Sourcing of an Alternative Pericyte-Like Cell Type from Peripheral Blood in Clinically Relevant Numbers for Therapeutic Angiogenic Applications

Abstract

Autologous cells hold great potential for personalized cell therapy, reducing immunological and risk of infections. However, low cell counts at harvest with subsequently long expansion times with associated cell function loss currently impede the advancement of autologous cell therapy approaches. Here, we aimed to source clinically relevant numbers of proangiogenic cells from an easy accessible cell source, namely peripheral blood. Using macromolecular crowding (MMC) as a biotechnological platform, we derived a novel cell type from peripheral blood that is generated within 5 days in large numbers (10–40 million cells per 100 ml of blood). This blood-derived angiogenic cell (BDAC) type is of monocytic origin, but exhibits pericyte markers PDGFR-β and NG2 and demonstrates strong angiogenic activity, hitherto ascribed only to MSC-like pericytes. Our findings suggest that BDACs represent an alternative pericyte-like cell population of hematopoietic origin that is involved in promoting early stages of microvasculature formation. As a proof of principle of BDAC efficacy in an ischemic disease model, BDAC injection rescued affected tissues in a murine hind limb ischemia model by accelerating and enhancing revascularization. Derived from a renewable tissue that is easy to collect, BDACs overcome current short-comings of autologous cell therapy, in particular for tissue repair strategies.

Introduction

Re-establishing microvasculature is an important goal in regenerative medicine.¹ A variety of cell types, mainly derived from bone marrow, have been examined for their suitability to achieve this therapeutic goal. In this regard, autologous cells were used to minimize immunological and risk of infections.^2,3,4 However, low yields of patient-specific functional cells necessitate prolonged cell expansion times in vitro. This is associated with increased costs and potential impairment of cellular functionality, thus diminishing the optimism associated previously with autologous cell therapy.⁵ The greatest challenge lies in the expansion of harvested cells within a therapeutically useful time window while controlling costs. Here, we consider peripheral blood as a renewable cell source that can be harvested with relative ease. Our focus is on monocytes, which represent 3–7% of all blood leukocytes, and differentiate spontaneously into macrophages upon isolation from the circulation.⁶

Macrophages are comprised of several subtypes (i.e., M1 and M2) that play different physiological roles, one of them being supporting angiogenesis.⁷ The M2-subtype is indispensable for directing and enhancing angiogenesis in both embryonic development^8,9,10 and adulthood.¹¹ It has been demonstrated that macrophages migrate within the embryo towards areas of future vascularization, where they direct vessel fusion during vascular network formation.^8,9 Proangiogenic macrophages have been also identified in various solid tumors^12,13,14,15 and have been suggested to be responsible for the angiogenic switch in tumors.¹⁶

Studies focusing on peripheral blood as a source of reparative cells have described monocyte-derived cells with fibroblastic characteristics, termed fibrocytes,^16,17,18,19 and endothelial-like cells (ELC),^20,21,22,23 which are proangiogenic in vitro and in vivo.

Several reports have found that pericytes during early stages of angiogenesis, identified by their perivascular location and the expression of pericytic markers such as PDGFR-β and NG2,^{24,25,26,27,28,29} also expressed leukocyte and monocyte markers. However, this observation is not congruent with the current opinion that pericytes are in essence mesenchymal stem cells (MSCs) residing in perivascular niches.^{30,31,32,33,34} Obviously, MSCs lack hematopoietic markers³⁵ and specifically, MSC-like pericytes do not express hematopoietic and monocytic markers when freshly isolated using CD146 selection, and after in vitro expansion.^30,36 In order to explain this discrepancy, we hypothesize the existence of at least two pericytic populations. One population would be of hematopoietic origin and may be found around newly forming vessels during early angiogenesis and contributing to vessel sprouting, whereas MSC-like pericytes enter at a later angiogenic stage to promote vessel stabilization and maturation. After completion of angiogenesis MSC-like pericytes would remain attached to the blood vessels and support their maintenance, whereas hematopoietic pericytes go on to retreat. Previously, we have introduced macromolecular crowding (MMC) as a novel cell culture technique to modulate cellular characteristics. MMC affects the biophysical state of a cell culture system by excluding volume, and thereby increasing the effective concentration, availability and efficacy of macromolecules such as growth factors³⁷ and also drives extracellular microenvironment formation.^37,38,39,40

Here, we describe the use of MMC to generate blood-derived angiogenic cells (BDACs) with pericyte characteristics from cultured buffy coats, in large numbers in a short period of time. Furthermore, we demonstrate that these cells have angiogenic potential and exert a strong therapeutic effect in a preclinical model of critical limb ischemia.

Results

Pulsing of mononuclear cell fraction with MMC results in a proliferative burst of adherent spindle-shaped cells

Peripheral blood mononuclear cells (PBMCs) were cultured in low glucose DMEM supplemented with 10% FBS on fibronectin-coated dishes, either in the absence (−MMC) or presence of a sucrose copolymer cocktail containing Ficoll 70 kDa (Fc70) and 400 kDa (Fc400) (+Fc) (Figure 1). The concentration for the Ficoll cocktail was optimized previously to have a fraction volume occupancy of 17% resembling physiological crowding conditions.²³ Macromolecular polymers as well as nonadherent cells were removed after 1 day through medium change, and adherent cells were cultivated in DMEM/FBS alone for further 4 days (Figure 1a,b).

Figure 1.

MMC induces BDAC proliferation and enhances growth factor signaling in monocyte-derived cultures. Spindle-shaped cells were generated from PBMCs without (−MMC), or in the presence of macromolecules (+Fc) during the first day of culture. Increased proliferation of adherent spindle-shaped cells under +Fc is evident (a) in phase contrast (PhC) images at day 5 (b) adherent cytometry of DAPI stained nuclei of adherent cells resulting from cell culture of 10⁶ PBMC at 6 hours (0.25 days), 1 day, 2 days and 5 days and (c) Quantitation of cell doublings during 5 days of culture. (d–e) MTS assay of spindle-shaped cells normalized to cell number on day 1 and 5, respectively. NADH/NADPH levels are displayed as percentages relative to –MMC controls. (f–g) G6PDH activity assay normalized to cell number on day 1 and 5, respectively. (h) M1 polarization of THP-1 derived macrophages under MMC (+Fc) or without macromolecules (−MMC). Read-out for successful polarization is TNF-α production. *P < 0.05; **P < 0.003. All experiments were done at least in triplicates with comparable results.

Spindle-shaped cells appeared under both conditions; –MMC and +Fc. However, under MMC, cells grew to a considerable length of ~100 µm (Figure 1a, +Fc), while the majority of cells in the control media appeared shorter in comparison (Figure 1a, −MMC). Adherent cytometry of DAPI-stained cells at different time points, revealed that MMC increased the number of adherent cells by almost twofold already after 6 hours (Figure 1b and Supplementary Figure S1). Furthermore, a MMC pulse during the first 24 hours resulted in almost three times more cells than in untreated controls at the end of the culture period (Figure 1b and Supplementary Figure S1). Quantification of cell doubling times over the 5 days of culture revealed that cells generated under MMC were more proliferative (Figure 1c and Supplementary Figure S1). Extrapolating from a 50 ml blood sample, we estimate that this protocol would be able to generate 40–160 million BDAC from one blood donation (400 ml) within 5 days. These elongated adherent cells generated under MMC demonstrated angiogenic properties, and are henceforth referred to as “blood derived angiogenic cells” (BDACs). Unless stated otherwise BDACs generated in the presence of MMC were used for all subsequent experiments.

MMC induces BDAC proliferation by switching metabolism from citrate cycle to the pentose phosphate pathway

Despite a significant proliferative burst of BDACs (Figure 1c), MTS assays showed only a slightly increased activity of mitochondrial oxidoreductases under MMC on day 1, when normalized for cell numbers (Figure 1d). However, on day 5, the normalized activity of mitochondrial oxidoreductases decreased significantly when compared to controls (Figure 1e). In contrast, the activity of glucose-6-phosphate dehydrogenase (G6PDH), a key-enzyme of the pentose phosphate pathway (PPP), was increased under MMC on days 1 and 5 (Figure 1f,g), indicating an anabolic switch towards cell mass production.

MMC enhances cytokine signaling in monocyte-derived cells

We generated BDACs from crude buffy coats to allow for a complex interaction between monocytes and other mononuclear cells. To dissect the effects of MMC on cytokine interactions, we employed as a pars pro toto model macrophages derived from THP-1 cells, which were polarized with LPS and various concentrations of IFN-γ into an M1 phenotype under either –MMC or +Fc environment. TNF-α secretion served as read-out for successful polarization (Figure 1h). While MMC alone did not act as a polarizing factor (Figure 1h, control), it enhanced LPS-induced polarization into M1 in the absence and at all IFN-γ concentrations (Figure 1h). Interestingly, the largest MMC effects were found at low IFN-γ concentrations, indicating a partial saturation of M1 polarizing signals at higher concentrations (Figure 1h). This suggests that MMC enhances cytokine signaling in monocyte-derived cells.

BDACs exhibit a unique marker profile

BDACs showed a characteristic surface marker profile (Figure 2). Flow cytometric analysis revealed strong expression of leukocyte marker CD45, and monocyte/macrophage marker CD11b and CD14 (Figure 2a). This indicates that BDACs are derived from the monocytic lineage. Interestingly, these cells lacked MHC II receptor HLA-DR, but strongly expressed M2 marker CD206 (Figure 2a). Dendritic and B-cell marker CD83 and CD19 were absent (Figure 2a). Although BDACs did not express the hematopoietic progenitor markers CD34 or CD133, they showed a strong expression of c-kit (CD117) (Figure 2a). From an array of potential endothelial cell markers, only VEGFR-2 was expressed, with some batch-to-batch variance. CD146 and VE-cadherin (CD144) were absent (Figure 2a). Therefore, BDACs did not have an endothelial phenotype (and thus can be distinguished from ELC) but were also significantly distinct from mesenchymal progenitors, expressing CD29, CD105, CD13, and CD166, while being negative for CD73 and CD90 (Figure 2a). Spindle-shaped cells generated under noncrowded conditions were analyzed for a range of selected markers. They displayed comparable levels of CD45, CD11b, CD206, and CD105. However, they expressed higher levels of CD14 and HLA-DR and lower levels of VEGFR-2 (Supplementary Figure S2).

Figure 2.

BDACs express a unique set of markers and are distinguishable from other angiogenic cells. (a) Flow cytometry analysis for hematopoietic, endothelial, and mesenchymal markers. Results are displayed as average ± SD from three independent blood samples. (b) Immunocytochemistry for vWF in BDAC and HUVEC cultures. (c) SDS-Page of pepsin-digested conditioned media by Pl-Prc, IMR-90s (fibroblasts), and BDACs. Collagen I standards (Col I Std) served as molecular weight standards. Bands from the same gel are displayed. (d) RT-PCR of collagen α1 and vWF mRNAs in Pl-Prc, IMR-90s, HUVEC and BDACs. (e) Immunocytochemistry for Tie-2, VEGFR-1, CD31, and antigen recognized by PM-2k antibody clone in HUVECs, BDAC, Pl-Prc, and bmMSCs. (f) Immunocytochemistry for PDGFR-β, NG2, α-SMA and desmin in cell types from e. All experiments were repeated at least three times with comparable results.

Fibrocytes and ELCs are two other spindle-shaped cell types derived from monocytes ascribed with angiogenic properties. We tested BDACs for ELC marker vWF,¹⁶ and the fibrocyte marker collagen I^16,17 (Figure 2b–d). Immunocytochemical analysis for vWF showed punctate staining indicative of contaminating platelets, while human umbilical cord endothelial cells (HUVEC) as controls showed strong perinuclear staining (Figure 2b, Supplementary Figure S3a).

Collagen I has been described as a distinct marker for fibrocytes.^7,17 Therefore, BDAC were cultivated in parallel with fibroblasts (IMR-90s) and placenta-derived pericytes (Pl-Prc) in the presence of ascorbic acid. SDS-Page analysis of pepsin-digested culture media demonstrated that IMR-90 and Pl-Prc secreted substantial amounts of collagen I, while BDACs did not (Figure 2c). Furthermore, RT-PCR with primers specific for vWF and COL1A1, showed significant collagen I mRNA levels only in IMR-90s and Pl-Prc, while significant vWF mRNA levels were observed only in HUVEC (Figure 2d). This confirmed that BDACs have a phenotype distinctive from fibrocytes and ELC.

BDACs express angiogenesis- and pericyte-related markers

BDACs were assessed immunocytochemically for angiogenesis-related markers. HUVEC, Pl-Prc, and bmMSCs served as controls for the markers examined. Conjugate controls for all cell types can be found in Supplementary Figure S3a. Along with HUVECs, they consistently expressed Tie-2, VEGFR-1, and CD31, although intensities varied from batch to batch (Figure 2e). Only BDACs showed a positive staining by the antibody clone PM-2k, confirming their monocytic origin (Figure 2e). Interestingly, Pl-Prc shared Tie-2 and VEGFR-1 expression with BDACs and HUVEC, although with weaker intensity, whereas bone marrow (bm) MSCs did not express any of these markers.

With regards to pericyte-related markers, BDACs were positive for PDGFR-β and NG2, but not for α-SMA (Figure 2f). Desmin expression varied from batch to batch (Figure 2f). Endothelial cells, which served as a negative control, did not express any of these pericyte-related markers (Figure 2f). Pl-Prc, which served as a positive control expressed all of these markers, although α-SMA and desmin immunostaining were variable (Figure 2f). BmMSCs, shared PDGFR-β and α-SMA expression with Pl-Prc, indicating the lack of specificity of this markers for pericytes (Figure 2f). Of note, when PBMCs were cultured without MMC, the resultant spindle-shaped cells lacked most of angiogenesis and pericyte-related markers (Supplementary Figure S4). Therefore, MMC contributes significantly to the generation and characteristics of BDACs, and their differential pericytic phenotype. When HUVEC, Pl-Prc, and bmMSCs were subjected to the same culture protocol as PBMCs, MMC did not affect the expression of any of the tested markers (Supplementary Figure S5).

BDACs do not exhibit standard macrophage function

To capture the substantial plasticity of monocyte-derived cells, a classification spectrum with two extremes of macrophage polarization has been proposed, namely M1 (inflammatory) and M2 (alternative).^7,41 We therefore generated BDACs and macrophages from the same blood samples and polarized them into either M1 or M2 using standard protocols.⁴¹ The results showed high variability, especially for macrophages, but all blood samples displayed the same trends: M1 polarization of macrophages resulted in a high secretion of TNFα (Figure 3a), whereas M2 polarization did not induce an elevated secretion of IL-10, an antiinflammatory M2 cytokine (Figure 3b). However, M2 polarization led to CD206 expression (Figure 3c, Supplementary Figure S3b). In contrast, BDACs responded much more weakly to M1 polarization protocols with much lower levels of TNF-α secretion (Figure 3a). BDACs also always secreted lower levels of IL-10 in comparison to macrophages (polarized or unpolarized), and furthermore M2 polarization of BDACs resulted in a decrease of IL-10 (Figure 3b). BDACs also showed a stable expression of CD206, although the cell morphology altered during polarization (Figure 3c, Supplementary Figure S3b). This indicated that BDAC might represent an alternatively activated (M2) monocyte-derived cell type with a stable (less plastic) phenotype. When subjected to a phagocytosis assay, only 10 ± 6% of BDACs took up fluorescent microbeads similar to HUVEC (9 ± 6%), which served as a negative control (Figure 3d–f). In contrast, 52 ± 4% of macrophages generated from the same blood samples exhibited phagocytosis (Figure 3d–f). Therefore, BDACs, although monocyte-derived, did not perform standard macrophage functions.

Figure 3.

Although monocyte-derived, BDACs differ from standard macrophages. (a–c) BDACs and macrophages derived from the same blood samples were polarized into either M1 or M2. (a) Graph shows ELISA results for M1 cytokine TNFα measured in supernatant. (b) ELISA for M2 cytokine IL10 in conditioned media. (c) Immunocytochemistry for CD206 expression in cell types from a) and b). (d–f) Phagocytosis assay. (d) Quantification of cells up-taking fluorescent microbeads by flow cytometry. Data are presented as average of three independent blood samples. HUVEC served as negative controls. (e) Exemplary flow cytometry results with (f) corresponding images of cells before harvesting (Phase contrast images (PhC) overlaid with green fluorescence are displayed). Scale bar 100 µm.

BDACs colocalize with endothelial tubular networks on matrigel and stabilize them

When cocultured with HUVEC on matrigel, BDACs colocalized with endothelial tube networks immediately at formation, and remained firmly attached as long as a network could be observed (Figure 4a, Supplementary Video S1). In particular, some cells were observed to move along endothelial tubes from junction point to junction point (Figure 4a, arrows, Supplementary Video S1). Over a time course of 24 hours, labeled cells accumulated at junction points (Figure 4a,b). We measured the total tube length in the field of view taken in order to estimate the stability of the network over time. Although no difference in total tube length between HUVEC controls and HUVEC in coculture with BDACs was observed during the initial formation of the network, quantification of total tube length at 16 hours after seeding revealed that more of the tubular network was sustained in the presence of BDAC and therefore BDACs were able to stabilize these networks (Figure 4c,d). BDACs did not form a network on their own.

Figure 4.

BDACs colocalize and stabilize endothelial tubular networks on matrigel. (a) Stills of live-cell imaging of coculture of BDAC (green label) with HUVEC on matrigel over 24 hours (Supplementary Video S1). (b) 40× magnification of BDAC (green label)-HUVEC coculture on matrigel. Phase contrast (PhC) and green fluorescence are displayed. Bar: 50 µm. (c) Culture of HUVECs alone or HUVECs together with BDAC on matrigel in starving media at 4 hours and 16 hours. Bar: 500 µm. (d) Quantification of cumulative tube length per field of view (FOV) at 16 hours of cultures in c. All experiments were repeated at least three times with comparable results.*P < 0.02.

BDACs colocalized with endothelial sprouts and enhanced endothelial sprouting in vitro

We tested the hypothesis that BDACs may enhance endothelial sprouting at an early stage of angiogenesis. Fluorescently labeled BDACs were cocultured with HUVEC to form mixed spheroids, which were subsequently suspended in a collagen I gel. Endothelial cells budded out and formed sprouts within the collagen I gel. Interestingly, BDACs were not only found in the spheroid cores, but also at the sprouting points (Figure 5a). Spheroids solely composed of BDACs were not stable and often disintegrated during the transfer into collagen I gels. In subsequent experiments and to ensure comparable sizes, spheroids containing only HUVEC were embedded in collagen I gels, which contained different concentrations of BDAC suspended as single cells. We observed that the cumulative sprout length per spheroid increased proportionately to the concentration of BDACs present as a suspension in the gel (Figure 5b and Supplementary Figure S6). Of note, spindle-shaped cells generated under noncrowded conditions did not enhance endothelial sprouting in vitro (Supplementary Figure S7). In BDAC-free gels, sprouts consisted of loosely attached endothelial cells and often disintegrated. In contrast, the presence of BDACs induced formation of smooth, continuous sprouts (Figure 5b,c).

Figure 5.

BDACs colocalize with endothelial sprouts and enhance endothelial sprouting in a cell-dose dependent manner in vitro. (a) Spheroids containing HUVEC and BDAC (green label) in a ratio of 2:1 were embedded into a collagen I gel and allowed to sprout for 3 days. Bar: 50 µm. (b) HUVECs spheroids were embedded in a collagen I gel with various BDAC concentrations suspended. Graphs show measured cumulative sprout length/spheroid and representative phase contrast (PhC) pictures are shown below. Bar: 100 µm. (c) Close-ups of sprouts for HUVEC control condition and in the presence of 2.7 x 10⁵ BDAC suspended per ml of collagen I. All experiments were repeated at least three times with comparable results.*P < 0.01; **P < 0.00001.

BDACs generated are proangiogenic in vivo

To assess the angiogenic potential of BDACs in vivo, these cells were fluorescently labeled and coinjected subcutaneously with human glioma cells (U87) in nude mice. In contrast to other in vivo revascularization models, where ischemia leads to necrosis and inflammation, this tumor model allows the observation of solely angiogenic events. A subcutaneous tumor mass is vascularized by ingrowth of vessels from the periphery. Therefore it is possible to distinguish easily between areas with different degrees of vascularization, and the localization of the cells of interest within these areas. Tumors were harvested after 8 days. At this early stage of tumor growth, no significant differences in tumor weight were observed. Tumors were snap frozen and cryosectioned. Methanol-fixed sections (causing a loss of BDAC labeling) were immunostained with antibodies against CD31 and vWF. Quantification of fluorescent areas per tumor section proved that tumors containing BDACs had 30–50% larger areas staining positively for vWF and CD31, respectively (Figure 6a,b and Supplementary Figure S3c). As expected at this early time point of tumor harvest, tumors were most vascularized in the periphery and still avascular in the tumor center.

Figure 6.

BDACs are proangiogenic in vivo. U87 human glioma cells were injected subcutaneously into nude mice, either alone (U87 control) or with BDACs (U87 + BDAC) (n = 5). Animals were sacrificed after 8 days and tumors analyzed. (a) Immunohistochemical analysis of CD31 staining. Graph: Total fluorescent area per tumor section was quantified. Representative images are displayed below. (b) Immunohistochemical analysis of vWF staining. Graph: Total fluorescent area per tumor section was quantified. Representative images are displayed below. (c) Images of labeled BDACs (red) and CD31 stained blood vessels (green staining) at various locations within a tumor. DAPI was used to visualize nuclei staining (blue). Arrows: BDACs in perivascular location. Bar: 100 µm *P < 0.05.

When tumor slices from the same tumors were fixed with formaldehyde, instead of methanol, the fluorescent red label of BDAC was retained. BDAC location before fixation (Supplementary Figure S8a) and with respect to stained blood vessels could be visualized in the tumor (Figure 6c and Supplementary Figures S3d and S8b). Most of the BDAC staining was found in tumor centers, which were still avascular at this early time point, and at areas of angiogenesis (junction of still-avascular tumor center and vascularized tumor periphery). Highly vascularized tumor areas found in the tumor periphery were devoid of BDACs (Figure 6c and Supplementary Figure S8b). At areas of angiogenesis, CD31 staining revealed presence of BDACs in both perivascular locations as expected for pericytes (Figure 6c, white arrows) as well as in the tumor stroma (Figure 6c).

MMP9 secreted by BDACs is a pivotal supporter of endothelial sprouting

Culture media conditioned by BDACs was analyzed for secreted angiogenic factors. A proteome array revealed presence of extracellular matrix proteases MMP-9, uPA and their inhibitors TIMP and PAI-I, respectively (Figure 7a). In addition, angiogenic factors were identified, including GM-CSF, VEGF, and IL-8 (Figure 7a); all are known to support endothelial cell survival, migration and proliferation.^42,43,44,45 Furthermore, HB-EGF and MCP-1 (Figure 7a) were identified; both factors are proangiogenic in vivo.^{29,46,47,48,49} Interestingly, BDACs lacked antiangiogenic factors, such as Activin A, TGFβ-1, IL-1, or thrombospondin, except CXCL4 (PF4) (Figure 7a, Supplementary Figure S9), which we believe to come from contaminating platelets (compare Figure 2b (vWF), Figure 2h (CD31)).

Figure 7.

BDACs secrete proangiogenic factors involved in early stages of angiogenesis, of which secreted MMP9 is a pivotal effector in the enhancement of endothelial sprouting in vitro. (a) Proteome array for various pro- and antiangiogenic factors. Only strong signals are labeled on the membrane. Quantifications of signals can be found in Supplementary Figure S9. (b) Gelatin and collagen zymography of gel samples from spheroid sprouting assay (from Figure 5b), from HUVECs and BDACs suspended as single cells in collagen I gels and from supernatant of monolayer cultures of HUVEC and BDACs. Western blot (WB) for MMP-9. (c) Inhibition of MMP9 by small molecules in spheroids sprouting assay. (d) Representative images of spheroids from c. All experiments were repeated at least three times with comparable results.*P < 0.003.

Gelatin and collagen I zymography confirmed that cultures in which endothelial cells sprouted in the presence or absence of BDAC, contained MMP activity associated with bands migrating at 60 and 80 kDa (Figure 7b). Monocultures of BDAC suspended in collagen I gels secreted MMPs mainly at 80 kDa, whereas endothelial monocultures at corresponding cell density mainly secreted MMPs at 60 kDa, although the band indicated smaller amounts of MMP (Figure 7b). Immunoblotting confirmed the identity of the 80 kDa band as MMP9 (Figure 7b). Therefore, it appears plausible that BDAC-derived MMP9 plays a major role in the enhancement of endothelial sprouting. In order to confirm this hypothesis, we demonstrated that the addition of an MMP9 inhibitor abrogated the BDAC-mediated enhancement of endothelial sprouting significantly (Figure 7c,d).

BDAC-treatment accelerates and enhances revascularization in a murine hind limb ischemia model

BDAC's proangiogenic properties were explored in a model of tissue repair to test their efficacy in a clinically relevant ischemia model. For this reason, the external iliac artery was ligated in the left limb of balb/c nude mice and BDACs or just the vehicle were injected intramuscularly. Complete ligation was confirmed and revascularization was observed by MRI angiography of the operated leg over 4 weeks. Representative reconstructed images of vessels taken by MRI are displayed in Figure 8a and quantifications of the total vessel volume in the region of interest (ROI) are shown in the graph in Figure 8b. MRI angiography 1 day after surgery confirmed no detectable blood vessels in the limb and therefore complete ligation of the iliac artery (Figure 8a,b). Revascularization was observed in both treatment groups (vehicle-injected and BDAC-injected), however already after 1 week a significantly higher vessel volume was measured in the BDAC-treated group (Figure 8a,b). During the second week, the vessel volume exceeded that of the baseline only in the BDAC group. At weeks 3 and 4, the ischemia appeared to be resolved in both experimental groups as vessel volume was comparable to the baseline before surgery (Figure 8a,b). All vehicle-treated animals showed necrosis of the toes and one animal needed to be sacrificed prior to the end of the study due to progressive foot necrosis (Figure 8c–e). Representative images at week 2 are shown in Figure 8c and a semiquantification following a scoring (Figure 8d) summarizing the observed ischemia is displayed in graph Figure 8e. In the BDAC-treated group, all animals except one were completely free of necrosis. The exception animal showed a necrosis of one toe during the first week and the necrosis did not progress any further during the study. We also observed functional recovery of the lower limb as evident by improved grabbing in the BDAC-treated group. Histological analysis of the calf muscles revealed fibrosis in the center of the muscle section, neutrophil infiltration and muscle degeneration with adipose replacement in all experimental animals (Figure 8f,g). Fiber regeneration, characterized by muscles fibers with a centralized nucleus could also be observed. However, in accordance with phenotypic changes of the lower limb, a higher degree of neutrophil infiltration and muscle degeneration with adipose replacement were observed in the vehicle-treated group (Figure 8f,g).

Figure 8.

BDACs accelerate and enhance revascularization in a murine critical limb ischemia model. Affected limbs in a murine hind limb ischemia model were treated with intramuscular injected BDACs or the vehicle only. Revascularization was monitored by MRI angiography. (a) Reconstructed MRI images with the region of interest (ROI) indicated by red box. (b) Quantification of vessel volume in ROI. (c) Representative pictures of affected limbs in both treatment groups. (d) Ischemic scoring table. (e) Semiquantitative ischemia scoring accordingly to d. (f,g) H&E staining of the murine gastrocnemicus muscle of the calf in the vehicle- and BDAC-treated group respectively. Arrows indicate neutrophil infiltration, arrowheads indicate fibrosis and stars (*) indicate muscle degeneration with adipose replacement.

Discussion

We describe here a modified protocol to generate blood-derived angiogenic cells (BDACs) with pericyte characteristics in large numbers and in a short period of time. This protocol is based on an earlier method for generating fibrocytes from PBMCs³⁰ and is subjecting crude PBMCs to a 24 hours pulse of MMC using mixed Ficoll species. Ficoll supplementation in media has been approved by regulatory authorities (FDA, 21 CFR §884.6180), for clinical in vitro fertilization applications. Hence, the protocols and materials developed herein can easily be translated into clinical practice. The employed method produces a spindle-shaped cell type, termed BDAC, with a unique marker and proliferation profile that is distinct from fibrocytes, ELC, or macrophages. BDACs exhibited proangiogenic behavior in vitro and in vivo. Interestingly, spindle-shaped cells cultured under noncrowded conditions displayed a different set of markers and did not show the same angiogenic potential in vitro. This indicates that MMC is essential for generating the distinct angiogenic BDAC phenotype. A single pulse of MMC is also responsible for the metabolic switch from citrate cycle to the PPP, resulting in enhanced proliferation. PPP produces ribose, a building block for RNA and DNA synthesis. Therefore an increase in PPP activity is associated with enhanced proliferation. The protocol is robust and has produced consistent results in terms of marker expression and functionality with PBMCs derived from eight different donors in the course of over 4 years. We adopted a crude cell mixture model (as opposed to a CD14-based prepurification) to allow for cross-talk between all components of PBMCs, including platelets. Our assumption that MMC would amplify cytokine signals in such a system was corroborated in a simpler macrophage model. Although increasing IFN-γ concentrations did not increase TNF-α secretion, MMC was able to enhance M1 polarization, suggesting that MMC was amplifying the effects of endogenous cytokines secreted by macrophages in this monoculture system. Future experiments may involve the closer examination of responsible factors and cells for the generation of BDACs.

In contrast to pericytes that are currently assumed to be in essence MSCs,^24,30,31 BDACs show a unique marker profile characterized by pericytic features along the hematopoietic/monocytic lineage, but without mesenchymal stem cell characteristics. Although it is generally accepted that angiogenic cells can be derived from blood, we propose that BDACs represent an alternative pericyte-like cell population of distinct hematopoietic origin, which is involved in early stages of angiogenesis. Although monocyte-derived cells are heterogenous,⁷ we propose that BDACs are related to hematopoietic pericytes, which thus far have only been observed in vivo at early stages of angiogenesis.^{24,25,26,28,36} Thus, our data support the existence of two different pericyte populations, one of hematopoietic and the other of mesenchymal origin, and this would address some of the discrepancies in the literature on the origin of pericytes. However, the question currently remains whether BDAC occur physiologically in humans. The BDAC phenotype occurs reproducibly once MMC is introduced as amplifier of signaling and metabolic tuner in the absence of additional factors, or genetic manipulation. Therefore, the relative ease of BDAC generation would suggest that this phenotype may not be merely an in vitro artefact, but one that could arise physiologically in repairing tissue. Future work will be necessary to determine this phenomenon.

Beyond expression of surface markers, their most compelling aspect of BDACs is their functionality.³⁶ BDACs colocalized with and stabilized endothelial tubular networks on matrigel. They also colocalized with endothelial sprouts in collagen I gels and improved their morphology. In a previous study,³⁶ we have characterized pericyte specific behavior in functional in vitro models, which have been also used in the present study. Furthermore, we have demonstrated that colocalization of cells with the endothelial tubular network on matrigel or with endothelial sprouts in the spheroid sprouting assay is not a pericyte specific behavior. Nonpericytic MSCs, and even fibroblasts show the same behavior. However, the stabilization of the tubular networks on matrigel with improved sprout morphology in the spheroid sprouting assay was a pericyte specific behavior.³⁶ Furthermore, when coinjected with tumor cells, BDACs increased the vascularity of tumors, and localized both in perivascular position (as expected for pericytes), but also in the tumor stroma. A similar distribution was reported for MSC-like pericytes when introduced into infarcted myocardium.⁵⁰ As expected, BDACs were found at sites awaiting vascularization (avascular center) and sites undergoing angiogenesis, and were absent from vascularized parts of the tumor (periphery), indicating that BDAC may take part in different stages of angiogenesis, whereas MSC-like pericytes are known to persist and support formed vessels.²⁴ BDAC location in tumors is in agreement with previous in vivo findings, where hematopoietic pericytes migrated ahead into avascular areas followed by endothelial sprouts at early stages of angiogenesis²⁸ and colocalizing with forming vessels after sprouts have formed.^14,24,25,26

BDACs differ from fibrocytes, EPC and macrophages in terms of marker expression. Yet they appear to promote angiogenesis via classical angiogenic factors such as MMP9, VEGF, and IL-8, which were also attributed to fibrocytes, ELC and tumor associated macrophages (TAM).^{3,15,19,21,23} Thus, it can be concluded that BDAC although exhibiting a unique phenotype have a mode of action similar to other monocyte-derived angiogenic cells. The issue of the clinical utility of BDAC hinges on their efficiency in a clinically relevant model of tissue repair and the number of cells that can be generated from a single blood sample. The ability of BDAC to significantly enhance revascularization in a hind limb ischemia model resulted in the rescue of the lower limb in contrast to control groups, which suffered from extensive necrosis. This efficacious outcome in a preclinical model of critical limb ischemia provides a proof-of-principle for the therapeutic efficacy of BDACs in ischemic diseases.

We estimate that it's possible to generate 40–160 million BDAC within 5 days, from a single blood donation (400 ml). In comparison, clinical trials use crude mixtures or isolated cells from bone marrow derived mononuclear cells or PBMC, after G-CSF mobilization. Cell numbers as low as 1.6 million CD133-selected cells⁴ or 16-2460 (ref. 3) million nonselected mononuclear cells, of which only a very small fraction would be functional, were delivered in clinical trials for the treatment of acute myocardial infarction.

We have demonstrated the capability to generate large numbers of proangiogenic cells with therapeutic efficacy in ischemic diseases, BDAC, from peripheral blood. This opens avenues to generate clinically relevant numbers of patient-derived, functional cells from an easily accessible source with little risk. Therefore BDAC might be well able to overcome current shortcomings in autologous cell therapy⁵ of ischemic diseases.

Materials and Methods

Cell culture. HUVEC were cultured in EGM-2 (Lonza, Basel, Switzerland). Pl-Prc were cultured in PGM (Promocell, Heidelberg, Germany). Human bmMSCs (Lonza) and IMR-90s (ATCC, Manassas, VA) were cultured in DMEM (Gibco, Life Technologies, Carlsbad, CA) (10% FBS, 1% P/S). THP-1 (ATCC) were maintained in RPMI 1640 (10% FBS, 1% P/S). THP1 at a density of 2 × 10⁵ cells/ml were differentiated using 16.2 nmol/l phorbol 12-myristate 13-acetate (PMA, Sigma-Aldrich, St Louis, MO) for 3 days followed by 5 days culture in PMA-free medium. (All incubations were at 37 °C in 5% CO₂.)

Isolation of PBMC. After approval was obtained from the IRB office, volunteers were recruited to donate blood samples of 50–100 ml. PBMC were isolated over a Ficoll gradient (GE Healthcare, Uppsala, Sweden) using manufacturer's protocol.

Generation of BDAC. PBMC were seeded in DMEM with (10% FBS, 1% P/S) supplemented with the Ficoll cocktail (Fc400 25 mg/ml and Fc70 37.5 mg/ml) at 2 million cells/ml on fibronectin (Sigma-Aldrich) coated dishes at 0.25 ml per cm². After one day media was removed and fresh DMEM (10% FBS, 1% P/S) was added. Adherent cells were cultured for further 4 days. Experiments were performed after 5 days of culture.

Generation and polarization of macrophages from PBMCs. Monocytes were isolated from PBMCs via adherence. Cells were cultured for 7 days in RPMI-1640 with 10% FBS, supplemented with M-CSF (100 ng/ml, PeproTech, Rocky Hill, NJ). Macrophages were polarized into M1 with LPS (100 ng/ml, Sigma-Aldrich) and IFN-γ (20 ng/ml, PeproTech) and into M2 with IL-4 (20 ng/ml, PeproTech).

Phagocytosis assay. Fluorescent microspheres (Invitrogen, Life Technologies, Carlsbad, CA, F8823, 1 µl beads/ 300 µl medium per well) were incubated with cells for 1 hour. Cells were harvested and analyzed by flow cytometry (Cyan ADP flow cytometer, Beckman Coulter, Brea, CA).

Adherent cytometry. Cells were stained with DAPI. Adherent fluorescent cytometry was based on a montage of nine sites per well taken by a coolSNAP HQ camera attached to a Nikon TE2000 microscope at 2× magnification, covering 83% of total well area. DAPI fluorescence was accessed with a single DAPI filter [Ex 350 nm/Em 465 nm].

MTS assay and G6PDH activity assay. MTS assay was performed using the CellTiter 96 AQueous One Solution Kit (Promega, Madison, WI) and measurements of G6PDH activity were performed on the same samples using commercial G6PDH Activity Assay Kit (Sigma-Aldrich) according to manufacturer's instruction.

ELISA for TNF-α and IL-10. Supernatants of indicated cell types were analyzed for secreted TNF-α and IL-10 using the DuoSet ELISA kits (R&D Systems, Minneapolis, MN) following the manufacturer's instructions. Results were normalized to cell numbers as quantified by adherent cytometry.

Flow cytometry (FC). Cells suspensions were stained at 2 × 10⁵ cells/sample in PBS containing 1% FBS (FC buffer). Each 50 µl of cell suspension was incubated with respective antibody (Supplementary Table S1) for 1 hour at 4 °C in the dark. Cells were analyzed using the Cyan ADP flow cytometer, Beckman Coulter. All data displayed represents cell population gated in R3. Gates R8 and R13 indicate signal intensity of positive signal.

Immunocytochemistry. Cells were fixed with 100% methanol and blocked for 1 hour in 3% BSA in PBS and incubated with respective primary antibodies for 90 minutes and with secondary antibodies for 30 minutes at room temperature (Supplementary Table S2). Samples incubated without primary antibodies served as conjugate controls (Supplementary Figure S3). Staining was examined using Olympus IX71 inverted microscope (Olympus, Center Valley, PA).

Induction of collagen I secretion and SDS-Page of pepsin digested culture. Collagen secretion was induced with 100 mmol/l L-ascorbic acid phosphate (Wako Pure Chemical Industries, Osaka, Japan). Conditioned culture medium was digested with porcine gastric mucosa pepsin (2,500 U/mg; Roche Diagnostics Asia Pacific, Singapore) in a final concentration of 100 mg/ml. Media were analyzed by SDS- PAGE with 5% acrylamide under nonreducing conditions. 1 mg/µl collagen I standards (Koken, Tokyo, Japan, IAC) served as a molecular weight standard.

RT-PCR. Total RNA was extracted using the RNAeasy kit (Qiagen, Venlo, the Netherlands) following the manufacturer's protocol. cDNA was synthesized from equal amounts of mRNA using Superscript reverse transcriptase II. RT-PCR reactions were performed using following primers: COL1A1 5′ agccagcagatcgagaacat 3′, 5′ cttgtccttggggtcttg 3′; vWF 5′ taagtctgaagtagaggtgg 3′, 5′ agagcagcaggagcactggt 3′; GAPDH 5′ gtccactggcgtcttcacca3′, 5′ gtggcagtgatggcatggac3′. RT-PCR was monitored on a Stratagene real-time PCR instrument (Stratagene, La Jolla, CA) with a PCR master mix based on Platinum Taq DNA polymerase (Invitrogen). Data analysis was performed using the MxPro software (Stratagene). Expression levels of COL1A1 and vWF mRNA levels were normalized to the mRNA levels of GAPDH.

Life cell labeling. Cells were labeled with green and red fluorescence according to manufacturer's instructions (Sigma-Aldrich, PKH67/PKH26).

Tube formation assay on matrigel. 48-well–plate wells were coated with 150 μl of matrigel (Becton and Dickinson, Franklin Lakes, NJ). HUVEC were seeded in 250 μl EGM-2 or DMEM (0.5% FBS) (starving media) at 30,000 cells/well. Where indicated, 15,000 BDAC were added. Pictures were taken by an epifluorescence microscope (Olympus IX71). Cumulative tube length per well was measured by the simple neurite tracer-plugin of Fiji software (http://fiji.sc/Downloads).

Spheroid sprouting assay. 500 HUVEC per well were seeded in EGM-2 containing 2.5 μg/ml methylcellulose into nonadherent round-bottom 96-well plates overnight to form spheroids. When indicated, 250 BDAC/well were added. 24 Spheroids were collected on the next day and resuspended in 1.5 ml of EGM-2 containing 2.5 μg/ml methylcellulose and 1 mg/ml naturalized collagen I in a 12 wp well. Pictures were taken with an epifluorescence microscope (Olympus IX71) after 3 days. Cumulative tube length was measured using Fiji software. Where indicated, MMP9 was inhibited using a MMP9/13 inhibitor (Santa Cruz, Dallas, TX, sc-311438) at a final concentration of 2 μmol/l.

In vivo xenograft tumor assay. All animal procedures were approved by the Institutional Animal Care and Use Committee (AICUC). Five million U87-MG cells were mixed alone or together with 2 million BDAC in 200 μl of L15 media and added to concentrated matrigel at a v/v ratio 1:1. Nude mice were anesthetized using isoflurane. Cell suspensions (200 μl) were injected subcutaneously into the flanks of the mice. After 8 days mice were sacrificed and tumors harvested. Tumors (five control and five supplemented with BDAC) were snap-frozen and cryosectioned at a thickness of 8 μm. Cryosections were fixed in ice-cold 100% methanol or with 4% formaldehyde. Sections were stained (Supplementary Table S3) according to immunocytochemistry protocol and viewed using an epifluorescence microscope (Olympus, IX71) or an Apotome (Zeiss). Quantifications for vWF were performed on one middle section per tumor, whereas quantifications for CD31 were performed on two middle sections per tumor using Fiji software (http://fiji.sc/Downloads). Image stitching was performed at the Nikon centre (SBIC) using Nikon Ni-E microscope.

Angiogenesis proteome array. BDAC were incubated with DMEM containing 0.5% FBS for 3 days. Media was collected and unconditioned media as a control were filtered through a 0.22 μm pore-size filter. Secretion profile of BDAC was established using an angiogenesis proteome array from (R&D Systems) (Cat. no. ARY007) following the manufacturers protocol.

Zymography. Zymograph SDS-PAGE gels composed of 10% acrylamide gel containing 1 mg/ml gelatin or neutralized collagen I and of a stacking gel containing 3% acrylamide. After sample separation by SDS-PAGE zymographs were washed with buffer containing 2.5% TritonX100, 50 mmol/l Tris, 5 mmol/l CaCl₂, and 1 μmol/l ZnCl₂ for 1 hour. Afterwards zymographs were rinsed with deionized water and incubated in buffer lacking 1 μmol/l TritonX100 at 37 °C with gentle agitation over-night. Zymographs were stained with Page Blue using manufacturer's protocol.

Murine hind limb ischemia model. All animal related procedures were in accordance with the Institutional Animal Care and Use Committee at Biological Resource Centre, A*STAR, Singapore. Unilateral hindlimb ischemia was induced in Balb/C nude mice (male, 10–19 weeks, 20–26 g). The mice were anaesthetized by intraperitoneal injection of Ketamine (150 mg/kg) and Xylazine (10 mg/kg). The external iliac artery was gently separated, ligated twice with 7/0 polypropylene suture (Premilene, Braun, Melsungen AG), and dissected between the two ligations. Animals received subcutaneous injection of Enrofloxacin (10 mg/kg, once daily) for 5 days and of Buprenorphine (0.1 mg/kg, twice daily) for 3 days after surgery. One day after surgery, the BDAC group (n = 7) was injected with 10⁶ BDACs suspended in 0.1 ml of Hank's Balanced Salt Solution (HBSS) intramuscularly at three different sites on the operated hindlimb, controls (n = 6) received HBSS only. BDAC from two independent donors were used for this experiment.

Magnetic resonance angiography and data analysis. The vasculature of the animals was accessed using time-of-flight magnetic resonance angiography (MRA) at 7T (ClinScan, Bruker, Ettlingen, Germany) (flip angle = 70, FOV = 25 × 18.75 mm, TR = 20 microseconds, TE = 3.84 microseconds, slice number = 90, slice thickness = 0.3 mm, distance factor = −2.00, average number = 6, acquisition time per animal = 43 minutes).

MRA data was analyzed using ImageJ software. Blood vessels were segmented by thresholding. The total volume of blood vessels was calculated by multiplying the total vessel area by slice thickness times (1 − distance factor/100). 30 slices covering from the start of thigh to lower calf were used for the quantification. The thresholded images were stacked using the VolumeViewer plugin in ImageJ to create three dimensional views of the vasculature.

Statistical analysis. Obtained values were averaged and are displayed as average value ± SD. P values were calculated using Student's t-test.

SUPPLEMENTARY MATERIAL Figure S1. Cell count of adherent cells with data from two other independent blood samples. Figure S2. Flow cytometry analysis of control spindle-shaped cells resulting from PBMCs cultured under noncrowded condition. Figure S3. Conjugate controls (CC) for immunocytochemistry and immunohistochemistry experiments. Figure S4. Immunocytochemical comparison of BDAC and noncrowded controls (−MMC) by the expression of angiogenic and pericytic markers. Figure S5. Immunocytochemical comparison of HUVEC, Pl-Prc and bmMSCs cultured under MMC or noncrowded conditions. Figure S6. Spheroid sprouting assay with data from two other independent blood samples. Figure S7. Spheroid sprouting assay comparing angiogenic potential of BDAC and controls cells cultured under noncrowded conditions. Figure S8. Analysis of tumors resulting from coinjection of BDAC with U87 subcutaneously into nude mice. Figure S9. Quantification of densitometric signal of angio proteome array. Table S1. Antibodies used for flow cytometry. Table S2. Antibodies used for immonocytochemistry. Table S3. Antibodies used for immunohistochemistry. Video S1.