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. 2015 Jul 14;23(9):1497–1506. doi: 10.1038/mt.2015.110

Mesenchymal Stromal Cells for Linked Delivery of Oncolytic and Apoptotic Adenoviruses to Non-small-cell Lung Cancers

Valentina Hoyos 1,2, Francesca Del Bufalo 1,2,*,2, Shigeki Yagyu 1, Miki Ando 1, Gianpietro Dotti 1, Masataka Suzuki 1,3, Lisa Bouchier-Hayes 1,4, Ramon Alemany 5, Malcolm K Brenner 1
PMCID: PMC4817892  PMID: 26084970

Abstract

Oncolytic adenoviruses (OAdV) represent a promising strategy for cancer therapy. Despite their activity in preclinical models, to date the clinical efficacy remains confined to minor responses after intratumor injection. To overcome these limitations, we developed an alternative approach using the combination of the OAdv ICOVIR15 with a replication incompetent adenoviral vector carrying the suicide gene of inducible Caspase 9 (Ad.iC9), both of which are delivered by mesenchymal stromal cells (MSCs). We hypothesized that coinfection with ICOVIR15 and Ad.iC9 would allow MSCs to replicate both vectors and deliver two distinct types of antitumor therapy to the tumor, amplifying the cytotoxic effects of the two viruses, in a non-small-cell lung cancer (NSCLC) model. We showed that MSCs can replicate and release both vectors, enabling significant transduction of the iC9 gene in tumor cells. In the in vivo model using human NSCLC xenografts, MSCs homed to lung tumors where they released both viruses. The activation of iC9 by the chemical inducer of dimerization (CID) significantly enhanced the antitumor activity of the ICOVIR15, increasing the tumor control and translating into improved overall survival of tumor-bearing mice. These data support the use of this innovative approach for the treatment of NSCLC.

Introduction

Oncolytic or conditionally replicating adenoviruses (OAdV/CRAd) represent a promising strategy for cancer therapy. CRAd can selectively replicate in and lyse tumor cells, and they are easy to manipulate genetically to incorporate genes of interest. Despite encouraging activity in preclinical models, to date CRAds have revealed only local, transient, and confined responses after intratumor injection in clinical trials.1,2,3 Intravenous administration of these adenoviruses is even less effective due to the widespread pre-existing immunity against this common pathogen. The virus also gets trapped in the liver.4,5,6 Moreover, CRAd replicates primarily in tumor cells, whereas resting/hypoxic areas of the tumor and tumor-associated stromal components may be infected without being killed.

In order to overcome the above limitations of CRAd therapy and increase its potency, we developed an alternative approach using our previously validated mesenchymal stromal cell (MSC) delivery system.7,8 MSCs home to inflammatory and tumor areas and are therefore an ideal cellular carrier for the systemic administration of CRAd.9,10,11 We have previously shown that when MSCs are forced to express the adenoviral E1A gene, they can replicate first-generation adenoviral vectors encoding an inducible caspase 9 (iC9) suicide gene and deliver these vectors to lung tumors in a model of human non-small-cell lung cancer (NSCLC).7 Following the administration of the chemical inducer of dimerization (CID), AP20187, iC9 expressed by the infected tumor cells activates the apoptosis pathway, thereby killing the cells. We hypothesize now that using MSC as producer cells for both CRAd and iC9 vectors could increase the potency and amplify the antitumor activity of the CRAd therapy. We determined if the CRAd component has the machinery necessary to replicate the two viruses both in MSCs and in tumor cells and thereby induce a self-amplifying circuit and potent antitumor effect. iC9 is aimed at extending the antitumor effect of the system by targeting the slow growing areas and the tumor-associated stroma, which are poorly sensitive to the oncolytic activity of the CRAd. We combined the CRAd ICOVIR15 (ref. 12) with a replication incompetent Ad5/35 iC9 in MSCs and present the results of this approach in vitro and in a human xenograft model of NSCLC.

Results

MSCs replicate both ICOVIR15 and a replication-incompetent adenoviral vector after coinfection

To assess the ability of the MSCs to replicate the replication-incompetent adenoviral vector after coinfection with ICOVIR15, we infected MSCs with either ICOVIR15 alone (50 vp/cell), a green fluorescent protein (GFP)-encoding first-generation Ad5/35 vector alone (Ad.GFP, 1,000 vp/cell) or both in combination at the same multiplicity of infection (MOI). On day 5 after infection, we transferred the supernatant to two NSCLC cell lines (A549 or H1299). After 4 days we verified that the supernatants contained ICOVIR 15 from the development of cytopathic effects. The replication of Ad.GFP in the MSC was assessed by immunofluorescence of the indicator cell lines after exposure to MSC supernatants. Supernatants from MSC infected with Ad.GFP alone produced no GFP expression in H1299 cells, whereas supernatants from MSC infected with ICOVIR15 alone produced progressive cytopathic effects on the indicator cells but no GFP expression (Figure 1a). Only when MSCs had been coinfected with ICOVIR15 and Ad.GFP were both oncolytic effects and GFP expression observed (Figure 1a), confirming the replication of both viruses by the MSCs. Identical results were obtained using A549 cells (data not shown).

Figure 1.

Figure 1

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ICOVIR15 enables MSCs to replicate the replication-incompetent adenoviral vector. (a) MSCs were infected with either Ad.GFP, ICOVIR15 or the combination of both and after 5 days the supernatant was transferred to a culture of A549 or H1299. Representative pictures of H1299 (light microscopy on the left panel, fluorescence microscopy on the right panel). Scale bars represent 100 µm. (b) MSCs were infected with either ICOVIR15 (I15) only or ICOVIR15 and Ad.iC9 (I15.iC9) and after 5 days the supernatant was transferred at serial dilutions on a 293T culture to perform a PFU assay. The viral titers measured after 5 days were compared to the PFU/ml at infection. Absolute titers (above panels) and the viral titer fold increase (below panel) are shown. (c) MSCs were infected with either Ad.iC9, ICOVIR15, or the combination of both. After 5 days, infected and uninfected MSCs were lysed, DNA was extracted and the copy numbers of E1a (representative of the ICOVIR15 production) and iC9 (representative of the Ad.iC9) were measured by real-time PCR. Results were normalized to the hGAPDH copy number. (d) Five days after infecting MSCs with the combination of ICOVIR15 and either Ad.GFP or Ad.iC9 and E-MSCs with either Ad.GFP or Ad.iC9, the supernatant was collected and serial dilutions were transferred to A549. The expression of GFP or ΔCD19 (linked to iC9), was measured by flow-cytometry (E-MSC supernatant, grey bars; MSCs supernatant, blue bars). Error bars represent standard deviation. MSCs, mesenchymal stromal cells.

We repeated these experiments, substituting Ad.iC9 for Ad.GFP and quantified the efficiency of MSCs infection by each virus alone or in combination. We infected the MSCs with ICOVIR15 and Ad.iC9 alone or in combination, and evaluated the iC9 and E1a copy numbers by quantitative polymerase chain reaction (qPCR) after 6 hours, in order to allow the viral infection without replication. Our results show that each virus alone can efficiently infect MSCs (copy numbers, normalized by human glyceraldehyde 3-phosphate dehydrogenase (GAPDH), of 7.84E-01 ± 2.12E-01 for iC9 and 1.52E00 ± 9.23E-01 for ICOVIR15, as compared to 2.57E-03 ± 3.31E-03 for iC9 and 2.16E-02 ± 2.75E-02 for ICOVIR15 of NT MSCs) and that there is no competition when viral combinations are used, likely because each chimeric adenovirus uses different cellular receptors (see Methods) (copy numbers, normalized by human GAPDH, of 1.97E00 ± 1.54E00 for iC9 and 7.96E-01 ± 2.56E-01 for ICOVIR15) (Supplementary Figure S1a).

To prove that MSCs can replicate both viruses and produce amplified viral progeny, we first infected MSCs with either ICOVIR15 or the combination of ICOVIR15 and Ad.iC9. After 5 days, we determined the overall titers of infectious virus present in the supernatants using a median tissue culture infective dose method (TCID50). The plaque forming unit (PFU)/ml measured were: at baseline, 2.66E02 ± 2.12E01 and 1.11E5 ± 6.74E04, and at 5 days after the infection, 5.15E10 ± 1.64E10 and 2.37E13 ± 1.11E13 (ICOVIR15 and ICOVIR15 + Ad.iC9, respectively). Since we used a TCID50 assay with E1 expressing 293T cells, the replication of the first-generation adenoviral vector is fully enabled, accounting for the difference in PFU/ml at baseline with and without the iC9 vector. Because this overall measurement of titer cannot distinguish the source of the viral product (ICOVIR versus iC9), we normalized the titer at d5 to the baseline and then measured the fold increase in the PFU/ml from each condition. The results were closely comparable (2.01E+08 ± 5.89E+07 for ICOVIR15 and 2.25E+08 ± 3.70E+07 for the combination of ICOVIR15 and Ad.iC9), as shown in Figure 1b, third panel, indicating that the ability of MSCs to replicate each adenovirus is not impaired by coinfection.

We then quantified the replication of each viral component by measuring the copy numbers of iC9 (i.e., Ad.iC9 replication) and E1A (i.e., ICOVIR15) in MSCs 5 days after infection either with one of the two viruses or with both. As expected, we obtained similarly increased copy numbers of E1A in both ICOVIR15 and ICOVIR15-Ad.iC9 MSCs, whereas iC9 could be detected only in the ICOVIR15-iC9 MSCs (Figure 1c).

MSCs infected with ICOVIR15 produce higher titers of Ad.iC9 or Ad.GFP than MSC expressing transgenic E1A

To determine the relative potency of our approach, we next compared the titer of replication incompetent virus produced by MSCs after coinfection with a CRAd to the titer of the virus made by “producer MSCs” expressing transgenic E1A4 (E-MSCs).7 We infected MSCs with ICOVIR15 and either Ad.GFP or Ad.iC9 and E-MSCs with either Ad.GFP or Ad.iC9. At day 5 after infection, serial dilutions of the supernatant were transferred from MSCs and E-MSCs to A549 and the expression of GFP or ΔCD19 (linked to iC9) was measured by flow-cytometry. The truncation mutant of CD19 has no activity but is still recognized by antibodies allowing us to measure the expression of the linked iC9 protein. Tumor cells transduced with the supernatants from coinfected MSCs had a higher percentage of transduced cells at a given concentration of supernatant (55 ± 1% at the highest concentration for CD19 expression; 99 ± 2% for GFP expression) and a higher mean fluorescence intensity (78 ± 0.64 for CD19 expression and 938 ± 31.27 for GFP expression) than tumor cells transduced with EMSC supernatant (20 ± 9% expression of CD19 and 93 ± 2% of GFP, P = 0.022 and P = 0.1, with mean fluorescence intensity of 2 ± 0.27 for CD19 and 105 ± 6.66 for GFP, P < 0.0001 and P = 0.0006) (Figure 1d). The superior transduction of tumor cells in the coinfected group is likely a consequence of the higher replication of Ad.GFP and Ad.iC9 in MSCs, producing a higher viral titer in the supernatant, and hence greater replication also in the A549 cells, demonstrating the amplification provided by this system. Thus, MSCs infected with both viruses replicate the adenoviral vectors (Ad.GFP or Ad.iC9) more efficiently than MSC expressing the E1A protein alone.

To further characterize the superior ability of ICOVIR15-infected MSCs to replicate the adenoviral vectors, compared to E-MSC (>70% transduced, Supplementary Figure S1b, left panel), we evaluated the level of expression of E1 gene through quantification of the relative mRNA expression by RT-PCR. As shown in the Supplementary Figure S1b (right graph), we found copy number values of 1.79E-02 ± 2.16E-02 for E-MSCs and 2.22E02 ± 4.11E01 for ICOVIR15-infected MSCs (at day 5), confirming the significantly higher replication in ICOVIR15-infected MSCs.

Tumor cell apoptosis is enhanced by the combination of ICOVIR15 and Ad.iC9

We then evaluated the antitumor effects of MSC infected with each virus alone and in combination. One day after infecting MSCs with either ICOVIR15, Ad.iC9 or ICOVIR15+Ad.iC9, we cocultured these cells with two different lung tumor cell lines (A549, H1299) at a MSC:tumor cell ratio of 1:5. On day 3 after coculture, 80 nmol/l of AP20187 (the chemical inducer of iC9 dimerization, CID) was added to the culture. After 24 hours, we measured apoptosis by flow cytometric analysis of Annexin V and 7-AAD staining of CD90 negative cells to exclude the MSCs. In both tumor cell lines, the administration of CID induced a statistically significant increase in apoptosis in the combination virus group (75 ± 1.73% in H1299, 87 ± 5% in A549) than ICOVIR15+Ad.iC9 without CID (52 ± 9.07%, P = 0.04 for H1299 and 68 ± 7%, P = 0.032 for A549), ICOVIR15 (52 ± 12.02%, P = 0.007 and 71 ± 2%, P = 0.013), iC9 (22 ± 2.08%, P = 0.001 and 16 ± 5%, P < 0.0001), NT (19 ± 5%, P = 0.004 and 11 ± 1%, P = 0.001) or untreated (4 ± 0.58%, P = 0.0006 and 5 ± 1%, P = 0.0001) (Figure 2).

Figure 2.

Figure 2

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The activation of iC9 by CID enhances tumor apoptosis in the ICOVIR15-Ad.iC9 treatment group. One day after infection of the MSCs with either ICOVIR15, Ad.iC9 or ICOVIR15+Ad.iC9, the cells were cocultured with two different lung tumor cell lines (A549, H1299) at a MSC: tumor cell ratio of 1:5 and 80 nmol/l of AP 20187 was added to the culture on day 3 after coculture. After 24 hours, we measured apoptosis of tumor cells (CD90 negative cells) by flow cytometric analysis of Annexin V and 7-AAD staining. Error bars represent standard deviation. MSCs, mesenchymal stromal cells.

After intravenous infusion MSCs persist only at tumor sites in lung

To evaluate the fate of intravenously infused MSC containing ICOVIR15 and Ad.iC9, we used a SCID-Beige mouse lung cancer xenograft model and infused 2.5 × 106 FFLuc-GFP labeled A549 cells intravenously. After 4 days, the mice were injected with: vehicle alone; 1 × 106 intravenous uninfected MSCs; Ad.iC9-MSCs; ICOVIR15-MSCs; or ICOVIR15-Ad.iC9-MSCs. After 4, 5, and 7 days, intraperitoneal CID (50 µg) was administered. At day 8, a second cycle of MSCs and CID was repeated.

To evaluate the distribution of the MSCs, three mice each time-point were euthanized, 3, 24, 48, and 72 hours after MSC administration. Lung tissues were snap-frozen, sectioned, and analyzed by immunofluorescence after staining with an anti-CD90 monoclonal antibody to detect human MSCs. As shown in Figure 3a (with additional pictures in Supplementary Figure S2a), 3 hours after the infusions MSCs (34 ± 5 cells/10 hpf) were found scattered randomly in the lung tissue. After 72 hours, however, MSCs (2 ± 1 cells/10 hpf) could only be detected in areas surrounding the tumor tissue and were absent elsewhere.

Figure 3.

Figure 3

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MSCs home to the tumor areas and deliver both ICOVIR15 and Ad.iC9. (a) SCID-Beige mice were engrafted with FFLuc-GFP labeled A549 cells (green) intravenously (IV) followed by IV infusion of either vehicle or uninfected MSCs, Ad.iC9-MSCs, ICOVIR15-MSCs, or ICOVIR15-Ad.iC9-MSCs. Twelve mice (three mice/group) were then sacrificed 3, 24, 48, and 72 hours after MSCs administration. Lung tissues were snap-frozen, sectioned, and analyzed by immunofluorescence after staining with an anti-CD90 monoclonal antibody and Alexa-Fluor 594-conjugated secondary antibody (red) to detect human MSCs and counterstaining with 4′,6-diamidino-2-phentlindole (DAPI) (blue). Green represents tumor cells (GFP-transduced). After 3 hours, 34 ± 5 MSCs/10 hpf could be detected in the lung tissue versus 2 ± 1/10 hpf at 72 hours. Representative confocal microscope images of 3 and 72 hours time points are shown. Scale bars represent 100 µm. (b) Three mice/group in the untreated, ICOVIR15-MSCs and ICOVIR15-Ad.iC9-MSCs groups were euthanized 5 days after infusion. Lung tissues were collected and qPCR for E1A on the extracted DNA was performed. (c) Lung tissues from untreated, Ad.iC9-MSCs, ICOVIR15-MSCs, and ICOVIR15-Ad.iC9-MSCs groups were snap-frozen and stained with antiadenoviral capsid proteins antibody and AlexaFluor594-conjugated secondary antibody (red) and counterstained with DAPI (blue). Green represents tumor cells (GFP-transduced). Representative confocal microscope images are shown. (d) Lung tissues from untreated, Ad.iC9-MSCs, and ICOVIR15-Ad.iC9-MSCs groups and from a tumor-free mouse, for comparison, were snap-frozen and stained with anti-CD19 antibody and AlexaFluor 594-conjugated secondary antibody (red) and counterstained with DAPI (blue). Green represents tumor cells (GFP-transduced). Representative confocal microscope images are shown. Scale bars represent 100 µm. MSCs, mesenchymal stromal cells.

MSCs deliver ICOVIR15 and Ad.iC9 to tumor tissues

To verify the delivery of the ICOVIR15 to the tumor sites, 6 days after the first MSC administration, three of the mice from the untreated group, three from the ICOVIR15 group and three from the ICOVIR15-Ad.iC9 group were sacrificed and lung tissues were analyzed for the presence of the CRAd by quantifying copy numbers of E1A (i.e., ICOVIR15). As shown in the Figure 3b, ICOVIR15 was present in both treatment groups (copy numbers, normalized by human GAPDH, of 76.3 ± 35.3 and 66.5 ± 27.1 in the ICOVIR15 and ICOVIR15-Ad.iC9 groups, respectively), whereas the untreated group was negative (1.64 ± 2.04 copy numbers). These data suggest that the ICOVIR15 is successfully delivered to the tumor environment by the MSCs. To further demonstrate that the delivery selectively targets the tumor, we stained lung sections of mice in the untreated and Ad.iC9-MSCs, ICOVIR15-MSCs, ICOVIR15-Ad.iC9-MSCs treatment groups for the adenoviral capsid proteins. As shown in Figure 3c (and in additional pictures in Supplementary Figure S2b), adenoviral infection was observed in the tumor areas of ICOVIR15- and ICOVIR15-Ad.iC9-MSCs treatment groups.

Having shown the delivery of the adenoviruses, we wanted to verify the ability to induce transduction of iC9 to the tumor cells by checking the expression of the CD19 marker by immunofluorescence in the untreated, Ad.iC9 and ICOVIR15-Ad.iC9 groups. In the first two groups, no CD19 was detected; by contrast, tumor tissues of the ICOVIR15-Ad.iC9-MSCs mouse were positive for the iC9-associated marker CD19 (Figure 3d with additional pictures in Supplementary Figure S3a).

Tumor tissues of mice treated with MSCs carrying ICOVIR15 and Ad.iC9 have increased apoptosis after iC9 activation

We evaluated the apoptotic effect of iC9 activation by immunofluorescence, using an in situ terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay. As shown in Figure 4 (with additional pictures in Supplementary Figure S3b), we detected minimal TUNEL staining in the lung cancer tissue after CID administration in the iC9-treated mice, likely due to the lack of amplification of the MSC-associated iC9 virus (Figure 4a). In the mice receiving MSCs containing both ICOVIR15-iC9 (in which the iC9 virus is amplified during ICOVIR replication) TUNEL staining was detected in the tumor areas and is significantly increased after iC9 activation, as shown by increased fluorescent signal measurements (0.63 ± 0.12 versus 1.55 ± 0.16; P = 0.01) (Figure 4b).

Figure 4.

Figure 4

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Tumor apoptosis is enhanced in the ICOVIR15-Ad.iC9-MSCs treated mice after CID administration. (a) Lung tissues of mice treated with Ad.iC9-MSCs+CID, ICOVIR15-Ad.iC9-MSCs and ICOVIR15-Ad.iC9-MSCs+CID were analyzed by confocal microscope after in situ TUNEL staining. Tumor (green), nuclei (blue) and TUNEL staining (red) are shown. Representative confocal microscope images are shown. Scale bars represent 100 µm. (b) Apoptosis-associated fluorescence was measured on 10 different fields using the NIH software ImageJ. MSCs, mesenchymal stromal cells.

Normal tissues showed no evidence of MSC infiltration, iC9 transduction, or adenoviral infection

To assess the safety of our approach, we collected heart, kidney, liver, and spleen tissues from mice in the ICOVIR15-iC9 treatment group and compared them with the same tissues from untreated mice. We initially reviewed the hematoxylin/eosin staining of these normal tissues with an independent pathologist who observed no signs of adenoviral infection/disease (Supplementary Figure S4a). To further analyze the biodistribution of the virus, we used qPCR amplification of adenoviral E1a in the same tissues and (as a positive control) in lung tissue from the ICOVIR15 + Ad.iC9 group (all analyses normalized against murine GAPDH). As shown in Supplementary Figure S4b, ICOVIR could not be detected in heart, spleen or kidney of treated mice; liver samples of treated mice had a low copy number of E1a as expected by the high tropism of the adenovirus for this tissue, but levels were 11-fold lower than present in the positive control lung.

We evaluated MSC infiltration 3 days after infusion by immunofluorescence using anti-CD90 staining: neither tumors nor MSCs were found in any of the tissues analyzed (Supplementary Figure S5a). Six days after MSC infusion, we assessed the same tissues for iC9-associated CD19 and for activation of the apoptotic pathway after in vivo CID administration using in situ immunofluorescent TUNEL assay. As shown in Supplementary Figure S5b, there was no evidence of iC9-CD19 or apoptosis in normal tissues.

The combination of ICOVIR15 and iC9 improves tumor control and enhances survival of the mice

Finally, we compared the in vivo antitumor activity of each treatment alone and in combination (Figure 5). Tumor bioluminescence was significantly lower in the ICOVIR15-iC9 group after CID administration compared to all other groups (iC9, ICOVIR15 and ICOVIR15-iC9 without CID groups) (Log10 signal change of 9.38; 10.77, P < 0.0001; 10.74, P < 0.0001; 9.8, P = 0.02; and 9.88, P = 0.008, respectively, from day 0 to day 38) (Figure 5a,b). As anticipated, this improved tumor control was associated with enhanced survival. Animals in the ICOVIR15-iC9 + CID group had a median survival of 60 days versus 38, 43, and 45 days in the untreated, iC9 treated, and the ICOVIR15-treated groups, respectively, with maximal survival 82 versus 48 days (P = 0.0003), 50 days (P = 0.002) and 65 days (P = 0.04), respectively (Figure 5c).

Figure 5.

Figure 5

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The combination of ICOVIR15 and iC9 improves tumor control and enhances survival of the mice. (a) SCID-Beige mice were engrafted with FFLuc-GFP labeled A549 cells intravenously (IV) followed by IV infusion of either vehicle or uninfected MSCs, Ad.iC9-MSCs, ICOVIR15-MSCs or ICOVIR15-Ad.iC9-MSCs. Mice received intraperitoneal injections of chemical inducer of dimerization (CID; 50 µg) as described in Methods. Tumor growth was monitored by in vivo imaging to measure bioluminescence. Representative images of bioluminescence in mice show the tumor site and tumor size on days 0 and 38. (b) Total tumor growth after 38 days is represented as log10 signal change. Error bars represent standard deviation. (c) Survival curves of the mice (10 mice per group). MSCs, mesenchymal stromal cells.

Discussion

MSCs can be infected by the combination of ICOVIR15 and Ad.iC9 and efficiently replicate both vectors, leading to amplified viral load and, subsequently, more potent antitumor activity. In the present study, we show that this combined system improves the delivery of target genes to tumor cells compared to the previously described E-MSCs.7 Upon coculture with tumor cells, MSCs release CRAd particles that infect tumor cells and induce apoptosis. MSCs can also successfully transduce tumor cells with the nonreplicating iC9 adenovirus, thereby increasing apoptosis after CID administration in vitro. Our in vivo data show that infected MSCs efficiently and selectively deliver the combination of ICOVIR15 and Ad.iC9 to lung tumor sites significantly improving tumor control, which translates into increased survival.

Combining a replication-defective adenovirus expressing a therapeutic gene with a CRAd may increase the spread of the transgene by propagating the otherwise replication-defective component.13 Such amplification should augment the transduction of adjacent tumor and/or stromal cells with the transgene of interest. To further increase the benefits of combining and replication-defective adenovirus and CRAD, we used two viruses with different fibers that bind to distinct cell receptors.

The use of a vehicle to deliver adenoviral vectors represents a promising strategy to overcome the main limitations of the intravenous use of this therapy. MSCs have several features that make them attractive as a cellular vehicle for cancer therapies: they can be infected efficiently with adenovirus and selectively deliver it to tumor sites, shielding the virus in the bloodstream. MSCs can also target both primary and metastatic sites due to their homing abilities.14 Moreover, the lack of major histocompatibility complex (MHC) class II molecules and low expression of costimulatory molecules allow for the use if MSCs in the allogeneic setting, as several studies already proved in various clinical scenarios.15,16,17,18,19 The delivery of adenoviral-based treatment to tumor sites using MSCs has already been exploited in several tumor models such as glioma, ovarian cancer, melanoma, and breast cancer.20,21,22,23,24 Hakkarainen et al.25 tested MSCs as vehicles of CRAd in a lung cancer model, but they were unable to demonstrate MSC-tumor tropism. The median survival of the treated group in that study remained below 30 days, confirming the aggressiveness and poor response rate of this tumor model.

We have recently shown that MSCs can be engineered to produce an adenoviral vector expressing iC9 and that this approach can be used to kill NSCLC in vitro and in an orthotopic mouse xenograft model.7 This strategy is particularly advantageous in treating lung cancer, since most of the MSCs are entrapped in the lung microvasculature after i.v. administration.11 Moreover, administering the bioinert, nontoxic CID activates iC9, inducing apoptosis irrespective of the cycle status of the cell.26 Therefore, iC9 can affect non-proliferating areas of the tumor, such as resting or hypoxic areas and tumor-associated stroma, both of which are less sensitive to CRAd.27 In our previous NSCLC model, we observed a high heterogeneity in the sensitivity of lung cancer cells to iC9, suggesting the need to further increase its potency in order to improve the antitumor activity. Now, we show that combining iC9 with an oncolytic adenovirus provides an amplification step to the suicide gene and increases cytotoxicity of both the approaches using the single viruses.

Similarly to other groups cited below, our results show that CRAd can efficiently infect MSCs, enabling its replication and providing preliminary amplification of the viral dose. In ICOVIR15, the oncolytic adenovirus we used, an E2F-responsive promoter restricts replication to cells with an activated RB-pathway and controls the replication of early virus genes. This mechanism explains viral replication in MSCs, which contain free E2F. The amount of E2F available in MSCs, however, is consistently inferior to that of tumor cells and the replication ability is inferior as well (data not shown).28 We proved for the first time, to the best of our knowledge, that MSCs can be infected efficiently with a combination of ICOVIR15 (Ad5-RGD) and a first-generation adenoviral vector (Ad5/35), and that this combination strategy significantly improves the replication of the adenoviral vector compared to the previously described E-MSCs, without impairing the replication of the CRAd. As shown by the lower copy numbers of iC9 than E1A 5 days after MSCs infection, however, our results suggest that in the coinfection the replication of the adenoviral vector is less efficient than the replication of the CRAd. Although the mechanism for selective replication remains uncertain, the same phenomenon has recently been reported with the combination of CRAd and helper-dependent adenoviral vectors29 Nonetheless, our dual-approach provides a significant amplification step for the replication-incompetent iC9 vector. Contrary to Hakkarainen et al., we were able to detect selective localization of MSCs at tumor sites within 48–72 hours after administration, allowing them to persist and release the viruses directly to the tumor mass.

An important limitation of the iC9 suicide gene strategy, compared to the more common HSV-TK gene, is the lack of the bystander effect, which limits its action to the first infected tumor cells. Our strategy has the potential to amplify iC9, providing the surrounding tumor cells with the ability to selectively replicate the Ad.iC9 after combined infection with ICOVIR15. As shown by our data, a spread of iC9 is seen in the tumor mass only in the combination treatment, proving the amplification and diffusion of the gene to the surrounding tumor cells. This mechanism enables the elimination of areas of the tumor that are not affected by the oncolytic ICOVIR15 but, expressing iC9, can undergo apoptosis after CID administration.

Our data confirm that the combination treatment enhances the cytotoxic effect on tumor cells as compared to ICOVIR15 alone, leading to a significant improvement of tumor control in this highly aggressive model, which translates into a median survival of 60 days. This effect can be attributed to combination of both oncolytic and apoptotic effects (mediated by ICOVIR15 and iC9, respectively) and also to amplification of both vectors mediated by the ICOVIR 15, which enables multiple rounds of replication of both viruses. Limitations of the animal model used, in which the tumor stroma is of murine origin, make it difficult to assess the likely contribution of ICOVIR15 versus iC9 vectors to the tumor control observed, but the improvement in such control in the mice receiving the vector combination nonethelesas remains significant.

The MSCs' delivery of ICOVIR15 and Ad.iC9 has proven to be safe in our in vivo model. No MSC infiltration, adenoviral cytopathic effect or iC9 expression were recorded on normal organs such as liver, heart, spleen, and kidneys, implying that normal organs are largely spared by viral infection. It is important to point out, however, that mouse models partially reproduce the toxicity that is associated with adenovirus because murine cells have a low ability to replicate adenovirus. Therefore, early phase toxicity, related to the proteins of the capsid, can still occur whereas the late-phase toxicity, dependent upon viral gene transcription, is reduced.30 Moreover, immunocompromised models potentially underestimate the toxicity, as immune antiviral responses often damage healthy tissues. However, using MSCs carrying CRAd was safe in a small clinical case series in which this approach was used to treat metastatic neuroblastoma. Autolimited fever and a mild, transient increase of hepatic alanine aminotransferase were the only toxicities reported.8

Despite the reassuring safety profile, there is concern that MSCs may favor tumor progression and enhance metastases.31 In our model, however, MSCs serve only as vehicles and once they deliver the viruses to the tumor sites, they are also eliminated by the cytopathic effect of the CRAd. As a safety switch, our MSCs also express iC9 and therefore undergo apoptosis upon CID administration.32 Lastly, as our data prove, mice receiving untreated MSCs did not have reduced survival or increased tumor growth compared to untreated mice (Figure 5b,c).

The combination of oncolytic adenovirus and Ad.iC9 has clear benefits for the treatment of NSCLC. However, we can further improve this approach to overcome some of its limitations. First, MSC delivery to metastases beyond lungs is limited because many of the intavenously-infused cells are blocked by the lung microvasculature filter. To improve delivery to secondary sites, we can exploit the technologies of human induced pluripotent stem cells (hiPSC). hiPSCs can be induced to differentiate into MSCs and used as cellular vehicles in their pre-MSCs state, where their smaller size should enable escape from the pulmonary filter. Although our approach showed significant improvement in survival, we did not reach the ultimate goal of tumor eradication; therefore, the strategy must be further improved. The induction of an immune response against adenovirus-infected tumor cells is considered essential for oncolytic virotherapy to yield relevant and sustained antitumor activity.3,33 Therefore, including an immunostimulatory signal could improve the antitumor activity of this approach.

In conclusion, this is the first study demonstrating the feasibility and effectiveness of delivering a combination of ICOVIR15 and Ad.iC9 through MSCs for NSCLC, due in part to the strong amplification of Ad.iC9 by coinfected MSCs and tumor cells. The combined treatment confers better tumor control in vivo, which translates into prolonged survival in mice in the very aggressive NSCLC tumor model.

Materials and Methods

Cloning. The construction of the iC9 suicide cassette was previously described.34 Briefly, iC9 was comprised of the catalytic domains of caspase-9 fused to FV (F36V), a modified FK506 binding protein (FKBP). The caspase recruitment domain (CARD) of the caspase-9 gene was replaced with a 12-kDa human FK506 binding domain (FKBP12; GenBank AH002 818) that allows dimerization of the caspase-9 and activation of the apoptotic pathway after exposure to the FK506 analog AP20187. This gene was fused via a 2A-derived nucleotide sequence to truncated human CD19 (iC9-ΔCD19). As previously demonstrated, fusion of FKBP to caspase-9 enables conditional dimerization and activation of the caspase by addition of a CID/AP20187, a ligand containing two FK506 moieties, to the cellular medium.35 Recombinant Adenovirus 5F35 was constructed using the Adeno-XTM Expression System 1 from Clontech. Briefly, the iC9-ΔCD19 construct was amplified by PCR and inserted into the pShuttle 2 plasmid. The recombinant expression cassette was then digested using I-Ceu I and PI-Sce I and subcloned into the Adeno-XTM genome. The vector encoding the fusion protein eGFP-Fireflyluciferase (eGFP-FFLuc) was previously described.36

Virus production. The ICOVIR15 has been previously described.12 The viral particle (VP) to (pfu) ratios for ICOVIR15 was 38. Ad.iC9 was produced by Baylor College of Medicine's (BCM) Vector Development Lab following standard operating procedures (www.bcm.edu/vector). Ad.GFP vector was purchased by BCM Vector Development Lab.

Generation of MSCs. MSCs were isolated from healthy donors (following a standard BCM Institutional Review Board approved protocol) as described previously.31 Discarded (postinfusion) healthy donor bone marrow collection bags and filters were washed with RPMI 1640 and plated on tissue culture flasks in RPMI with 10% fetal bovine serum (FBS), 2 mmol/l L-glutamine, 100 units/ml penicillin, and 100 mg/ml streptomycin (Invitrogen). After 48 hours, the supernatant was discarded and the cells were cultured in complete culture medium: α-minimum essential medium (Invitrogen) with 16.5% FBS, 2 mmol/l L-glutamine and 100 mg/ml streptomycin. Cells were grown to less than 80% confluency and replated at a lower density. The bone marrow-derived MSC used in the present study exhibited a spindle shape, expressed MSC-defining markers measured by flow cytometry analysis, and differentiated into adipocytes, osteocytes, and chondrocytes in appropriate culture media (data not shown).

Cell culture. Two human non-small-cell lung cancer cell lines (H1299 and A549) were obtained from American Type Culture Collection. H1299 were cultured in RPMI 1640 and A549 were cultured in Dulbecco's modified Eagle's medium. All media was supplemented with 10% fetal bovine serum, 2 mmol/l L-glutamine and 100 mg/ml streptomycin. Cells were grown at 37 °C in a humidified incubator at 5% CO2.

Adenoviral transduction and loading into MSCs. Cells were plated in a six-well tissue culture treated plate (2 × 105 cells/well) and cultured in corresponding complete medium for 24 hours. Medium was aspirated and replaced with 350 µl serum-free Iscove's Modified Dulbecco's Medium per well. ICOVIR15 50 vp/cell and/or Ad.iC9 1,000V vp/cell were added and plates were rocked every 10 minutes for 1 hour and every 20 minutes for the following 2 hours. Next, 650 mcl of corresponding complete medium was added per well. After 24 hours, cells were washed thrice with PBS and 2 ml of fresh complete medium was added per well. To further transduce tumor cells with MSC-produced viruses, supernatant from MSCs infected as described was collected 5 days after infection and transferred to A549 and H1299 at sequential dilution. To prepare MSCs for the in vivo infusion, the same protocol has been followed, loading 50 vp/cell of ICOVIR15 and 1,000 vp/cell of Ad.iC9. After 24 hours, cells were washed thrice with PBS, collected, and resuspended in 200 mcl of PBS for the infusion.

Flow cytometry. Cells were collected at a specific time point and stained with CD19 APC (Beckman Coulter) and CD90 FITC (BD Biosciences) antibodies according to the manufacturer's instructions. The percentage of cells expressing GFP, CD19, or CD90 and the mean fluorescence intensity was measured using Gallios (Beckman Coulter) and analyzed using the Kaluza Flow analysis software (Beckman Coulter).

Viral production assay. MSC cell cultures (2 × 105 cells seeded in six-well plates) were infected with ICOVIR15 alone or in combination with iC9 as described above. Five days after infection, the supernatant was collected and the viral titer was determined using the median tissue culture infective dose method (TCID50).

Quantitative PCR analysis of viral DNA. A qPCR was performed to determine the copy number of the E1A and the iC9 genes in MSCs and mice lung tissues, as previously described.29 Briefly, MSCs were plated and infected with ICOVIR15 (50 vp/cell) alone or in combination with Ad.iC9 (1,000 vp/cell), as described above. After 5 days, cells were collected. DNA was extracted from each sample using QIAamp DNA mini kit (QIAGEN), as per manufacturer's protocol. To determine the copy number of E1A in mice lung tissues, lungs were collected and DNA was extracted using QIAamp DNA mini kit (QIAGEN), as per manufacturer's protocol. DNA samples were analyzed by quantitative real-time PCR using a Bio-Rad iQ5 Real-time PCR Detection system (Bio-Rad) and Applied Biosystems SYBR green PCR master mix (Life Technologies). The specific primers targeting the iC9 or the E1A genes are shown in the Table 1. The gene dosages were then normalized to that of GAPDH.

Table 1. Primer sets information.

graphic file with name mt2015110t1.jpg

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Reagents. CID (B/B homodimerizer) was purchased from Clontech and diluted in 100% Ethanol.

Measurement of apoptosis. Cells were treated as indicated, trypsinized, and collected by centrifugation. Cells were stained with Annexin-V PE and 7-amino-actinomycin D (7-AAD) for 15 minutes according to the manufacturer's instructions (BD Biosciences). Annexin V/7-AAD-positive cells were quantified by flow cytometry (Gallios, Beckman Coulter) and analyzed with Kaluza software (Beckman Coulter).

Xenograft mouse model. All mouse experiments followed BCM animal husbandry guidelines. Seven-week-old severe combined immunodeficient female BEIGE mice (CB17-SCID; Charles River) were inoculated via tail vein injection with 2.5 × 106 A549 cells labeled with firefly luciferase (FFLuc). Tumor engraftment was measured using the Xenogen-IVIS Imaging System (Caliper Life Sciences, Hopkinton, MA). Briefly, mice were injected intraperitoneally (i.p.) twice a week with D-luciferin (150 mg/kg), and signal intensity was measured as total photon/sec/cm2/sr (p/s/cm2/sr). Four days after tumor establishment, engrafted mice were injected with either vehicle or 1 × 106 intravenous uninfected MSCs, Ad.iC9-MSCs, ICOVIR15-MSCs, or ICOVIR15-Ad.iC9-MSCs. After 4, 5, and 7 days, intraperitoneal CID (50 µg) was administered. The whole treatment cycle (MSCs and CID) was repeated twice. Mice were euthanized when signs of discomfort (more than 20% weight loss, immobility, huddled posture, inability to eat, ruffled fur, self mutilation, vocalization, hypothermia) were detected by the investigator or as recommended by the veterinarian who monitored the mice three times a week.

Immunofluorescence staining and immunohistochemistry. Murine tissue (lungs, heart, liver, spleen, and kidneys) was collected and fixed in 4% formaldehyde for 24 hours at room temperature (for paraffin embedding and hematoxylyn/eosin or immunohistochemistry staining) or frozen in optimal cutting temperature (Sakura Finetek, Zoeterwoude, Netherlands). Human CD90 and CD19 immunodetection was performed by incubating optimal cutting temperature-embedded tissue sections with a primary polyclonal mouse anti-hCD90 antibody (Calbiochem, CP28) or a rabbit anti-hCD19 antibody (Abcam, ab134114) and a secondary AlexaFluor594-labeled donkey anti-mouse antibody (Jackson ImmunoResearch, 715-585-150) AlexaFluor 594-labeled donkey anti-rabbit antibody (Jackson ImmunoResearch, 711-585-152). Adenoviral capsid proteins immunodetection was performed with the polyclonal rabbit anti-adenovirus antibody (Abcam, ab6982) and a secondary AlexaFluor 594-labeled donkey anti-rabbit antibody (Jackson ImmunoResearch, 711-585-152). Slides were counterstained and mounted with ProLong Gold antifade reagent with 4,6-diamidino-2-phentlindole (Life Technologies) and visualized under a spinning disk confocal microscope. Imaging data were acquired and analyzed using Zen software (Zeiss). Apoptosis detection was performed by direct TUNEL assay using the ApopTag Fluorescein Direct In Situ Apoptosis Detection Kit (Millipore), following manufacturer's protocol.

Statistical analysis. All statistical analyses used GraphPad Prism 4.0 software (GraphPad Software). Paired two-tailed Student's t-test was used to determine the statistical significance of differences between samples. All numerical data are represented as mean ± standard deviation. Results were considered statistically significant when P < 0.05.

SUPPLEMENTARY MATERIAL Figure S1. I15-MSCs are efficiently co-infected and express higher levels of E1A than E1A-MSCs. Figure S2. MSCs persists at tumor site and deliver ICOVIR15 (additional pictures). Figure S3. Tumor is transduced by iC9 and undergoes apoptosis (additional pictures). Figure S4. Normal tissues do not show morphologic signs of adenoviral toxic effects. Figure S5. MSCs cannot be found in normal tissues and no evidence of virus replication and iC9 transduction in normal tissue is revealed.

Acknowledgments

The authors thank Catherine Gillespie for the editing of the paper and Jeffrey Howard for his help in the set up of immunofluorescent stainings. This work was supported by the Cancer Prevention Research Institute of Texas (RP110553 P1 to M.K.B.) and the National Institutes of Health (R00HL098692 to M.S.).

Supplementary Material

Supplementary Figures
Click here for additional data file. (1MB, pdf)

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