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Angiogenic properties of human placenta-derived adherent cells and efficacy in hindlimb ischemia
Angiogenic properties of human placenta-derived adherent cells and efficacy in hindlimb ischemia Aleksandar Francki, PhD, Kristen Labazzo, PhD, Shuyang He, PhD, Ellen Z. Baum, PhD, Stewart E. Abbot, PhD, Uri Herzberg, DVM, PhD, MBA, Wolfgang Hofgartner, MD, DSc, and Robert Hariri, MD, PhD, on behalf of the Celgene Cellular Therapeutics Research Group, Warren, NJ Objective: Human placenta-derived adherent cells (PDACs) are a culture-expanded, undifferentiated mesenchymal-like population from full-term placental tissue and were previously shown to possess anti-inflammatory and immunomodulatory properties. PDACs (formulated as PDA-002) are in clinical trials for peripheral arterial disease with diabetic foot ulcer. In the current study, we examined their angiogenic and tissue reparative properties. Methods: The effects of PDACs on survival and tube formation of human umbilical vein endothelial cells (HUVECs) were tested using conditioned media and noncontact coculture. Angiogenic effects were assessed in the chick chorioallantoic membrane assay. Hindlimb ischemia (HLI) was induced in mice and rats by femoral artery transection, and blood flow and blood vessel density were monitored in vivo by laser Doppler and angiography in the ischemic and control limbs. Tissue damage and regeneration in HLI were examined in histologic sections of quadriceps muscle stained with hematoxylin and eosin, and newly synthesized blood vessels were detected by indoxyl-tetrazolium staining for alkaline phosphatase. Results: PDACs enhanced the survival of serum-starved HUVECs and stimulated HUVEC tube formation, and in the chick chorioallantoic membrane assay, PDACs stimulated blood vessel formation. In HLI, intramuscular administration of PDACs resulted in improved blood flow and vascular density, and in quadriceps muscle, tissue regeneration and increased numbers of blood vessels were observed. Conclusions: PDACs exhibited various activities consistent with angiogenesis and tissue repair, supporting the continued investigation of this cell therapy as treatment for vascular disease-related indications. (J Vasc Surg 2015;-:1-11.) Clinical Relevance: Peripheral arterial disease with diabetic foot ulcer is a major complication in patients with diabetes and is estimated to lead to w80,000 lower extremity amputations each year. A significant need exists for new treatment options that will promote revascularization and healing of damaged tissue. This report discusses the angiogenic and tissue reparative properties of PDA-002, a culture-expanded mesenchymal-like cell population derived from human placenta that is currently in clinical trials for peripheral arterial disease with diabetic foot ulcer (NCT01859117).
Angiogenesis is a complex process that facilitates blood vessel formation and tissue perfusion. It is associated with a balance of stimulatory and inhibitory steps, including secretion of various soluble factors, degradation of extracellular matrix, and proliferation and differentiation of endothelial cells into tubelike vessels.1 The term therapeutic angiogenesis refers to situations in which angiogenic stimulation is a From the Research & Development Department, Celgene Cellular Therapeutics. The Appendix (online only) lists the names of the Celgene Cellular Therapeutics Research Group members. Author conflict of interest: All authors except I.L. are employees of Celgene and hold stock or stock options in Celgene. I.L. indicates no potential conflicts of interest and received funding from Celgene Cellular Therapeutics through a contract service agreement for conducting the in vivo studies. Additional material for this article may be found online at www.jvascsurg.org. Reprint requests: Aleksandar Francki, PhD, Research & Development Department, Celgene Cellular Therapeutics, 7 Powderhorn Dr, Warren, NJ 07059 (e-mail: [email protected]). The editors and reviewers of this article have no relevant financial relationships to disclose per the JVS policy that requires reviewers to decline review of any manuscript for which they may have a conflict of interest. 0741-5214 Copyright Ó 2015 by the Society for Vascular Surgery. Published by Elsevier Inc. http://dx.doi.org/10.1016/j.jvs.2015.04.387
clinically desirable outcome (eg, tissue repair or treatment of local hypovascularity).2 To date, no small-molecule or biologic-based approach for therapeutic angiogenesis has proved consistently efficacious, and several compounds that showed promise in preclinical and early clinical trials failed to demonstrate efficacy in larger randomized trials.3 These failures may reflect the fact that angiogenesis and tissue repair are not regulated by a single molecule or biologic pathway but by an intricate, temporally coordinated network of biologic processes. It has been suggested that cell therapy using mesenchymal stromal cells (MSCs) or mesenchymal-like cells available from a variety of sources, such as bone marrow and placenta, can promote angiogenesis and tissue regeneration through multiple biologic activities, offering a potentially transformational means to treat diseases with significant unmet clinical needs.4 These cell therapies have been shown to modulate pathologic immune responses and inflammatory processes5 as well as the secretion of specific trophic factors that stimulate resident endothelial cells or progenitors to initiate angiogenic processes in injured tissue.6 Placenta-derived adherent cells (PDACs) are a cultureexpanded, undifferentiated mesenchymal-like population derived from full-term placental tissue.7 The cells have 1
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the genotype of the newborn and display features characteristic of MSCs, but with a unique phenotype associated with their placental origin. The cells are anchorage dependent in culture and adhere to plastic during expansion in vitro.7 PDAC-mediated activities have been studied in vitro, in animal models, and in clinical trials and display immunomodulatory, anti-inflammatory, proregenerative, and neuroprotective properties,7-10 suggesting that the cells may have potential utility across a variety of disease indications. PDA-002, an intramuscular formulation of PDACs, is in clinical development for the treatment of peripheral arterial disease with diabetic foot ulcer (http:// clinicaltrials.gov/show/NCT01859117). In this study, we investigated whether in addition to previously described immunomodulatory capabilities, PDACs display angiogenic properties in various assays and animal models relevant to angiogenesis. We show that PDACs secrete a variety of proangiogenic factors (most prominently hepatocyte growth factor [HGF]), support endothelial cell survival, induce blood vessel formation, sustain blood vessel growth and maintenance, and induce muscle repair in rodent hindlimb ischemia (HLI) models, demonstrating significant proangiogenic and reparative functions. These results support the continued investigation of PDACs as cell therapy for vascular disease-related indications. METHODS Preparation of PDACs, human umbilical vein endothelial cells (HUVECs), human dermal fibroblast (HDF), and human MSCs. PDACs were prepared by mechanical and enzymatic digestion of human placental tissue of newborn origin obtained from a normal, full-term birth as described.7 PDACs were expanded until passage 6 in PDAC medium7 and subsequently used as described in the Supplementary Methods (online only). HUVECs, HDF, and human bone marrow-derived MSCs were obtained from Lonza (Walkersville, Md) and culture expanded according to the manufacturer’s instructions. Flow cytometric analysis of PDACs. Flow cytometric surface marker staining of PDACs was performed using labeled monoclonal mouse antihuman immunoglobulin G (BD Biosciences, San Jose, Calif), and expression was analyzed as described in the Supplementary Methods (online only). Preparation of cell-conditioned medium (CM). PDACs, HDF, or MSCs were seeded onto tissue culturetreated polystyrene 24-well plates (Corning, Lowell, Mass) at 4000 cells/cm2 for overnight establishment in PDAC medium before incubation in serum-free PDAC medium for 48 hours (37 C, 5% CO2, 21% O2, 90% relative humidity). CM was then collected and frozen at 80 C. HUVEC survival assay. HUVECs were subjected to serum starvation in the presence and absence of PDACs, HDF, or MSCs, using indirect coculture systems as detailed in the Supplementary Methods (online only).
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After 24 hours, the Transwell (Corning) cocultures and media were removed, and HUVEC counts were determined by the CellTiter-Glo assay (Promega, Madison, Wisc). Tube formation assay. HUVECs were cultured to passage 3 in endothelial cell growth medium 2 for 3 days to w75% confluency. HUVECs were washed and resuspended in Dulbecco’s modified Eagle medium (DMEM) and diluted to 4000 cells/mL with human placental collagen (1.5 mg/mL in DMEM), and the tube formation assay was performed as described in the Supplementary Methods (online only). After incubation (24 hours), HUVEC drops were stained with a Diff-Quik Cell Staining Kit (Dade Behring, Inc, Newark, Del). Images of each well were acquired with the Zeiss SteREO Discovery V8 microscope (Carl Zeiss AG, Jena, Germany) and analyzed for average network area. Quantitation of secreted factors. The concentrations of angiogenic factors basic fibroblast growth factor 2 (FGF-2), granulocyte colony-stimulating factor (G-CSF), interleukin-8 (IL-8), platelet-derived growth factor BB (PDGF-BB), vascular endothelial growth factor (VEGF), interleukin-6 (IL-6), and monocyte chemotactic protein 1 (MCP-1) were analyzed in PDAC-CM using Bio-Plex Pro multiplex bead array systems (Bio-Rad Laboratories, Hercules, Calif). HGF concentration was determined using a solid-phase sandwich enzyme-linked immunosorbent assay system (Quantikine Human HGF Immunoassay; R&D Systems, Minneapolis, Minn). Chicken embryo chorioallantoic membrane (CAM) assay. CAM assays in fertilized chicken eggs were used to assess neovascularization and were performed by SRI International (Menlo Park, Calif) as described11 and detailed in the Supplementary Methods (online only). Animal care. All animal work was performed at Pharmaseed Ltd (Ness Ziona, Israel) using adult Balb/c mice (w25 g) or Sprague-Dawley rats (w200 g) according to the National Institutes of Health and the Association for Assessment and Accreditation of Laboratory Animal Care. Unilateral HLI induction and administration of PDACs. The surgical procedure followed the Prox-A protocol12 as detailed in the Supplementary Methods (online only). At 24 hours after surgery, each animal was treated with PDACs, recombinant human VEGF 165 (3.3 mg/animal; Sigma-Aldrich, St. Louis, Mo), or vehicle (cell freezing media) by intramuscular injection proximal and distal to the surgical wound. Mice were injected with 25 mL in each of two sites (total 50 mL); rats were injected with 50 mL in each of four sites (total 200 mL). PDAC doses were w4 104, 4 105, or 4 106 cells/kg. Luciferase assay for PDAC retention. PDACs constitutively expressing luciferase were prepared with the Cignal lenti positive control (luc) system (SABiosciences, Valencia, Calif), and 1 106 cells (in 25 mL) were injected intramuscularly into the ischemic hindlimb of Balb/c mice. Luciferin substrate (150 mg/kg; Caliper Life Sciences, Hopkinton, Mas) was administered intravenously, and
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luciferin intensity was measured in the whole animal with an in vivo real-time imaging system (IVIS Spectrum 200; Caliper Life Sciences). Laser Doppler analysis. Blood flow in surgical and control (nonsurgical) hindlimbs was measured with a noncontact laser Doppler before surgery and on days 1, 7, 15, 21, 28, and 35 for both mice and rats. In addition, mice were measured at 42 and 49 days. Blood flow in the nonischemic control limb is defined as 100%; measurements are expressed as percentage flow in ischemic limb compared with control limb. Angiography of rat hindlimbs. On day 35 after surgery, half of the animals in each group were anesthetized with ketamine/xylazine. Barium sulfate solution (2.5 mL) was infused into the infrarenal aorta after ligation of the proximal aorta and inferior vena cava, and the number of contrast-filled blood vessels was determined by X-ray roentgen assays. Quantitation of intersections between contrast-filled vessels and gridlines of the angiographs is presented as an angioscore, defined as the number of blocks on a grid filled by radiopaque dye, and was performed in blinded fashion. Histology of hindlimb muscle. After sacrifice, quadriceps muscles of ischemic and control limbs were removed, weighed, fixed in Hepes-glutamic acid buffer mediated Organic solvent Protection Effect (HOPE) fixative (DCS Innovative, Hamburg, Germany), and embedded in paraffin. Paraffin blocks were cooled to 20 C for 30 minutes and sectioned with a microtome at 5 mm thickness. Muscle anatomy was visualized by staining two serial sections with hematoxylin and eosin and evaluated microscopically for morphologic changes. Quantitation of blood vessel density in muscle sections. The indoxyl-tetrazolium method for alkaline phosphatase was used to detect capillary endothelial cells in newly synthesized vessels. Staining was performed using the BCIP/NBT kit from Vector Labs (Burlingame, Calif; catalog No. SK-5400), and nuclei were counterstained with methyl green (Vector Labs; catalog No. H-3402). Muscle capillary density was evaluated in blinded fashion in sections from three representative animals by counting positively stained capillaries within three randomly chosen fields per section at 20 magnification. RESULTS Flow cytometric characterization of PDACs. As previously described,7 PDACs expressed surface markers that are typical of MSCs, including CD90, CD73, and CD105, as well as CD200, which is associated with placental cells but absent on other MSC-like cells, thereby distinguishing PDACs from MSCs. Further immunophenotypic characterization showed that PDACs do not express CD31, KDR, CD133, VE cadherin (CD144), and Tie2. In conjunction with the described lack of CD34 expression, these findings confirmed the absence of endothelial progenitor cells and mature endothelial cells in PDAC preparations.
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PDACs enhance the survival of endothelial cells. The effect of PDACs on the survival of serum-starved HUVECs was examined in a noncontact coculture system. After serum starvation and 24 hours of indirect coculture of the two cell types, only 35% of the cell count of HUVECs remained in the absence of PDACs (Fig 1, A, serum-free DMEM). In contrast, indirect coculture with PDACs resulted in 72% HUVEC survival. Although we refer to the observed increase in the HUVEC count in the presence of PDACs compared with control as “enhanced survival (reduced cell death),” enhanced HUVEC proliferation may also be a component. Other cell types secrete factors that may render them active in HUVEC survival, tube formation, and CAM assays; these in vitro and ex vivo angiogenic activities are not unique to PDACs, HDF, or MSCs. To address potential differences between the aforementioned cell types and PDACs, we tested HDF and MSCs in the HUVEC survival assay under equal experimental conditions. All three tested cell types showed an effect that was significantly higher in comparison to the control medium; however, the effect of PDACs and the CM was consistently 20% higher compared with the effects of HDF and MSCs and their respective CM (data not shown). These results support that PDACs have unique in vitro effects on the angiogenic functions of human endothelial cells compared with other relevant cell types. Dose-response studies were performed to determine the ratio of PDACs to HUVECs required to promote HUVEC survival (Fig 1, B). A range of PDACs (0-40,000) were used, whereas the input HUVECs were kept constant at w50,000 cells (PDAC:HUVEC of 0.1:1 to 0.8:1). When cultured in DMEM alone, the HUVEC count decreased by w60%, to 20,000 cells. As few as 5000 PDACs had a statistically significant effect on the HUVEC count (P < .01), with a plateau observed at approximately 20,000 PDACs, resulting in twofold higher final HUVEC count compared with the DMEM control (P < .01). Based on the starting HUVEC count, PDACs exerted a beneficial effect on HUVEC survival at as little as 0.1:1, with a plateau observed at w0.4:1 PDAC:HUVEC. The lack of direct contact between PDACs and HUVECs in these experiments suggests that the effect is mediated through factors elaborated by PDACs. PDAC-CM stimulates endothelial cell tube formation. The ability of CM from PDACs to promote HUVEC tube formation was examined by the application of PDAC-CM to HUVECs cultured within a threedimensional environment of collagen. After 24 hours of incubation, tube formation (as measured by average network area) was w70% greater for CM than for serumfree media (P < .01; Fig 2). Trophic factor protein secretion of PDACs. Because direct PDAC contact with HUVECs was not required to stimulate HUVEC survival or tube formation (Figs 1 and 2), it was of interest to examine protein secretion by PDACs. Assessment of eight proteins (Fig 3) revealed that PDACs secrete several mitogens and cytokines
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Fig 1. A, Effect of placenta-derived adherent cells (PDACs) on human umbilical vein endothelial cell (HUVEC) survival. HUVECs were exposed to PDACs through a Transwell (PDAC:HUVEC, w2:1). The initial value of HUVECs (after 6-hour serum starvation in endothelial cell basal medium and before Transwell exposure) was set to 100%. The difference between HUVEC survival in serum-free Dulbecco’s modified Eagle medium (SF-DMEM) overnight (w35%) and in the presence of PDACs (72%) was statistically significant (P < .01; n ¼ 7). B, Dosedependent effect of PDACs on HUVEC survival (n ¼ 3). The error bars indicate standard deviation.
relevant to angiogenesis. FGF-2, G-CSF, IL-8, and PDGF-BB were secreted in low but detectable levels (<100 pg/mL); VEGF and IL-6 were secreted at moderate levels (140 and 480 pg/mL, respectively). MCP-1 and HGF were secreted at substantially higher levels (1200 and 1600 pg/mL, respectively). Ex vivo proangiogenic effects of PDACs. To investigate the effect of PDACs on vascularization in an ex vivo model, PDACs were cultured on exposed CAM of chick embryos, and the change in blood vessel density was assessed. As shown in Fig 4, PDACs caused a substantial increase (w5- to 10-fold) in blood vessel density compared with medium and vehicle controls and was indistinguishable from positive control FGF-2, a known proangiogenic factor. PDAC administration increases blood flow in rodent models of HLI. The proangiogenic effects of PDACs led us to test their efficacy in rodent models of HLI. After induction of HLI, blood flow was measured by laser Doppler with and without intramuscular administration of PDACs. Throughout the mouse study (Fig 5, A), an increase in blood flow was observed in all animal groups treated with PDACs compared with vehicle-treated
control. The improvement in blood flow was visible from day 28 and continued to increase through day 49 after surgery, similar to the positive control, VEGF. The increase in blood flow in all PDAC treatment groups compared with vehicle control was statistically significant (from day 28 through day 49; P < .01). In a rat study, an increase in blood flow was observed in all animal groups treated with PDACs (4 104, 4 105, and 4 106 cells/kg) compared with the vehicle-treated control (Fig 5, B). This improvement was observed on day 35 after HLI initiation in all PDAC-treated groups and was statistically significant compared with vehicle control (P < .001 for day 35 at all PDAC doses). To determine whether PDAC viability is required for the observed increase in blood flow, nonviable cells were prepared by repeated freeze-thaw cycles before administration in the rat HLI model; nonviable PDACs failed to improve blood flow. Because rodents subjected to HLI continued to experience improvement in blood flow for an extended time (49 days after intramuscular administration of PDACs into leg muscle), it was of interest to examine the in vivo location and retention time of PDACs using luciferin
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Fig 2. Stimulation of human umbilical vein endothelial cell (HUVEC) network tube formation by placenta-derived adherent cell (PDAC)-conditioned medium (CM). Representative photomicrographs of PDAC-induced (upper panel) and serum-free Dulbecco’s modified Eagle medium (DMEM) control (lower panel) network (tube) formation are shown. The arrows depict representative tubes.
substrate and PDACs expressing luciferase in a mouse HLI model. As shown in Fig 5, C, the PDAC signal was detectable at 4, 24, and 72 hours after injection only at the leg injection sites, but not in any major organs (eg, brain, heart, lung, liver, kidneys; data not shown). However, no luciferin signal was detected at the injection site or elsewhere in the animals after 8 days, suggesting only short-term persistence of PDACs in the mouse leg and no migration or integration of the cells in the treated animals. Thus, recovery of blood flow in PDAC-treated mice continued for several weeks after the administered cells were no longer detectable. Thus, in both mouse and rat models of HLI, intramuscular injection of PDACs was effective in restoring blood flow to the ischemic limb beyond the measured persistence of PDACs. Angiography assessment of rat hindlimbs. To further investigate the restoration of blood flow observed in the rat HLI model, blood vessel density was examined by angiography of ischemic limb from rats with and without PDAC treatment, 35 days after induction of ischemia (Fig 6). By visual inspection, higher blood vessel density was observed on PDAC treatment (Fig 6, B) compared with the vehicle control (Fig 6, A) and is
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quantified in Fig 6, C. PDAC treatment resulted in a twofold increase in angioscore compared with vehicle control (P < .01 or .05 for day 35 at both PDAC doses). Histology of ischemic limb muscle. To determine the effect of PDACs on the injured muscle tissue in the HLI models, hematoxylin and eosin staining was performed on representative tissue sections, allowing optimal visualization of potential repair of the injured muscle. Muscle fibers in mouse quadriceps muscle from ischemic and healthy hindlimbs were examined (Fig 7). In nonischemic limb, muscle fibers had a polygonal appearance with relatively narrow borders of connective tissue (Fig 7, A). In ischemic limb (Fig 7, B), the myofibers appeared smaller and irregular, with correspondingly larger borders of surrounding connective tissue, and the muscle tissue frequently showed inflammatory infiltrates adjacent to blood vessels. Treatment with PDACs (Fig 7, D) led to the reduction of inflammatory infiltrates and marked improvement in the structure of muscle fibers, with an appearance similar to uninjured muscle in the nonischemic limb (Fig 7, A), but with myofibers containing multiple, centrally located nuclei, indicative of tissue regeneration. PDACs had an effect similar to the VEGF control (compare Fig 7, C and D); however, PDAC treatment resulted in an apparent further reduction of interspersing connective tissue. The effects of PDAC treatment on rat muscle histology were similar (data not shown). Murine blood vessel density measurement. Vascular structures in PDAC-treated mice were visualized (Fig 8, A) and quantitated (Fig 8, B), using VEGF as the positive control. PDAC treatment (4 104/kg or 4 105/kg) resulted in markedly higher numbers of all sizes of blood vessels (capillaries, 50-100 and 300-700 mm2; arterioles, 300-700 mm2; and larger blood vessels, >700 mm2) in comparison to vehicle control. VEGF was similar to PDAC treatment in vessel density in the size ranges #700 mm2, but for larger vessels (>700 mm2), PDAC treatment resulted in threefold higher density compared with VEGF. For smaller vessels (#300 mm2), VEGF and PDAC treatment each exceeded the vessel density of the healthy control limb. Thus, treatment with PDACs demonstrated not only muscle regeneration at the level of muscle fibers but also an increase in the number of blood vessels of all sizes examined, relative to vehicle-treated mice. Finally, as stated before, other cell types secrete factors that may render them active in the HLI model; these in vivo activities are not unique to PDACs, MSCs, or even fibroblasts. Therefore, we deemed the use of PDACs alone as well as the use of dead PDACs in the HLI in vivo studies appropriate. However, to address potential differences between the aforementioned cell types and to exclude nonspecific responses related to cross-species transfer of cells into an injury model, we tested HDF at equal cell numbers in the in vivo HLI models. In both mouse and rat in vivo models, HDF failed to restore blood flow or to induce repair of muscle tissue in the ischemic limb (data not shown). Thus, the angiogenic and tissue reparative effects of PDACs observed in rodent HLI are not
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Fig 3. Quantitation of factors secreted by placenta-derived adherent cells (PDACs). Culture supernatants were analyzed as described for the indicated set of secreted factors. Of the factors examined, hepatocyte growth factor (HGF) is the most abundant, followed by monocyte chemotactic protein 1 (MCP-1) and interleukin-6 (IL-6; n ¼ 5). The error bars indicate standard deviation. FGF-2, Fibroblast growth factor 2; G-CSF, granulocyte colony-stimulating factor; IL-8, interleukin-8; PDGF-BB, platelet-derived growth factor BB; VEGF, vascular endothelial growth factor.
Fig 4. Placenta-derived adherent cells (PDACs) promote vascularization. Percentage change of blood vessel (BV) density was measured in the chorioallantoic membrane (CAM) assay. PDACinduced vascularization of the CAM was statistically indistinguishable from that of the positive control fibroblast growth factor 2 (FGF-2) and significantly greater than the BV density induced by either the vehicle control (P < .01) or the basal medium (P < .01; n ¼ 7). The error bars indicate standard deviation.
driven by a nonspecific xenogeneic inflammatory response to human cells. DISCUSSION Human PDACs are a novel MSC-like population previously shown to possess potent immunomodulatory, anti-inflammatory, neuroprotective, and proregenerative properties.7-9,13 In the current study, we explored the angiogenic properties of PDACs. In vitro, PDACs exhibited various activities consistent with angiogenesis, including enhancement of endothelial cell survival under conditions of serum starvation and stimulation of tube formation. PDACs also promoted vascularization in the CAM
assay. Importantly, in clinically relevant disease models of HLI in both rats and mice, intramuscular injection of PDACs resulted in increased blood flow, increased blood vessel density, and repair of damaged tissue. These studies support continued investigation of PDACs as an angiogenic therapeutic. PDACs (formulated as PDA-002 for local injection) are currently in clinical trials for peripheral arterial disease with diabetic foot ulcer. It is becoming evident that treatment of certain vascular diseases with a single factor (eg, FGF, VEGF, or HGF) may have limited efficacy14,15 and may not be sufficient to ameliorate complex pathologic processes with parallel excessive inflammation or tissue degeneration. In contrast to single agents, MSCs and mesenchymal-like cells have been referred to as an “injury drugstore,”16 possessing immunomodulatory, anti-inflammatory, proregenerative, and angiogenic activities and acting in a paracrine fashion to provide trophic support, often without clear evidence of engraftment.17 Mesenchymal-like cells from a variety of sources can promote angiogenesis and tissue regeneration through a wide variety of biologic activities, offering a potentially transformational means of treating diseases that have significant unmet clinical needs.18-21 Caplan and Correa16 posit that MSCs are able to “establish a regenerative microenvironment by secreting bioactive molecules and regulating the local immune response.” Our experiments demonstrate PDAC multifunctionality consistent with that viewpoint. In particular, our experiments using CM from PDACs or noncontact coculture of HUVECs and PDACs strongly suggest that soluble factors secreted by PDACs mediate effects on endothelial cells including cell survival (or proliferation) and tube formation, consistent with results from mesenchymal-like cells from other sources.16,17,20 Angiogenic factors detected in
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Fig 5. A and B, Placenta-derived adherent cell (PDAC) administration increases blood flow in rodent models of hindlimb ischemia (HLI). Blood flow was measured by noncontact laser Doppler at the indicated times after surgery. PDACs were administered at the indicated doses. Blood flow to the undamaged limb is defined as 100%. Representative experiments from duplicate studies are shown. A, Mouse model. The increase in blood flow in all PDAC treatment groups compared with the vehicle control was statistically significant (from day 28 through day 49; P < .01; n ¼ 15). The error bars indicate standard deviation. B, Rat model. At 35 days, the difference in blood flow between PDACtreated groups and the vehicle control group is statistically significant (P < .001). Nonviable PDAC control was statistically indistinguishable from the vehicle control (n ¼ 12). The error bars indicate standard deviation. C, Representative in vivo real-time images at different time points of four mice injected in the ischemic hindlimb with PDACs expressing the luciferase reporter gene; mouse No. 5 received vehicle. PDACs persisted <8 days in mice. ROI, Region of interest; VEGF, vascular endothelial growth factor.
media supernatants from cultured PDACs included FGF-2, G-CSF, IL-8, PDGF-BB, VEGF, IL-6, MCP-1, and most prominently HGF. HGF plays a major role in embryonic development, adult organ regeneration, and wound healing.22 Furthermore, HGF is central to angiogenesis and tissue regeneration23 and is thought to exert positive effects during the repair of vascular pathologic processes, including stroke.23,24 The role of HGF in angiogenesis includes not
only induction of new blood vessel formation but also maturation of nascent blood vessels, suggesting effects distinct from VEGF25 and confirming the requirement for multiple soluble factors to drive the angiogenic process. The lack of significant success of HGF monotherapy in clinical trials for critical limb ischemia26-28 also points to the importance of additional mediators. Candidate factors include VEGF,29 IL-6,29 and IL-830 as well as secreted molecules with multiple demonstrated functionalities like
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Fig 6. Placenta-derived adherent cell (PDAC) treatment increases the angioscore in rat ischemic limb. A, Representative angiographs taken at day 35 after surgery. Rats were treated with (A) vehicle or (B) PDACs (4 104/kg). The arrows indicate the surgical site. C, Angioscore of rat hindlimb, 35 days after induction of ischemia. Rats were treated with the indicated doses of PDACs or vehicle control; angioscore of the undamaged limb is defined as 100% (n ¼ 5). The error bars indicate standard deviation.
Fig 7. Histology of cross sections of mouse quadriceps muscle. Quadriceps slices from mice (49 days after surgery) were stained with hematoxylin and eosin; representative micrographs (40 magnification) are shown. The pink shapes are muscle fibers; the white borders are connective tissue. A, Nonsurgical limb. B, Surgical limb from animals treated with vehicle control. C, Vascular endothelial growth factor (VEGF) control. D, Placenta-derived adherent cells (PDACs) at 4 104/kg dose. Scale bar ¼ 50 mm. The black dots in B likely represent infiltration of immune cells in the vicinity of a blood vessel.
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Fig 8. A, Evaluation of capillary density in cross sections of mouse quadriceps muscle. Quadriceps slices from mice (49 days after surgery) were stained with indoxyl-tetrazolium to detect alkaline phosphatase, a marker of newly synthesized capillary endothelial cells. Representative micrographs (40 magnification) are shown for surgical limbs treated with vehicle control (left panel) or placenta-derived adherent cells (PDACs, 4 104/kg dose; right panel). The arrows designate representative areas of alkaline phosphatase staining. Scale bar ¼ 100 mm (n ¼ 5). B, Size distribution of small and large blood vessels in mouse limbs 49 days after surgery. PDACs were administered at the indicated doses (n ¼ 5). The error bars indicate standard deviation.
prostaglandins. Prostaglandin E2 has been shown to positively regulate the angiogenic response of endothelial cells, particularly under hypoxic conditions, and thereby may contribute to the angiogenic and tissue repair processes during vascular or ischemia muscle injuries.31 Interestingly, we have previously shown that PDACs can secrete prostaglandin E2 and thereby modulate the differentiation of dendritic cells,7 which in turn have been suggested to partake in the angiogenic process.31 In combination with the data shown herein, these findings support the hypothesis that PDACs can respond to specific stimuli in a context-sensitive manner by the secretion of various soluble factors that regulate a multitude of biologic processes. Mechanism of action studies are currently under way to evaluate the role of secreted factors like prostaglandins and interleukins in tissue repair processes and to understand the effect of PDACs on the interplay between immunomodulation and angiogenesis. Taken together, these findings highlight the multiple functions that PDACs may offer for the treatment of diseases that do not respond to standard therapies or single biologics. In the mouse and rat HLI models, PDACs demonstrated sustained effects beyond their observed persistence of 8 days. The effects also lasted longer than the endogenous healing process that usually reaches a plateau at 21 to 28 days after surgery. Indeed, blood flow in the injured limb of mice continued to increase at 49 days after PDAC administration, and histologic evaluation of the injury site indicated that the numbers of both small and large blood vessels were increased in comparison to untreated controls. Furthermore, greater increases in large blood vessels were observed with PDAC treatment compared with VEGF
treatment. The secretion of high levels of HGF and modest levels of VEGF by PDACs may contribute to the observed angiogenic effects. However, nonviable PDACs as well as HDF were ineffective in the HLI model, which has been shown previously,32 suggesting that PDACs are not merely a delivery system for growth factors and do not convey their effects through a nonspecific inflammatory response to human cells. The diverse beneficial effects of live PDACs, including generation of new small blood vessels and support for their subsequent maturation, protection of the existing larger blood vessels in the injured tissue, and reduction of inflammatory infiltrates and promotion of muscle regeneration, suggest that PDACs exert an immediate, local angiogenic effect that triggers downstream events leading to a long-term effect on the endogenous homeostasis of the injured tissue. The current study demonstrating the angiogenic capacity of PDACs is consistent with our previous results in a rodent model of stroke,8 in which PDACs (administered intravenously) promoted endothelial cell proliferation and vascular density in ischemic brain, and with results from other laboratories examining MSCs from various sources (mainly bone marrow and adipose tissue) in rodent HLI.19,20 However, human placenta has unique attributes as the source of mesenchymal-like cells for therapeutic use; in addition to the ready availability of healthy, young, and abundant donor tissue from full-term pregnancies, the status of the placenta as “immune privileged”33 may confer a possible advantage for allogeneic, “off-the-shelf” use in patients. In addition, we note that PDACs were reproducibly effective at a dose of 40,000 cells/kg intramuscularly, which translates to 1000 cells per mouse or rat. In contrast
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to our results with PDACs, studies with MSCs from other sources in HLI models reported efficacy at intramuscular doses of 105 to 107 MSCs per mouse.19 The reason for this disparity is not clear and may be due to differences in cell types, expansion methods, or formulation. CONCLUSIONS We have demonstrated that PDACs can promote angiogenesis and tissue regeneration, suggesting that the cells may be a promising candidate for treatment of complex peripheral arterial diseases. PDACs enhance endothelial cell survival in vitro and promote blood vessel formation and maturation resulting in increased blood flow and vascular density, reduce inflammation, and promote muscle regeneration in animal models of ischemic injury. A live cell therapy with diverse functions, and potentially with the ability to dynamically respond to environmental signals and to stimulate repair functions that last beyond their presence in vivo, PDAC treatment may be more effective than single angiogenic growth factors or gene therapy approaches. Studies in progress are aimed at elucidating the mechanisms of action by which this cell therapy exerts its effects. AUTHOR CONTRIBUTIONS Conception and design: AF, KL, SH, SA, UH, WH, RH Analysis and interpretation: AF, KL, SH, SA, EB, UH Data collection: KL, SH Writing the article: AF, EB, SA, KL, SH Critical revision of the article: AF, EB, UH, WH, RH Final approval of the article: AF, KL, SH, SA, EB, UH, WH, RH Statistical analysis: KL, SH Obtained funding: Not applicable Overall responsibility: AF REFERENCES 1. Zheng ZZ, Liu ZX. Activation of the phosphatidylinositol 3-kinase/ protein kinase Akt pathway mediates CD151-induced endothelial cell proliferation and cell migration. Int J Biochem Cell Biol 2007;39: 340-8. 2. Hockel M, Schlenger K, Doctrow S, Kissel T, Vaupel P. Therapeutic angiogenesis. Arch Surg 1993;128:423-9. 3. Annex BH. Therapeutic angiogenesis for critical limb ischaemia. Nat Rev Cardiol 2013;10:387-96. 4. Strioga M, Viswanathan S, Darinskas A, Slaby O, Michalek J. Same or not the same? Comparison of adipose tissue-derived versus bone marrow-derived mesenchymal stem and stromal cells. Stem Cells Dev 2012;21:2724-52. 5. Ma S, Xie N, Li W, Yuan B, Shi Y, Wang Y. Immunobiology of mesenchymal stem cells. Cell Death Differ 2014;21:216-25. 6. Kwon HM, Hur SM, Park KY, Kim CK, Kim YM, Kim HS, et al. Multiple paracrine factors secreted by mesenchymal stem cells contribute to angiogenesis. Vascul Pharmacol 2014;63:19-28. 7. Liu W, Morschauser A, Zhang X, Lu X, Gleason J, He S, et al. Human placenta-derived adherent cells induce tolerogenic immune responses. Clin Trans Immunology 2014;3:e14. 8. Shehadah A, Chen J, Pal A, He S, Zeitlin A, Cui X, et al. Human placenta-derived adherent cell treatment of experimental stroke promotes functional recovery after stroke in young adult and older rats. PLoS One 2014;9:e86621.
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9. He S, Khan J, Gleason J, Eliav E, Fik-Rymarkiewicz E, Herzberg U, et al. Placenta-derived adherent cells attenuate hyperalgesia and neuroinflammatory response associated with perineural inflammation in rats. Brain Behav Immun 2013;27:185-92. 10. Mayer L, Pandak WM, Melmed GY, Hanauer SB, Johnson K, Payne D, et al. Safety and tolerability of human placenta-derived cells (PDA001) in treatment-resistant Crohn’s disease: a phase 1 study. Inflamm Bowel Dis 2013;19:754-60. 11. Ribatti D, Vacca A, Roncali L, Dammacco F. The chick embryo chorioallantoic membrane as a model for in vivo research on angiogenesis. Int J Dev Biol 1996;40:1189-97. 12. Goto T, Fukuyama N, Aki A, Kanabuchi K, Kimura K, Taira H, et al. Search for appropriate experimental methods to create stable hind-limb ischemia in mouse. Tokai J Exp Clin Med 2006;31:128-32. 13. Li X, Ling W, Pennisi A, Wang Y, Khan S, Heidaran M, et al. Human placenta-derived adherent cells prevent bone loss, stimulate bone formation, and suppress growth of multiple myeloma in bone. Stem Cells 2011;29:263-73. 14. Belch J, Hiatt WR, Baumgartner I, Driver IV, Nikol S, Norgren L, et al. Effect of fibroblast growth factor NV1FGF on amputation and death: a randomised placebo-controlled trial of gene therapy in critical limb ischaemia. Lancet 2011;377:1929-37. 15. Khan TA, Sellke FW, Laham RJ. Gene therapy progress and prospects: therapeutic angiogenesis for limb and myocardial ischemia. Gene Ther 2003;10:285-91. 16. Caplan AI, Correa D. The MSC: an injury drugstore. Cell Stem Cell 2011;9:11-5. 17. Murphy MB, Moncivais K, Caplan AI. Mesenchymal stem cells: environmentally responsive therapeutics for regenerative medicine. Exp Mol Med 2013;45:e54. 18. Li Y, Chopp M. Marrow stromal cell transplantation in stroke and traumatic brain injury. Neuroscience Lett 2009;456:120-3. 19. Liew A, O’Brian T. Therapeutic potential for mesenchymal stem cell transplantation in critical limb ischemia. Stem Cell Res Ther 2012;3:28. 20. Bronckaers A, Hilkens P, Martens W, Gervois P, Ratajczak J, Struys T, et al. Mesenchymal stem/stromal cells as a pharmacological and therapeutic approach to accelerate angiogenesis. Pharmacol Ther 2014;143:181-96. 21. DiMarino AM, Caplan AI, Bonfield TL. Mesenchymal stem cells in tissue repair. Front Immunol 2013:e201. 22. Xu KP, Yu FS. Cross talk between c-Met and epidermal growth factor receptor during retinal pigment epithelial wound healing. Invest Ophthalmol Vis Sci 2007;48:2242-8. 23. Shimamura M, Sato N, Waguri S, Uchiyama Y, Hayashi T, Iida H, et al. Gene transfer of hepatocyte growth factor gene improves learning and memory in the chronic stage of cerebral infarction. Hypertension 2006;47:742-51. 24. Yang ZJ, Zhang YR, Chen B, Zhang SL, Jia EZ, Wang LS, et al. Phase I clinical trial on intracoronary administration of Ad-hHGF treating severe coronary artery disease. Mol Biol Rep 2009;36:1323-9. 25. Trapp T, Kogler G, El-Khattouti A, Sorg RV, Besselmann M, Focking M, et al. Hepatocyte growth factor/c-MET axis-mediated tropism of cord blood-derived unrestricted somatic stem cells for neuronal injury. J Biol Chem 2008;283:32244-53. 26. Kaga T, Kawano H, Sakaguchi M, Nakazawa T, Taniyama Y, Morishita R. Hepatocyte growth factor stimulated angiogenesis without inflammation: differential actions between hepatocyte growth factor, vascular endothelial growth factor and basic fibroblast growth factor. Vascul Pharmacol 2012;57:3-9. 27. Morishita R, Makino H, Aoki M, Hashiya N, Yamasaki K, Azuma J, et al. Phase I/IIa clinical trial of therapeutic angiogenesis using hepatocyte growth factor gene transfer to treat critical limb ischemia. Arterioscler Thromb Vasc Biol 2011;31:713-20. 28. Powell RJ, Simons M, Mendelsohn FO, Daniel G, Henry TD, Koga M, et al. Results of a double-blind, placebo-controlled study to assess the safety of intramuscular injection of hepatocyte growth factor plasmid to improve limb perfusion in patients with critical limb ischemia. Circulation 2008;118:58-65. 29. Ebrahem Q, Minamoto A, Hoppe G, Anand-Apte B, Sears JE. Triamcinolone acetonide inhibits IL-6- and VEGF-induced
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angiogenesis downstream of the IL-6 and VEGF receptors. Invest Ophthalmol Vis Sci 2006;47:4935-41. 30. Heidemann J, Ogawa H, Dwinell MB, Rafiee P, Maaser C, Gockel HR, et al. Angiogenic effects of interleukin 8 (CXCL8) in human intestinal microvascular endothelial cells are mediated by CXCR2. J Biol Chem 2003;278:8508-15. 31. Motz GT, Coukos G. The parallel lives of angiogenesis and immunosuppression: cancer and other tales. Nat Rev Immunol 2011;11: 702-11. 32. Madonna R, Delli Pizzi S, Di Donato L, Mariotti A, Di Carlo L, D’Ugo E, et al. Non-invasive in vivo detection of peripheral limb
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ischemia improvement in the rat after adipose tissue-derived stromal cell transplantation. Circ J 2012;76:1517-25. 33. Guller S, LaChapelle L. The role of placental Fas ligand in maintaining immune privilege at maternal-fetal interfaces. Semin Reprod Endocrinol 1999;17:39-44.
Submitted Nov 12, 2014; accepted Apr 4, 2015.
Additional material for this article may be found online at www.jvascsurg.org.
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APPENDIX (online only) As part of the Celgene Cellular Therapeutics (CCT) Research Group, the following group members significantly contributed to the work described in this manuscript: Aleksandr Kaplunovsky, Jennifer Paredes, Allan Reduta, Eric Law, Ewa Fik, Sascha Abramson, Vivian R. Albert, and Itschak Lamensdorfa (aPharmaseed Ltd). SUPPLEMENTARY METHODS (online only) Preparation of placenta-derived adherent cells (PDACs), human umbilical vein endothelial cells (HUVECs), human dermal fibroblast (HDF), and human mesenchymal stromal cells (MSCs). PDACs were prepared by mechanical and enzymatic digestion of human placental tissue of newborn origin obtained from a normal, full-term birth as described.7 PDACs were expanded until passage 6 in PDAC medium,7 which, unless otherwise indicated, contains 2% fetal bovine serum (Invitrogen, Carlsbad, Calif). Cryopreserved PDACs (passage 6 at 7.5 106 or 20 106 cells/mL in 5% dimethyl sulfoxide, 10% human serum albumin, and 5.5% dextran in 0.9% NaCl) were thawed in a 37 C water bath. Viability and cell counts were assessed by trypan blue exclusion; cells were used within 4 hours of thaw. Flow cytometric analysis of PDACs. Flow cytometric surface marker staining of PDACs was performed with labeled monoclonal mouse antihuman immunoglobulin G (BD Biosciences, San Jose, Calif). For surface marker expression analysis, PDAC single-cell suspensions were washed with fluorescence-activated cell sorting (FACS) wash buffer (phosphate-buffered saline with 2% fetal bovine serum) and labeled with antibodies by incubation at 4 C for 30 minutes. Flow cytometry analysis was performed on a FACSCanto II (BD Biosciences). Data were acquired with FACSDiva software (BD Biosciences) and analyzed with FlowJo flow cytometry software (Tree Star, Ashland, Ore). HUVEC survival assay. HUVECs were subjected to serum starvation in the presence and absence of PDACs, HDF, or MSCs, using indirect coculture systems. HUVECs were seeded (2 104 cells/well) in a 24-well tissue culture-treated polystyrene precoated with fibronectin (Sigma-Aldrich, St. Louis, Mo). After overnight incubation in complete endothelial cell growth medium 2 (Lonza, Walkersville, Md) to facilitate cell attachment and proliferation, HUVECs were serum starved in endothelial cell basal medium (Lonza) for 6 hours. Concomitant with HUVEC seeding, 0 to 5 104 PDACs, HDF, or MSCs were independently seeded into 8-mm-pore Transwell culture inserts (Corning, Lowell,
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Mass) in a separate multiwell plate in PDAC medium and incubated 2 hours to allow attachment. Subsequently, PDAC medium was replaced with serum-free Dulbecco’s modified Eagle medium (DMEM, Invitrogen) for 24 hours to prepare cell-conditioned medium (CM) containing factors secreted by PDACs, HDF, or MSCs. The following day, HUVEC culture medium was replaced with PDACCM. In addition, PDACs, HDF, or MSCs Transwell inserts were placed into the HUVEC multiwell plates for indirect coculture between PDACs, HDF, or MSCs and HUVECs overlaid with PDAC-CM. After 24 hours, the Transwell cocultures and media were removed, and HUVEC counts were determined with the CellTiter-Glo assay (Promega, Madison, Wisc). Tube formation assay. HUVECs were cultured to passage 3 in endothelial cell growth medium 2 for 3 days to w75% confluency. HUVECs were washed and resuspended in DMEM and diluted to 4000 cells/mL with human placental collagen (1.5 mg/mL in DMEM). After pipetting (3 mL/well) into 96-well plates (n ¼ 5 per condition), HUVEC/human placental collagen drops were incubated for 90 minutes (37 C, 5% CO2, and 90% relative humidity) to facilitate collagen solidification, followed by addition of 200 mL of PDAC-CM or control medium. After incubation (24 hours), HUVEC drops were stained with a Diff-Quik Cell Staining Kit (Dade Behring, Inc, Newark, Del). Images of each well were acquired with the Zeiss SteREO Discovery V8 microscope (Carl Zeiss AG, Jena, Germany) and analyzed for average network area. Chicken embryo chorioallantoic membrane assay. Chorioallantoic membrane assays in fertilized chicken eggs were used to assess neovascularization and were performed by SRI International (Menlo Park, Calif) as described.11 The test articles (n ¼ 8-10) applied on day 6 were PDACs (7.7 105 viable cells) in 40 mL of DMEM/Matrigel vehicle mixture (1:1), positive control (100 ng/mL basic fibroblast growth factor in DMEM/ Matrigel mixture), Matrigel vehicle control, and DMEM control. On day 8, blood vessel density was determined by an image capturing system at a magnification of 100. Unilateral hindlimb ischemia induction and administration of PDACs. The surgical procedure followed the Prox-A protocol.12 Under anesthesia, an incision (mice, 0.5-1.0 cm; rats, 1.0-2.0 cm) was made in the skin in the inguinal area. The femoral artery was ligated twice with 6-0 (mice) or 4-0 (rats) silk thread and transected between the ligatures. The wound was closed with 4-0 (mice) or 3-0 (rats) silk thread, and the animal was allowed to recover.
Angiogenesis is a complex process that facilitates blood vessel formation and tissue perfusion. It is associated with a balance of stimulatory and inhibitory steps, including secretion of various soluble factors, degradation of extracellular matrix, and proliferation and differentiation of endothelial cells into tubelike vessels.1 The term therapeutic angiogenesis refers to situations in which angiogenic stimulation is a From the Research & Development Department, Celgene Cellular Therapeutics. The Appendix (online only) lists the names of the Celgene Cellular Therapeutics Research Group members. Author conflict of interest: All authors except I.L. are employees of Celgene and hold stock or stock options in Celgene. I.L. indicates no potential conflicts of interest and received funding from Celgene Cellular Therapeutics through a contract service agreement for conducting the in vivo studies. Additional material for this article may be found online at www.jvascsurg.org. Reprint requests: Aleksandar Francki, PhD, Research & Development Department, Celgene Cellular Therapeutics, 7 Powderhorn Dr, Warren, NJ 07059 (e-mail: [email protected]). The editors and reviewers of this article have no relevant financial relationships to disclose per the JVS policy that requires reviewers to decline review of any manuscript for which they may have a conflict of interest. 0741-5214 Copyright Ó 2015 by the Society for Vascular Surgery. Published by Elsevier Inc. http://dx.doi.org/10.1016/j.jvs.2015.04.387
clinically desirable outcome (eg, tissue repair or treatment of local hypovascularity).2 To date, no small-molecule or biologic-based approach for therapeutic angiogenesis has proved consistently efficacious, and several compounds that showed promise in preclinical and early clinical trials failed to demonstrate efficacy in larger randomized trials.3 These failures may reflect the fact that angiogenesis and tissue repair are not regulated by a single molecule or biologic pathway but by an intricate, temporally coordinated network of biologic processes. It has been suggested that cell therapy using mesenchymal stromal cells (MSCs) or mesenchymal-like cells available from a variety of sources, such as bone marrow and placenta, can promote angiogenesis and tissue regeneration through multiple biologic activities, offering a potentially transformational means to treat diseases with significant unmet clinical needs.4 These cell therapies have been shown to modulate pathologic immune responses and inflammatory processes5 as well as the secretion of specific trophic factors that stimulate resident endothelial cells or progenitors to initiate angiogenic processes in injured tissue.6 Placenta-derived adherent cells (PDACs) are a cultureexpanded, undifferentiated mesenchymal-like population derived from full-term placental tissue.7 The cells have 1
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the genotype of the newborn and display features characteristic of MSCs, but with a unique phenotype associated with their placental origin. The cells are anchorage dependent in culture and adhere to plastic during expansion in vitro.7 PDAC-mediated activities have been studied in vitro, in animal models, and in clinical trials and display immunomodulatory, anti-inflammatory, proregenerative, and neuroprotective properties,7-10 suggesting that the cells may have potential utility across a variety of disease indications. PDA-002, an intramuscular formulation of PDACs, is in clinical development for the treatment of peripheral arterial disease with diabetic foot ulcer (http:// clinicaltrials.gov/show/NCT01859117). In this study, we investigated whether in addition to previously described immunomodulatory capabilities, PDACs display angiogenic properties in various assays and animal models relevant to angiogenesis. We show that PDACs secrete a variety of proangiogenic factors (most prominently hepatocyte growth factor [HGF]), support endothelial cell survival, induce blood vessel formation, sustain blood vessel growth and maintenance, and induce muscle repair in rodent hindlimb ischemia (HLI) models, demonstrating significant proangiogenic and reparative functions. These results support the continued investigation of PDACs as cell therapy for vascular disease-related indications. METHODS Preparation of PDACs, human umbilical vein endothelial cells (HUVECs), human dermal fibroblast (HDF), and human MSCs. PDACs were prepared by mechanical and enzymatic digestion of human placental tissue of newborn origin obtained from a normal, full-term birth as described.7 PDACs were expanded until passage 6 in PDAC medium7 and subsequently used as described in the Supplementary Methods (online only). HUVECs, HDF, and human bone marrow-derived MSCs were obtained from Lonza (Walkersville, Md) and culture expanded according to the manufacturer’s instructions. Flow cytometric analysis of PDACs. Flow cytometric surface marker staining of PDACs was performed using labeled monoclonal mouse antihuman immunoglobulin G (BD Biosciences, San Jose, Calif), and expression was analyzed as described in the Supplementary Methods (online only). Preparation of cell-conditioned medium (CM). PDACs, HDF, or MSCs were seeded onto tissue culturetreated polystyrene 24-well plates (Corning, Lowell, Mass) at 4000 cells/cm2 for overnight establishment in PDAC medium before incubation in serum-free PDAC medium for 48 hours (37 C, 5% CO2, 21% O2, 90% relative humidity). CM was then collected and frozen at 80 C. HUVEC survival assay. HUVECs were subjected to serum starvation in the presence and absence of PDACs, HDF, or MSCs, using indirect coculture systems as detailed in the Supplementary Methods (online only).
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After 24 hours, the Transwell (Corning) cocultures and media were removed, and HUVEC counts were determined by the CellTiter-Glo assay (Promega, Madison, Wisc). Tube formation assay. HUVECs were cultured to passage 3 in endothelial cell growth medium 2 for 3 days to w75% confluency. HUVECs were washed and resuspended in Dulbecco’s modified Eagle medium (DMEM) and diluted to 4000 cells/mL with human placental collagen (1.5 mg/mL in DMEM), and the tube formation assay was performed as described in the Supplementary Methods (online only). After incubation (24 hours), HUVEC drops were stained with a Diff-Quik Cell Staining Kit (Dade Behring, Inc, Newark, Del). Images of each well were acquired with the Zeiss SteREO Discovery V8 microscope (Carl Zeiss AG, Jena, Germany) and analyzed for average network area. Quantitation of secreted factors. The concentrations of angiogenic factors basic fibroblast growth factor 2 (FGF-2), granulocyte colony-stimulating factor (G-CSF), interleukin-8 (IL-8), platelet-derived growth factor BB (PDGF-BB), vascular endothelial growth factor (VEGF), interleukin-6 (IL-6), and monocyte chemotactic protein 1 (MCP-1) were analyzed in PDAC-CM using Bio-Plex Pro multiplex bead array systems (Bio-Rad Laboratories, Hercules, Calif). HGF concentration was determined using a solid-phase sandwich enzyme-linked immunosorbent assay system (Quantikine Human HGF Immunoassay; R&D Systems, Minneapolis, Minn). Chicken embryo chorioallantoic membrane (CAM) assay. CAM assays in fertilized chicken eggs were used to assess neovascularization and were performed by SRI International (Menlo Park, Calif) as described11 and detailed in the Supplementary Methods (online only). Animal care. All animal work was performed at Pharmaseed Ltd (Ness Ziona, Israel) using adult Balb/c mice (w25 g) or Sprague-Dawley rats (w200 g) according to the National Institutes of Health and the Association for Assessment and Accreditation of Laboratory Animal Care. Unilateral HLI induction and administration of PDACs. The surgical procedure followed the Prox-A protocol12 as detailed in the Supplementary Methods (online only). At 24 hours after surgery, each animal was treated with PDACs, recombinant human VEGF 165 (3.3 mg/animal; Sigma-Aldrich, St. Louis, Mo), or vehicle (cell freezing media) by intramuscular injection proximal and distal to the surgical wound. Mice were injected with 25 mL in each of two sites (total 50 mL); rats were injected with 50 mL in each of four sites (total 200 mL). PDAC doses were w4 104, 4 105, or 4 106 cells/kg. Luciferase assay for PDAC retention. PDACs constitutively expressing luciferase were prepared with the Cignal lenti positive control (luc) system (SABiosciences, Valencia, Calif), and 1 106 cells (in 25 mL) were injected intramuscularly into the ischemic hindlimb of Balb/c mice. Luciferin substrate (150 mg/kg; Caliper Life Sciences, Hopkinton, Mas) was administered intravenously, and
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luciferin intensity was measured in the whole animal with an in vivo real-time imaging system (IVIS Spectrum 200; Caliper Life Sciences). Laser Doppler analysis. Blood flow in surgical and control (nonsurgical) hindlimbs was measured with a noncontact laser Doppler before surgery and on days 1, 7, 15, 21, 28, and 35 for both mice and rats. In addition, mice were measured at 42 and 49 days. Blood flow in the nonischemic control limb is defined as 100%; measurements are expressed as percentage flow in ischemic limb compared with control limb. Angiography of rat hindlimbs. On day 35 after surgery, half of the animals in each group were anesthetized with ketamine/xylazine. Barium sulfate solution (2.5 mL) was infused into the infrarenal aorta after ligation of the proximal aorta and inferior vena cava, and the number of contrast-filled blood vessels was determined by X-ray roentgen assays. Quantitation of intersections between contrast-filled vessels and gridlines of the angiographs is presented as an angioscore, defined as the number of blocks on a grid filled by radiopaque dye, and was performed in blinded fashion. Histology of hindlimb muscle. After sacrifice, quadriceps muscles of ischemic and control limbs were removed, weighed, fixed in Hepes-glutamic acid buffer mediated Organic solvent Protection Effect (HOPE) fixative (DCS Innovative, Hamburg, Germany), and embedded in paraffin. Paraffin blocks were cooled to 20 C for 30 minutes and sectioned with a microtome at 5 mm thickness. Muscle anatomy was visualized by staining two serial sections with hematoxylin and eosin and evaluated microscopically for morphologic changes. Quantitation of blood vessel density in muscle sections. The indoxyl-tetrazolium method for alkaline phosphatase was used to detect capillary endothelial cells in newly synthesized vessels. Staining was performed using the BCIP/NBT kit from Vector Labs (Burlingame, Calif; catalog No. SK-5400), and nuclei were counterstained with methyl green (Vector Labs; catalog No. H-3402). Muscle capillary density was evaluated in blinded fashion in sections from three representative animals by counting positively stained capillaries within three randomly chosen fields per section at 20 magnification. RESULTS Flow cytometric characterization of PDACs. As previously described,7 PDACs expressed surface markers that are typical of MSCs, including CD90, CD73, and CD105, as well as CD200, which is associated with placental cells but absent on other MSC-like cells, thereby distinguishing PDACs from MSCs. Further immunophenotypic characterization showed that PDACs do not express CD31, KDR, CD133, VE cadherin (CD144), and Tie2. In conjunction with the described lack of CD34 expression, these findings confirmed the absence of endothelial progenitor cells and mature endothelial cells in PDAC preparations.
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PDACs enhance the survival of endothelial cells. The effect of PDACs on the survival of serum-starved HUVECs was examined in a noncontact coculture system. After serum starvation and 24 hours of indirect coculture of the two cell types, only 35% of the cell count of HUVECs remained in the absence of PDACs (Fig 1, A, serum-free DMEM). In contrast, indirect coculture with PDACs resulted in 72% HUVEC survival. Although we refer to the observed increase in the HUVEC count in the presence of PDACs compared with control as “enhanced survival (reduced cell death),” enhanced HUVEC proliferation may also be a component. Other cell types secrete factors that may render them active in HUVEC survival, tube formation, and CAM assays; these in vitro and ex vivo angiogenic activities are not unique to PDACs, HDF, or MSCs. To address potential differences between the aforementioned cell types and PDACs, we tested HDF and MSCs in the HUVEC survival assay under equal experimental conditions. All three tested cell types showed an effect that was significantly higher in comparison to the control medium; however, the effect of PDACs and the CM was consistently 20% higher compared with the effects of HDF and MSCs and their respective CM (data not shown). These results support that PDACs have unique in vitro effects on the angiogenic functions of human endothelial cells compared with other relevant cell types. Dose-response studies were performed to determine the ratio of PDACs to HUVECs required to promote HUVEC survival (Fig 1, B). A range of PDACs (0-40,000) were used, whereas the input HUVECs were kept constant at w50,000 cells (PDAC:HUVEC of 0.1:1 to 0.8:1). When cultured in DMEM alone, the HUVEC count decreased by w60%, to 20,000 cells. As few as 5000 PDACs had a statistically significant effect on the HUVEC count (P < .01), with a plateau observed at approximately 20,000 PDACs, resulting in twofold higher final HUVEC count compared with the DMEM control (P < .01). Based on the starting HUVEC count, PDACs exerted a beneficial effect on HUVEC survival at as little as 0.1:1, with a plateau observed at w0.4:1 PDAC:HUVEC. The lack of direct contact between PDACs and HUVECs in these experiments suggests that the effect is mediated through factors elaborated by PDACs. PDAC-CM stimulates endothelial cell tube formation. The ability of CM from PDACs to promote HUVEC tube formation was examined by the application of PDAC-CM to HUVECs cultured within a threedimensional environment of collagen. After 24 hours of incubation, tube formation (as measured by average network area) was w70% greater for CM than for serumfree media (P < .01; Fig 2). Trophic factor protein secretion of PDACs. Because direct PDAC contact with HUVECs was not required to stimulate HUVEC survival or tube formation (Figs 1 and 2), it was of interest to examine protein secretion by PDACs. Assessment of eight proteins (Fig 3) revealed that PDACs secrete several mitogens and cytokines
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Fig 1. A, Effect of placenta-derived adherent cells (PDACs) on human umbilical vein endothelial cell (HUVEC) survival. HUVECs were exposed to PDACs through a Transwell (PDAC:HUVEC, w2:1). The initial value of HUVECs (after 6-hour serum starvation in endothelial cell basal medium and before Transwell exposure) was set to 100%. The difference between HUVEC survival in serum-free Dulbecco’s modified Eagle medium (SF-DMEM) overnight (w35%) and in the presence of PDACs (72%) was statistically significant (P < .01; n ¼ 7). B, Dosedependent effect of PDACs on HUVEC survival (n ¼ 3). The error bars indicate standard deviation.
relevant to angiogenesis. FGF-2, G-CSF, IL-8, and PDGF-BB were secreted in low but detectable levels (<100 pg/mL); VEGF and IL-6 were secreted at moderate levels (140 and 480 pg/mL, respectively). MCP-1 and HGF were secreted at substantially higher levels (1200 and 1600 pg/mL, respectively). Ex vivo proangiogenic effects of PDACs. To investigate the effect of PDACs on vascularization in an ex vivo model, PDACs were cultured on exposed CAM of chick embryos, and the change in blood vessel density was assessed. As shown in Fig 4, PDACs caused a substantial increase (w5- to 10-fold) in blood vessel density compared with medium and vehicle controls and was indistinguishable from positive control FGF-2, a known proangiogenic factor. PDAC administration increases blood flow in rodent models of HLI. The proangiogenic effects of PDACs led us to test their efficacy in rodent models of HLI. After induction of HLI, blood flow was measured by laser Doppler with and without intramuscular administration of PDACs. Throughout the mouse study (Fig 5, A), an increase in blood flow was observed in all animal groups treated with PDACs compared with vehicle-treated
control. The improvement in blood flow was visible from day 28 and continued to increase through day 49 after surgery, similar to the positive control, VEGF. The increase in blood flow in all PDAC treatment groups compared with vehicle control was statistically significant (from day 28 through day 49; P < .01). In a rat study, an increase in blood flow was observed in all animal groups treated with PDACs (4 104, 4 105, and 4 106 cells/kg) compared with the vehicle-treated control (Fig 5, B). This improvement was observed on day 35 after HLI initiation in all PDAC-treated groups and was statistically significant compared with vehicle control (P < .001 for day 35 at all PDAC doses). To determine whether PDAC viability is required for the observed increase in blood flow, nonviable cells were prepared by repeated freeze-thaw cycles before administration in the rat HLI model; nonviable PDACs failed to improve blood flow. Because rodents subjected to HLI continued to experience improvement in blood flow for an extended time (49 days after intramuscular administration of PDACs into leg muscle), it was of interest to examine the in vivo location and retention time of PDACs using luciferin
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Fig 2. Stimulation of human umbilical vein endothelial cell (HUVEC) network tube formation by placenta-derived adherent cell (PDAC)-conditioned medium (CM). Representative photomicrographs of PDAC-induced (upper panel) and serum-free Dulbecco’s modified Eagle medium (DMEM) control (lower panel) network (tube) formation are shown. The arrows depict representative tubes.
substrate and PDACs expressing luciferase in a mouse HLI model. As shown in Fig 5, C, the PDAC signal was detectable at 4, 24, and 72 hours after injection only at the leg injection sites, but not in any major organs (eg, brain, heart, lung, liver, kidneys; data not shown). However, no luciferin signal was detected at the injection site or elsewhere in the animals after 8 days, suggesting only short-term persistence of PDACs in the mouse leg and no migration or integration of the cells in the treated animals. Thus, recovery of blood flow in PDAC-treated mice continued for several weeks after the administered cells were no longer detectable. Thus, in both mouse and rat models of HLI, intramuscular injection of PDACs was effective in restoring blood flow to the ischemic limb beyond the measured persistence of PDACs. Angiography assessment of rat hindlimbs. To further investigate the restoration of blood flow observed in the rat HLI model, blood vessel density was examined by angiography of ischemic limb from rats with and without PDAC treatment, 35 days after induction of ischemia (Fig 6). By visual inspection, higher blood vessel density was observed on PDAC treatment (Fig 6, B) compared with the vehicle control (Fig 6, A) and is
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quantified in Fig 6, C. PDAC treatment resulted in a twofold increase in angioscore compared with vehicle control (P < .01 or .05 for day 35 at both PDAC doses). Histology of ischemic limb muscle. To determine the effect of PDACs on the injured muscle tissue in the HLI models, hematoxylin and eosin staining was performed on representative tissue sections, allowing optimal visualization of potential repair of the injured muscle. Muscle fibers in mouse quadriceps muscle from ischemic and healthy hindlimbs were examined (Fig 7). In nonischemic limb, muscle fibers had a polygonal appearance with relatively narrow borders of connective tissue (Fig 7, A). In ischemic limb (Fig 7, B), the myofibers appeared smaller and irregular, with correspondingly larger borders of surrounding connective tissue, and the muscle tissue frequently showed inflammatory infiltrates adjacent to blood vessels. Treatment with PDACs (Fig 7, D) led to the reduction of inflammatory infiltrates and marked improvement in the structure of muscle fibers, with an appearance similar to uninjured muscle in the nonischemic limb (Fig 7, A), but with myofibers containing multiple, centrally located nuclei, indicative of tissue regeneration. PDACs had an effect similar to the VEGF control (compare Fig 7, C and D); however, PDAC treatment resulted in an apparent further reduction of interspersing connective tissue. The effects of PDAC treatment on rat muscle histology were similar (data not shown). Murine blood vessel density measurement. Vascular structures in PDAC-treated mice were visualized (Fig 8, A) and quantitated (Fig 8, B), using VEGF as the positive control. PDAC treatment (4 104/kg or 4 105/kg) resulted in markedly higher numbers of all sizes of blood vessels (capillaries, 50-100 and 300-700 mm2; arterioles, 300-700 mm2; and larger blood vessels, >700 mm2) in comparison to vehicle control. VEGF was similar to PDAC treatment in vessel density in the size ranges #700 mm2, but for larger vessels (>700 mm2), PDAC treatment resulted in threefold higher density compared with VEGF. For smaller vessels (#300 mm2), VEGF and PDAC treatment each exceeded the vessel density of the healthy control limb. Thus, treatment with PDACs demonstrated not only muscle regeneration at the level of muscle fibers but also an increase in the number of blood vessels of all sizes examined, relative to vehicle-treated mice. Finally, as stated before, other cell types secrete factors that may render them active in the HLI model; these in vivo activities are not unique to PDACs, MSCs, or even fibroblasts. Therefore, we deemed the use of PDACs alone as well as the use of dead PDACs in the HLI in vivo studies appropriate. However, to address potential differences between the aforementioned cell types and to exclude nonspecific responses related to cross-species transfer of cells into an injury model, we tested HDF at equal cell numbers in the in vivo HLI models. In both mouse and rat in vivo models, HDF failed to restore blood flow or to induce repair of muscle tissue in the ischemic limb (data not shown). Thus, the angiogenic and tissue reparative effects of PDACs observed in rodent HLI are not
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Fig 3. Quantitation of factors secreted by placenta-derived adherent cells (PDACs). Culture supernatants were analyzed as described for the indicated set of secreted factors. Of the factors examined, hepatocyte growth factor (HGF) is the most abundant, followed by monocyte chemotactic protein 1 (MCP-1) and interleukin-6 (IL-6; n ¼ 5). The error bars indicate standard deviation. FGF-2, Fibroblast growth factor 2; G-CSF, granulocyte colony-stimulating factor; IL-8, interleukin-8; PDGF-BB, platelet-derived growth factor BB; VEGF, vascular endothelial growth factor.
Fig 4. Placenta-derived adherent cells (PDACs) promote vascularization. Percentage change of blood vessel (BV) density was measured in the chorioallantoic membrane (CAM) assay. PDACinduced vascularization of the CAM was statistically indistinguishable from that of the positive control fibroblast growth factor 2 (FGF-2) and significantly greater than the BV density induced by either the vehicle control (P < .01) or the basal medium (P < .01; n ¼ 7). The error bars indicate standard deviation.
driven by a nonspecific xenogeneic inflammatory response to human cells. DISCUSSION Human PDACs are a novel MSC-like population previously shown to possess potent immunomodulatory, anti-inflammatory, neuroprotective, and proregenerative properties.7-9,13 In the current study, we explored the angiogenic properties of PDACs. In vitro, PDACs exhibited various activities consistent with angiogenesis, including enhancement of endothelial cell survival under conditions of serum starvation and stimulation of tube formation. PDACs also promoted vascularization in the CAM
assay. Importantly, in clinically relevant disease models of HLI in both rats and mice, intramuscular injection of PDACs resulted in increased blood flow, increased blood vessel density, and repair of damaged tissue. These studies support continued investigation of PDACs as an angiogenic therapeutic. PDACs (formulated as PDA-002 for local injection) are currently in clinical trials for peripheral arterial disease with diabetic foot ulcer. It is becoming evident that treatment of certain vascular diseases with a single factor (eg, FGF, VEGF, or HGF) may have limited efficacy14,15 and may not be sufficient to ameliorate complex pathologic processes with parallel excessive inflammation or tissue degeneration. In contrast to single agents, MSCs and mesenchymal-like cells have been referred to as an “injury drugstore,”16 possessing immunomodulatory, anti-inflammatory, proregenerative, and angiogenic activities and acting in a paracrine fashion to provide trophic support, often without clear evidence of engraftment.17 Mesenchymal-like cells from a variety of sources can promote angiogenesis and tissue regeneration through a wide variety of biologic activities, offering a potentially transformational means of treating diseases that have significant unmet clinical needs.18-21 Caplan and Correa16 posit that MSCs are able to “establish a regenerative microenvironment by secreting bioactive molecules and regulating the local immune response.” Our experiments demonstrate PDAC multifunctionality consistent with that viewpoint. In particular, our experiments using CM from PDACs or noncontact coculture of HUVECs and PDACs strongly suggest that soluble factors secreted by PDACs mediate effects on endothelial cells including cell survival (or proliferation) and tube formation, consistent with results from mesenchymal-like cells from other sources.16,17,20 Angiogenic factors detected in
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Fig 5. A and B, Placenta-derived adherent cell (PDAC) administration increases blood flow in rodent models of hindlimb ischemia (HLI). Blood flow was measured by noncontact laser Doppler at the indicated times after surgery. PDACs were administered at the indicated doses. Blood flow to the undamaged limb is defined as 100%. Representative experiments from duplicate studies are shown. A, Mouse model. The increase in blood flow in all PDAC treatment groups compared with the vehicle control was statistically significant (from day 28 through day 49; P < .01; n ¼ 15). The error bars indicate standard deviation. B, Rat model. At 35 days, the difference in blood flow between PDACtreated groups and the vehicle control group is statistically significant (P < .001). Nonviable PDAC control was statistically indistinguishable from the vehicle control (n ¼ 12). The error bars indicate standard deviation. C, Representative in vivo real-time images at different time points of four mice injected in the ischemic hindlimb with PDACs expressing the luciferase reporter gene; mouse No. 5 received vehicle. PDACs persisted <8 days in mice. ROI, Region of interest; VEGF, vascular endothelial growth factor.
media supernatants from cultured PDACs included FGF-2, G-CSF, IL-8, PDGF-BB, VEGF, IL-6, MCP-1, and most prominently HGF. HGF plays a major role in embryonic development, adult organ regeneration, and wound healing.22 Furthermore, HGF is central to angiogenesis and tissue regeneration23 and is thought to exert positive effects during the repair of vascular pathologic processes, including stroke.23,24 The role of HGF in angiogenesis includes not
only induction of new blood vessel formation but also maturation of nascent blood vessels, suggesting effects distinct from VEGF25 and confirming the requirement for multiple soluble factors to drive the angiogenic process. The lack of significant success of HGF monotherapy in clinical trials for critical limb ischemia26-28 also points to the importance of additional mediators. Candidate factors include VEGF,29 IL-6,29 and IL-830 as well as secreted molecules with multiple demonstrated functionalities like
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Fig 6. Placenta-derived adherent cell (PDAC) treatment increases the angioscore in rat ischemic limb. A, Representative angiographs taken at day 35 after surgery. Rats were treated with (A) vehicle or (B) PDACs (4 104/kg). The arrows indicate the surgical site. C, Angioscore of rat hindlimb, 35 days after induction of ischemia. Rats were treated with the indicated doses of PDACs or vehicle control; angioscore of the undamaged limb is defined as 100% (n ¼ 5). The error bars indicate standard deviation.
Fig 7. Histology of cross sections of mouse quadriceps muscle. Quadriceps slices from mice (49 days after surgery) were stained with hematoxylin and eosin; representative micrographs (40 magnification) are shown. The pink shapes are muscle fibers; the white borders are connective tissue. A, Nonsurgical limb. B, Surgical limb from animals treated with vehicle control. C, Vascular endothelial growth factor (VEGF) control. D, Placenta-derived adherent cells (PDACs) at 4 104/kg dose. Scale bar ¼ 50 mm. The black dots in B likely represent infiltration of immune cells in the vicinity of a blood vessel.
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Fig 8. A, Evaluation of capillary density in cross sections of mouse quadriceps muscle. Quadriceps slices from mice (49 days after surgery) were stained with indoxyl-tetrazolium to detect alkaline phosphatase, a marker of newly synthesized capillary endothelial cells. Representative micrographs (40 magnification) are shown for surgical limbs treated with vehicle control (left panel) or placenta-derived adherent cells (PDACs, 4 104/kg dose; right panel). The arrows designate representative areas of alkaline phosphatase staining. Scale bar ¼ 100 mm (n ¼ 5). B, Size distribution of small and large blood vessels in mouse limbs 49 days after surgery. PDACs were administered at the indicated doses (n ¼ 5). The error bars indicate standard deviation.
prostaglandins. Prostaglandin E2 has been shown to positively regulate the angiogenic response of endothelial cells, particularly under hypoxic conditions, and thereby may contribute to the angiogenic and tissue repair processes during vascular or ischemia muscle injuries.31 Interestingly, we have previously shown that PDACs can secrete prostaglandin E2 and thereby modulate the differentiation of dendritic cells,7 which in turn have been suggested to partake in the angiogenic process.31 In combination with the data shown herein, these findings support the hypothesis that PDACs can respond to specific stimuli in a context-sensitive manner by the secretion of various soluble factors that regulate a multitude of biologic processes. Mechanism of action studies are currently under way to evaluate the role of secreted factors like prostaglandins and interleukins in tissue repair processes and to understand the effect of PDACs on the interplay between immunomodulation and angiogenesis. Taken together, these findings highlight the multiple functions that PDACs may offer for the treatment of diseases that do not respond to standard therapies or single biologics. In the mouse and rat HLI models, PDACs demonstrated sustained effects beyond their observed persistence of 8 days. The effects also lasted longer than the endogenous healing process that usually reaches a plateau at 21 to 28 days after surgery. Indeed, blood flow in the injured limb of mice continued to increase at 49 days after PDAC administration, and histologic evaluation of the injury site indicated that the numbers of both small and large blood vessels were increased in comparison to untreated controls. Furthermore, greater increases in large blood vessels were observed with PDAC treatment compared with VEGF
treatment. The secretion of high levels of HGF and modest levels of VEGF by PDACs may contribute to the observed angiogenic effects. However, nonviable PDACs as well as HDF were ineffective in the HLI model, which has been shown previously,32 suggesting that PDACs are not merely a delivery system for growth factors and do not convey their effects through a nonspecific inflammatory response to human cells. The diverse beneficial effects of live PDACs, including generation of new small blood vessels and support for their subsequent maturation, protection of the existing larger blood vessels in the injured tissue, and reduction of inflammatory infiltrates and promotion of muscle regeneration, suggest that PDACs exert an immediate, local angiogenic effect that triggers downstream events leading to a long-term effect on the endogenous homeostasis of the injured tissue. The current study demonstrating the angiogenic capacity of PDACs is consistent with our previous results in a rodent model of stroke,8 in which PDACs (administered intravenously) promoted endothelial cell proliferation and vascular density in ischemic brain, and with results from other laboratories examining MSCs from various sources (mainly bone marrow and adipose tissue) in rodent HLI.19,20 However, human placenta has unique attributes as the source of mesenchymal-like cells for therapeutic use; in addition to the ready availability of healthy, young, and abundant donor tissue from full-term pregnancies, the status of the placenta as “immune privileged”33 may confer a possible advantage for allogeneic, “off-the-shelf” use in patients. In addition, we note that PDACs were reproducibly effective at a dose of 40,000 cells/kg intramuscularly, which translates to 1000 cells per mouse or rat. In contrast
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to our results with PDACs, studies with MSCs from other sources in HLI models reported efficacy at intramuscular doses of 105 to 107 MSCs per mouse.19 The reason for this disparity is not clear and may be due to differences in cell types, expansion methods, or formulation. CONCLUSIONS We have demonstrated that PDACs can promote angiogenesis and tissue regeneration, suggesting that the cells may be a promising candidate for treatment of complex peripheral arterial diseases. PDACs enhance endothelial cell survival in vitro and promote blood vessel formation and maturation resulting in increased blood flow and vascular density, reduce inflammation, and promote muscle regeneration in animal models of ischemic injury. A live cell therapy with diverse functions, and potentially with the ability to dynamically respond to environmental signals and to stimulate repair functions that last beyond their presence in vivo, PDAC treatment may be more effective than single angiogenic growth factors or gene therapy approaches. Studies in progress are aimed at elucidating the mechanisms of action by which this cell therapy exerts its effects. AUTHOR CONTRIBUTIONS Conception and design: AF, KL, SH, SA, UH, WH, RH Analysis and interpretation: AF, KL, SH, SA, EB, UH Data collection: KL, SH Writing the article: AF, EB, SA, KL, SH Critical revision of the article: AF, EB, UH, WH, RH Final approval of the article: AF, KL, SH, SA, EB, UH, WH, RH Statistical analysis: KL, SH Obtained funding: Not applicable Overall responsibility: AF REFERENCES 1. Zheng ZZ, Liu ZX. Activation of the phosphatidylinositol 3-kinase/ protein kinase Akt pathway mediates CD151-induced endothelial cell proliferation and cell migration. Int J Biochem Cell Biol 2007;39: 340-8. 2. Hockel M, Schlenger K, Doctrow S, Kissel T, Vaupel P. Therapeutic angiogenesis. Arch Surg 1993;128:423-9. 3. Annex BH. Therapeutic angiogenesis for critical limb ischaemia. Nat Rev Cardiol 2013;10:387-96. 4. Strioga M, Viswanathan S, Darinskas A, Slaby O, Michalek J. Same or not the same? Comparison of adipose tissue-derived versus bone marrow-derived mesenchymal stem and stromal cells. Stem Cells Dev 2012;21:2724-52. 5. Ma S, Xie N, Li W, Yuan B, Shi Y, Wang Y. Immunobiology of mesenchymal stem cells. Cell Death Differ 2014;21:216-25. 6. Kwon HM, Hur SM, Park KY, Kim CK, Kim YM, Kim HS, et al. Multiple paracrine factors secreted by mesenchymal stem cells contribute to angiogenesis. Vascul Pharmacol 2014;63:19-28. 7. Liu W, Morschauser A, Zhang X, Lu X, Gleason J, He S, et al. Human placenta-derived adherent cells induce tolerogenic immune responses. Clin Trans Immunology 2014;3:e14. 8. Shehadah A, Chen J, Pal A, He S, Zeitlin A, Cui X, et al. Human placenta-derived adherent cell treatment of experimental stroke promotes functional recovery after stroke in young adult and older rats. PLoS One 2014;9:e86621.
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9. He S, Khan J, Gleason J, Eliav E, Fik-Rymarkiewicz E, Herzberg U, et al. Placenta-derived adherent cells attenuate hyperalgesia and neuroinflammatory response associated with perineural inflammation in rats. Brain Behav Immun 2013;27:185-92. 10. Mayer L, Pandak WM, Melmed GY, Hanauer SB, Johnson K, Payne D, et al. Safety and tolerability of human placenta-derived cells (PDA001) in treatment-resistant Crohn’s disease: a phase 1 study. Inflamm Bowel Dis 2013;19:754-60. 11. Ribatti D, Vacca A, Roncali L, Dammacco F. The chick embryo chorioallantoic membrane as a model for in vivo research on angiogenesis. Int J Dev Biol 1996;40:1189-97. 12. Goto T, Fukuyama N, Aki A, Kanabuchi K, Kimura K, Taira H, et al. Search for appropriate experimental methods to create stable hind-limb ischemia in mouse. Tokai J Exp Clin Med 2006;31:128-32. 13. Li X, Ling W, Pennisi A, Wang Y, Khan S, Heidaran M, et al. Human placenta-derived adherent cells prevent bone loss, stimulate bone formation, and suppress growth of multiple myeloma in bone. Stem Cells 2011;29:263-73. 14. Belch J, Hiatt WR, Baumgartner I, Driver IV, Nikol S, Norgren L, et al. Effect of fibroblast growth factor NV1FGF on amputation and death: a randomised placebo-controlled trial of gene therapy in critical limb ischaemia. Lancet 2011;377:1929-37. 15. Khan TA, Sellke FW, Laham RJ. Gene therapy progress and prospects: therapeutic angiogenesis for limb and myocardial ischemia. Gene Ther 2003;10:285-91. 16. Caplan AI, Correa D. The MSC: an injury drugstore. Cell Stem Cell 2011;9:11-5. 17. Murphy MB, Moncivais K, Caplan AI. Mesenchymal stem cells: environmentally responsive therapeutics for regenerative medicine. Exp Mol Med 2013;45:e54. 18. Li Y, Chopp M. Marrow stromal cell transplantation in stroke and traumatic brain injury. Neuroscience Lett 2009;456:120-3. 19. Liew A, O’Brian T. Therapeutic potential for mesenchymal stem cell transplantation in critical limb ischemia. Stem Cell Res Ther 2012;3:28. 20. Bronckaers A, Hilkens P, Martens W, Gervois P, Ratajczak J, Struys T, et al. Mesenchymal stem/stromal cells as a pharmacological and therapeutic approach to accelerate angiogenesis. Pharmacol Ther 2014;143:181-96. 21. DiMarino AM, Caplan AI, Bonfield TL. Mesenchymal stem cells in tissue repair. Front Immunol 2013:e201. 22. Xu KP, Yu FS. Cross talk between c-Met and epidermal growth factor receptor during retinal pigment epithelial wound healing. Invest Ophthalmol Vis Sci 2007;48:2242-8. 23. Shimamura M, Sato N, Waguri S, Uchiyama Y, Hayashi T, Iida H, et al. Gene transfer of hepatocyte growth factor gene improves learning and memory in the chronic stage of cerebral infarction. Hypertension 2006;47:742-51. 24. Yang ZJ, Zhang YR, Chen B, Zhang SL, Jia EZ, Wang LS, et al. Phase I clinical trial on intracoronary administration of Ad-hHGF treating severe coronary artery disease. Mol Biol Rep 2009;36:1323-9. 25. Trapp T, Kogler G, El-Khattouti A, Sorg RV, Besselmann M, Focking M, et al. Hepatocyte growth factor/c-MET axis-mediated tropism of cord blood-derived unrestricted somatic stem cells for neuronal injury. J Biol Chem 2008;283:32244-53. 26. Kaga T, Kawano H, Sakaguchi M, Nakazawa T, Taniyama Y, Morishita R. Hepatocyte growth factor stimulated angiogenesis without inflammation: differential actions between hepatocyte growth factor, vascular endothelial growth factor and basic fibroblast growth factor. Vascul Pharmacol 2012;57:3-9. 27. Morishita R, Makino H, Aoki M, Hashiya N, Yamasaki K, Azuma J, et al. Phase I/IIa clinical trial of therapeutic angiogenesis using hepatocyte growth factor gene transfer to treat critical limb ischemia. Arterioscler Thromb Vasc Biol 2011;31:713-20. 28. Powell RJ, Simons M, Mendelsohn FO, Daniel G, Henry TD, Koga M, et al. Results of a double-blind, placebo-controlled study to assess the safety of intramuscular injection of hepatocyte growth factor plasmid to improve limb perfusion in patients with critical limb ischemia. Circulation 2008;118:58-65. 29. Ebrahem Q, Minamoto A, Hoppe G, Anand-Apte B, Sears JE. Triamcinolone acetonide inhibits IL-6- and VEGF-induced
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angiogenesis downstream of the IL-6 and VEGF receptors. Invest Ophthalmol Vis Sci 2006;47:4935-41. 30. Heidemann J, Ogawa H, Dwinell MB, Rafiee P, Maaser C, Gockel HR, et al. Angiogenic effects of interleukin 8 (CXCL8) in human intestinal microvascular endothelial cells are mediated by CXCR2. J Biol Chem 2003;278:8508-15. 31. Motz GT, Coukos G. The parallel lives of angiogenesis and immunosuppression: cancer and other tales. Nat Rev Immunol 2011;11: 702-11. 32. Madonna R, Delli Pizzi S, Di Donato L, Mariotti A, Di Carlo L, D’Ugo E, et al. Non-invasive in vivo detection of peripheral limb
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ischemia improvement in the rat after adipose tissue-derived stromal cell transplantation. Circ J 2012;76:1517-25. 33. Guller S, LaChapelle L. The role of placental Fas ligand in maintaining immune privilege at maternal-fetal interfaces. Semin Reprod Endocrinol 1999;17:39-44.
Submitted Nov 12, 2014; accepted Apr 4, 2015.
Additional material for this article may be found online at www.jvascsurg.org.
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APPENDIX (online only) As part of the Celgene Cellular Therapeutics (CCT) Research Group, the following group members significantly contributed to the work described in this manuscript: Aleksandr Kaplunovsky, Jennifer Paredes, Allan Reduta, Eric Law, Ewa Fik, Sascha Abramson, Vivian R. Albert, and Itschak Lamensdorfa (aPharmaseed Ltd). SUPPLEMENTARY METHODS (online only) Preparation of placenta-derived adherent cells (PDACs), human umbilical vein endothelial cells (HUVECs), human dermal fibroblast (HDF), and human mesenchymal stromal cells (MSCs). PDACs were prepared by mechanical and enzymatic digestion of human placental tissue of newborn origin obtained from a normal, full-term birth as described.7 PDACs were expanded until passage 6 in PDAC medium,7 which, unless otherwise indicated, contains 2% fetal bovine serum (Invitrogen, Carlsbad, Calif). Cryopreserved PDACs (passage 6 at 7.5 106 or 20 106 cells/mL in 5% dimethyl sulfoxide, 10% human serum albumin, and 5.5% dextran in 0.9% NaCl) were thawed in a 37 C water bath. Viability and cell counts were assessed by trypan blue exclusion; cells were used within 4 hours of thaw. Flow cytometric analysis of PDACs. Flow cytometric surface marker staining of PDACs was performed with labeled monoclonal mouse antihuman immunoglobulin G (BD Biosciences, San Jose, Calif). For surface marker expression analysis, PDAC single-cell suspensions were washed with fluorescence-activated cell sorting (FACS) wash buffer (phosphate-buffered saline with 2% fetal bovine serum) and labeled with antibodies by incubation at 4 C for 30 minutes. Flow cytometry analysis was performed on a FACSCanto II (BD Biosciences). Data were acquired with FACSDiva software (BD Biosciences) and analyzed with FlowJo flow cytometry software (Tree Star, Ashland, Ore). HUVEC survival assay. HUVECs were subjected to serum starvation in the presence and absence of PDACs, HDF, or MSCs, using indirect coculture systems. HUVECs were seeded (2 104 cells/well) in a 24-well tissue culture-treated polystyrene precoated with fibronectin (Sigma-Aldrich, St. Louis, Mo). After overnight incubation in complete endothelial cell growth medium 2 (Lonza, Walkersville, Md) to facilitate cell attachment and proliferation, HUVECs were serum starved in endothelial cell basal medium (Lonza) for 6 hours. Concomitant with HUVEC seeding, 0 to 5 104 PDACs, HDF, or MSCs were independently seeded into 8-mm-pore Transwell culture inserts (Corning, Lowell,
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Mass) in a separate multiwell plate in PDAC medium and incubated 2 hours to allow attachment. Subsequently, PDAC medium was replaced with serum-free Dulbecco’s modified Eagle medium (DMEM, Invitrogen) for 24 hours to prepare cell-conditioned medium (CM) containing factors secreted by PDACs, HDF, or MSCs. The following day, HUVEC culture medium was replaced with PDACCM. In addition, PDACs, HDF, or MSCs Transwell inserts were placed into the HUVEC multiwell plates for indirect coculture between PDACs, HDF, or MSCs and HUVECs overlaid with PDAC-CM. After 24 hours, the Transwell cocultures and media were removed, and HUVEC counts were determined with the CellTiter-Glo assay (Promega, Madison, Wisc). Tube formation assay. HUVECs were cultured to passage 3 in endothelial cell growth medium 2 for 3 days to w75% confluency. HUVECs were washed and resuspended in DMEM and diluted to 4000 cells/mL with human placental collagen (1.5 mg/mL in DMEM). After pipetting (3 mL/well) into 96-well plates (n ¼ 5 per condition), HUVEC/human placental collagen drops were incubated for 90 minutes (37 C, 5% CO2, and 90% relative humidity) to facilitate collagen solidification, followed by addition of 200 mL of PDAC-CM or control medium. After incubation (24 hours), HUVEC drops were stained with a Diff-Quik Cell Staining Kit (Dade Behring, Inc, Newark, Del). Images of each well were acquired with the Zeiss SteREO Discovery V8 microscope (Carl Zeiss AG, Jena, Germany) and analyzed for average network area. Chicken embryo chorioallantoic membrane assay. Chorioallantoic membrane assays in fertilized chicken eggs were used to assess neovascularization and were performed by SRI International (Menlo Park, Calif) as described.11 The test articles (n ¼ 8-10) applied on day 6 were PDACs (7.7 105 viable cells) in 40 mL of DMEM/Matrigel vehicle mixture (1:1), positive control (100 ng/mL basic fibroblast growth factor in DMEM/ Matrigel mixture), Matrigel vehicle control, and DMEM control. On day 8, blood vessel density was determined by an image capturing system at a magnification of 100. Unilateral hindlimb ischemia induction and administration of PDACs. The surgical procedure followed the Prox-A protocol.12 Under anesthesia, an incision (mice, 0.5-1.0 cm; rats, 1.0-2.0 cm) was made in the skin in the inguinal area. The femoral artery was ligated twice with 6-0 (mice) or 4-0 (rats) silk thread and transected between the ligatures. The wound was closed with 4-0 (mice) or 3-0 (rats) silk thread, and the animal was allowed to recover.