). No major differences in proliferative capacity, cytokine profile and suppressive ability were observed among Tr1 cells generated with unstimulated DC^IL−10, DC^{IL−10−LPS}, and DC^{IL−10−POLI}, showing that DC^IL−10 maintained their tolerogenic activity upon pro-inflammatory stimulation (Figures 6C,D). Overall, these data indicate that activated DC^IL−10 are as powerful as their steady state counterpart in modulating T cell responses and in promoting Tr1 cells in vitro, suggesting that activation does not impair the modulatory activity and tolerogenic potential of DC^IL−10.

To assess the modulatory activity of DC^IL−10 in vivo we took advantage of the recently developed protocol for the repopulation of NSG mice with human cord blood CD34⁺ cells. Intra-liver injection of human CD34⁺ cells in sub-lethally irradiated neonate NSG mice allowed efficient engraftment of human CD45⁺ hematopoietic cells in bone marrow (BM) and differentiation of lymphoid (B and T effector, and T regulatory) and myeloid cells in the periphery (28). We immunized reconstituted huMice by injection of irradiated allogeneic human CD3⁻ cells, which act as antigen presenting cells, and provided, after seven days, a second challenge by injecting autologous DC^UT. To assess the modulatory activity of DC^IL10, we co-injected DC^IL−10 (DC^UT+DC^IL10) or, as control, DC^GFP (DC^UT+DC^GFP) (Figure 7A). While treatment with DC^GFP induced a boost in CD4⁺ T cell proliferation in vivo, as assessed by Ki67 staining of peripheral blood lymphocytes 3 days after the re-challenge, DC^IL−10 dampened the response induced by allogeneic DC^UT. In all the three conditions, the immune system returned at steady state after 5 days from the re-challenge, with comparable proliferation levels observed in all mice (Figure 7B).

We next investigated the potential therapeutic effect of DC^IL−10 in a pre-clinical model of acute graft-vs. host disease (GvHD). To do so, we generated and characterized the murine counterpart of the human DC^IL−10. We transduced Balb/c (H-2d) BM cells with LV-IL-10 and LV-GFP at day 2 during DC differentiation and activated them with LPS in the last 2 days of culture. Murine DC were efficiently transduced with LV-IL-10 and LV-GFP, as demonstrated by an average 67.3 and 67.1% of ΔNGFR⁺ cells, respectively (Figure 8A). In contrast to DC^GFP, DC^IL−10 showed lower expression levels of CD83, while the expression of MHC class II, CD80, and CD86 was comparable (Figure 8A). As expected, DC^IL−10 showed human IL-10 expression at steady state and upon activation, which was not detected in DC^GFP (Figure 8B). Similar to human DC^IL−10, murine DC^IL−10 promoted a significantly lower proliferative allogeneic T cell response compared to that elicited by DC^GFP, both in CD4⁺ and CD8⁺ T cells (Figure 8C).

To study murine DC^IL−10 biodistribution, we transduced BM cells during DC differentiation with a bdLV encoding for luciferase and IL-10 (DC^IL−10) or ΔNGFR (DC^NGFR). DC^IL−10 and control DC^NGFR were then intravenously injected alone or in combination with autologous BM cells in lethally irradiated female Balb/c mice. The difference in the 1st day signals between DC^IL−10- and DC^NGFR-injected mice was very likely due to a higher transduction of DC^IL−10 (Figure 9A). In all groups we observed a consistent decrease in total flux in the first 2 days after DC injection, and both cell types localized in the lung (Figure 9B). Between day 2 and 5 lung signal dropped, and increased signal was registered in the legs of all mice (Figure 9C). When BM cells were co-injected, a different behavior was observed between DC^IL−10 and DC^NGFR. DC^NGFR migrated to the legs with the same kinetic irrespectively of the presence or absence of BM cells, at least until day 6. Conversely, migration of DC^IL−10 to the bones is limited by the presence of BM cells, while when mice are not replenished with BM cells it is increased, even at higher rate compared to DC^NGFR, probably due to an enhanced recruitment induced by the aplastic/hypoplastic bone marrow upon irradiation.

Finally, we evaluated the potential therapeutic effect of DC^IL−10 in GvHD. Lethally irradiated female Balb/c recipients were transplanted with BM cells and splenocytes isolated from female C57Bl/6 mice, and 2 days after transplantation mice received Balb/c-derived DC^IL−10 or DC^GFP, or were left untreated as control (Figure 10A). This experimental setting induced a strong GvHD, since all mice of the control group died within 24 days (Figure 10B). Administration of Balb/c DC^IL−10 improved the survival time of mice compared to both control groups (50% survival at day 24), while Balb/c DC^GFP treatment enhanced GvHD lethality (Figure 10B). Interestingly, treatment with DC^IL−10 clearly modulated not only the survival but also the severity of GvHD phenotype in mice, compare to both untreated and DC^GFP-treated mice groups, as shown by weight loss and score assessment at day 24 (Figure 10C). These results support the potential clinical application of DC^IL−10 as cell-based approach to control T cell responses in allogeneic transplantation setting.