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a Departments of Anatomy and Cell Biology,
b Exercise Science, and
c Dermatology, University of Iowa, Iowa City, Iowa, USA
Key Words. Angiogenesis • Endothelial cell • Diabetes • Monocyte • Progenitor cells • Somatic stem cells • Vascular development • Bone marrow cells
Correspondence: Gina Schatteman, Ph.D., Exercise Science 412 FH, University of Iowa, Iowa City, Iowa 52242, USA. Telephone: 319-335-9486; Fax: 319-335-6966; e-mail: gina-schatteman{at}uiowa.edu
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ABSTRACT |
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INTRODUCTION |
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Studying putative EC progenitor dysfunction in diabetes is not simple, because the antigenic phenotype of bone marrow–derived cells capable of differentiating into ECs remains poorly defined, probably because it is plastic [14]. Nevertheless, studies on various subsets of blood and BMCs have provided ample evidence that there are at least two distinct classes of bone marrow–derived EC progenitors, primitive progenitors and monocytic-like progenitors. Primitive EC progenitors (prECPs) were the first identified, initially by expression of the hematopoietic stem cell antigens, CD34 and flk-1 [6], and subsequently by these and other hematopoietic stem cell antigens, notably CD133 (AC133) [15–17]. Later, several studies demonstrated that monocytes or monocyte-like cells can also function as EC progenitors, and it is these monocytic-like cells that are most commonly referred to as EPCs [18–21]. Further studies have shown that these two types of EC progenitors have distinct in vitro and in vivo properties [19, 22].
Both of these EC progenitor classes have been studied in the context of diabetes, but no distinction has been made between these two populations. Human CD34+ peripheral blood mononuclear cells are enriched for prECPs [6]. In culture, blood-derived CD34+ cells from type 1 diabetic but not type 2 diabetic subjects produced fewer ECs than those from nondiabetic controls [13]. Fewer ECs also were produced in cultures of adherent peripheral blood mononuclear cells, that is monocytic ECPs (mECPs), from type 1 and type 2 diabetic blood than nondiabetic blood [23, 24]. In addition, ECs derived from human type 2 diabetic mECPs exhibited reduced integration into vascular tubes in vitro [23].
In vivo, human nondiabetic blood-derived CD34+ cells promoted revascularization of skin wounds in type 1 diabetic mice [25]. In a nude mouse model of hind limb ischemia, exogenous nondiabetic blood-derived CD34+ cells had no effect on the restoration of blood flow to an ischemic limb in nondiabetic mice, but the same cells profoundly accelerated blood flow restoration in type 1 diabetic mice. Similarly, mouse BMCs enriched for murine hematopoietic stem cells dramatically improved vascularization of skin wounds in obese type 2 diabetic Leprdb but not congenic lean nondiabetic C57Bl/6 mice [26]. Moreover, when skin wounds of Leprdb mice were treated with Leprdb-derived hematopoietic stem cell–enriched BMCs, wound vascularization was severely inhibited [26]. In contrast, administration of whole BMCs from both nondiabetic and type 1 diabetic mice improved blood flow restoration in ischemic hind limbs of both nondiabetic and type 1 diabetic mice. However, the effect was greater in mice treated with nondiabetic than diabetic cells [27].
These studies indicate that the ability of bone marrow–derived cells to promote neovascularization as well as to differentiate into ECs may be impaired by diabetes. In type 1 diabetes, both functions may be compromised [13, 24], but the picture is less clear in type 2 diabetes, in which the data are conflicting and less complete. One explanation for some of the conflicting data is that the diabetes affects prECP and mECPs (i.e., adherent whole bone marrow after 4 days in culture) differently, but this has never been examined. Also, although the behavior of BMCs in diabetic and nondiabetic environments differs [13, 26], whether there is a negative synergism between the diabetic environment and diabetic BMCs has not been explored.
To study these issues, we developed a culture system for growth and differentiation of murine EC progenitors and investigated various functional properties of murine hematopoietic stem cells (i.e., mouse prECPs) and adherent BMCs (i.e., myeloid/monocytic EC progenitors) from Leprdb mice. We also compared the ability of nondiabetic and Leprdb prECPs to promote vascular growth in vivo in nondiabetic mice. Our data demonstrate that the obese type 2 diabetic syndrome induces intrinsic defects in prECPs but possibly not in mECPs. The defects in prECPs were evident in vitro by decreases in prECP-derived EC numbers after stress and in vivo in nondiabetic mice by their inhibition of vascular growth in skin wounds and exacerbation of ischemia-induced tissue damage in limb muscle.
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MATERIALS AND METHODS |
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Isolation and Culture of Mouse BMCs
C57Bl/6 or Leprdb male mice (8–10 weeks) were injected intraperitoneally with 150 mg/kg pentobarbital sodium. BMCs were collected from femurs and tibias and enriched for hematopoietic stem cell using Sca-1+ (Ly-6A/E+) magnetic bead–positive selection (Miltenyi, Auburn, CA) for in vitro studies and SpinSep lineage depletion (StemCell Technologies, Vancouver, BC, Canada) so that no beads were injected for in vivo studies, according to the manufacturer’s instructions. Hematopoietic stem cell–enriched cells will be referred to as prECPs hereafter. Enriched cells were R-PE rat anti-mouse Sca-1 (BD Pharmingen, San Diego) labeled and analyzed by fluorescence-activated cell sorting as described to assess purity [26]. Whole bone marrow was harvested from additional mice and preplated on tissue culture plastic and then placed in a tissue culture incubator for 1 hour to remove endothelial and stromal cell contaminants.
Freshly isolated diabetic and nondiabetic-derived prE-CPs and preplated BMCs from Leprdb and C57Bl/6 mice were plated in Medium D [19] with heat-inactivated fetal calf serum reduced from 20% to 7.5%. Cells were plated on 5 µg/cm2 pronectin-coated plastic (Deepwater, Woodward, OK) in 96-well trays at 1.2 x 105 cells per well for 3(4,5-dimethylthiazol-2-y1)-2,5-diphenyltetrazolium bromide (MTT) assays or in eight-well plastic chamber slides at 1.5 x 105 cells per well for immunolabeling at approximately 30% confluence. Some plates were placed in a hypoxia chamber (5% O2). Additional wells were stressed with 200 µM H2O2 at the time of plating or 1 and 4 days after plating. Medium was replaced at 4 days, at which time nonadherent cells were removed. The residual adherent mouse mECPs in the whole BMC cultures are analogous to human EPCs.
At 2 or 8 days, cell medium and any unattached cells were removed and relative cell numbers were determined by MTT assay according to the manufacturer’s instructions (ATCC, Manassas, VA). Briefly, cells were incubated with MTT reagent for 2 hours at room temperature and then in lysis reagent for 2 hours, and optical densities at 540 nm were measured. Assays were performed in duplicate or triplicate four to seven times for mECPs and in single wells or duplicate four to six times for prECPs. Cells in eight-well chamber slides were stimulated and stressed identically to cells in 96-well trays. At 8 days, cells were fixed with methanol and then immunolabeled with 2.5 µg/ml antivascular EC-cadherin (anti-VE-cadherin) (Cayman Chemical Co., Ann Arbor, MI) or isotype-matched immunoglobulin G (IgG) for 2 hours. Label was visualized by 1-hour incubation with 10 µg/ml Alexa 488 anti-rabbit IgG (Molecular Probes, Eugene, OR). Aortic smooth muscle cells served as negative and human umbilical vein ECs (HUVECs) as positive control cells.
To verify the EC phenotype of cultured cells, cells were fixed in methanol after 4 or 8 days in culture and labeled with Bandeiraea simplicifolia isolectin B4 (BSLB4) or immunolabeled with anti-tie 2 or anti-von Willebrand factor (vWF) as described [19] or with anti-VE-cadherin as above. Isotype-matched IgG and cell controls were performed as above.
Wounding
Male 8- to 10-week C57Bl/6J nondiabetic mice were anesthetized with isoflurane, and their back skin was depilated with Nair (Church & Dwight Co., Inc., Princeton, NJ). One day later, mice were again anesthetized, and two 6-mm bilateral full-thickness skin wounds were created on the dorsorostral back skin as described [26]. Three days later, mice were again anesthetized, and 2.5 x 105 freshly isolated prECPs in 25 µl 0.9% NaCl from either C57Bl/6 mice or type 2 diabetic male Leprdb (B6.Cg-m+/+Leprdb) mice [28] were injected under each wound. Controls received 25 µl 0.9 % NaCl. Forallmice, both wounds were injected with the same substance to avoid the possibility that injected cells could migrate or secrete substances into the contralateral wound.
Histology and Morphometry
Thirteen days after wounding, mice were anesthetized and depilated. The next day, they were lethally injected with sodium pentobarbital as above. Wound beds and underlying muscle surrounded by a margin of normal skin were harvested, fixed, and paraffin embedded as described [26]. Eight wounds (two from four different mice) in each group were serially sectioned (7 µm) perpendicular to the wound surface. Every 10th section throughout the entire wound bed was hematoxylin and eosin stained, and the adjacent section was immunolabeled with anti-CD31 (BD Pharmingen) to visualize blood vessels. Sections were treated for 3 minutes at 37°C with 100 µg/ml proteinase K (BD Pharmingen) before 1-hour incubation with 2.5 µg/ml anti-CD31 or rat IgG as control at 37°C in 0.75 µg/ml biotinylated anti-rat IgG and then 1:200 alkaline phosphatase-streptavidin complex (Vector, Burlingame, CA) followed by visualization with Vector Red (Vector) and hematoxylin counterstaining. The number of sections analyzed ranged from 15 to 20 per wound depending on the size of the wound.
Wound morphometry was performed as previously described [26]. Briefly, wound area was measured by digitally tracing the wound periphery (epidermis and dermis) of hematoxylin and eosin–stained sections. Wound volume was estimated by interpolation from the wound areas measured in every 10th section (i.e., every 70 µm) throughout the entire wound as described [25]. Similarly, the area of anti-CD31 immunolabeled blood vessels in the wound was measured digitally, and vascular volume was estimated by interpolation. The vessel volume density (vessel volume/wound volume) then was computed. Additionally, the vessel density (number of vessels per wound area) was computed. Finally, vessel size (vessel area per vessel number) was computed. Data were compared among groups using a one-way analysis of variance (ANOVA) with Tukey’s honestly significant difference post-hoc analysis, and p < .05 was considered statistically significant [29].
Ischemic Limb
Left hind limbs of mice were depilated as above. One day later, surgical induction of unilateral hind limb ischemia was performed on 21 C57Bl/6 mice as described previously [13], and mice were divided into three groups. Two to 5 hours after surgery, mice were anesthetized with isoflurane and the medial thigh of the ischemic limb was injected intramuscularly with 5 x 105 freshly isolated prECPs from Leprdb (n = 6) or C57Bl/6 (n = 6) mice or vehicle (n = 9).
Blood Flow Analysis
Scanning LASER Doppler blood flow imaging (Moor Instruments Inc., Wilmington, DE) was used to assess blood flow restoration in mice after surgery as previously described [13, 30]. Blood flow was analyzed immediately before and after surgery and at 2, 4, 6, 8, and 11 days after surgery. Only mice whose mean flux in the operated limb immediately after surgery was 
12% of that of the unoperated control limb were analyzed. Statistical comparisons of blood flow over time among groups were done by a repeated-measures ANOVA followed by Tukey’s honestly significant difference post-hoc analysis using SigmaStat software (SPSS Science, Chicago). p .05 was considered statistically significant. Data are presented as percent mean blood flux in the operated ischemic limb relative to flux in the unoperated control limb.
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RESULTS |
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).
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. vWF expression was not detected in 4-day but was observed in 8-day cultures (Figs. 1G, 1H
). BSLB4 binding and tie-2 immunolabeling were present at low levels in 4-day cultures. Both persisted at 8 days (Fig. 1E), but the levels did not seem to increase over this time period as tie-2 and Ulex lectin binding do in human cell cultures [18, 19]. (Human ECs do not bind BSLB4 but do bind Ulex lectin.) VE-cadherin expression also differed from that observed for human BMC-derived ECs. Whereas VE-cadherin was expressed only after relatively long-term culture of human BMCs [18, 19], 90% or more of cells in every 4-day culture examined (mean, 96%) labeled brightly with anti-VE-cadherin. This labeling persisted in 8-day cultures (Figs. 1B, 1C). No labeling was present in isotype-matched controls (Figs. 1D, 1F, 1H) or cultures of mouse hepatoma cells, whereas robust VE-cadherin and vWF expression was observed on HUVEC-positive controls (data not shown). VE-cadherin immunolabeling and BSLB4 lectin binding of prECP-derived cells also was performed at 4 and 8 days in culture, and both labels were detected at both times. Tie-2 immunolabeling was performed at 8 days, by which time prECPs were weakly labeled (data not shown).
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). Cell numbers were then assessed 8 days after plating to assay cell growth (proliferation minus cell death).Cellnumbers did not differ significantly between nondiabetic and diabetic mECPs or prECPs at this time point (Fig. 2B). Because in humans it has been reported that diabetes leads to reduced proliferation of mECPs in culture at 7 days, this result was surprising. We considered the possibility that mouse mECPs survived in the diabetic mouse cultures but did not differentiate. Cells in 8-day cultures of mECPs were immunolabeled with VE-cadherin antibodies to identify ECs. Diabetes did not significantly affect the percentage of VE-cadherin–expressing cells; 95.8% ± 0.4% of nondiabetic and 97.2% ± 1.1% of diabetic cells were VE-cadherin positive.
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). Consistent with what we previously observed for nondiabetic human mECPs [22], culture in hypoxia also had no significant effect on nondiabetic and diabetic mouse mECP number in 8-day cultures (Fig. 3B).
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H2O2 treatment reduced cell numbers in both Leprdb mECP (p < .05) and C57Bl/6 mECP (p < .01) cultures by 2 days, although the reduction in cell numbers was similar in the two groups (Fig. 3C). The number of cells remained lower in H2O2-treated cultures relative to controls at day 8, but the reduction in cell number was similar to that observed at day 2 (Figs. 3C, 3D). Because in humans mECPs have not begun to proliferate by 2 days in culture and the cells in the 2-day cultures were assayed only 12 hours after the addition of H2O2 in these experiments, the data suggest that H2O2 induces cell death but does not affect subsequent proliferation. No significant effect of H2O2 treatment on differentiation was apparent. The percentage of VE-cadherin–expressing cells was 95.8% ± 0.4% and 97.6% ± 1.2% in nondiabetic controls and H2O2-treated mECPs, respectively, and 97.2% ± 1.1% and 98.3% ± 1.0% in diabetic controls and H2O2-treated mECPs, respectively.
Because increased oxidative stress and hypoxia occur concomitantly in ischemic tissue, we also assessed the effect of the combination of the two on diabetic mECPs. The additional stress of hypoxia led to no further reduction in cell number in the dual treatment cultures compared with H2O2 alone (Fig. 3E).
Diabetic prECPs Under Stress
Because prECPs and mECPs have distinct properties, we also studied the effects of hypoxia and oxidative stress on prECPs. As with mECPs, hypoxia had no significant effect on prECP plating efficiency (Fig. 4A). In 8-day cultures, however, hypoxia stimulated nondiabetic prECPs (p < .01). Cell numbers in nondiabetic cell cultures increased by almost 50%, whereas hypoxia failed to significantly stimulate diabetic prECPs (Fig. 4B). Also, as with mECPs, H2O2 treatment resulted in decreased cell numbers in both Leprdb (p < .01) and C57Bl/6 (p < .05) prECP cultures (Fig. 4C). The reduction in cell numbers, however, was significantly greater in H2O2-treated diabetic than nondiabetic cultures, being reduced by one third relative to H2O2-treated nondiabetic controls (p < .01) (Fig. 4C).
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Full-thickness skin wounds were created in nondiabetic C57Bl/6 mice. Cells or vehicle was injected under the wounds 3 days after wounding, and wounds were harvested 11 days later (14 days after wounding). In histological sections, marked differences between the three groups were apparent, with an increase in vascularization noted in nondiabetic cell treated and a decrease in Leprdb-cell treated wounds relative to controls (Figs. 5A–C). To quantitate these findings, the vascular volume density (vessel volume/wound volume) and vessel density (vessels per unit area) for cell- and vehicle-treated wounds were determined. In wounds treated with nondiabetic prECP, vascular volume density increased significantly (p < .05) (Fig. 5D), but vascular density was not significantly affected (Fig. 5E) relative to vehicle controls. Consistent with this, mean vascular size was increased in wounds treated with nondiabetic prECPs (p < .01) (Fig. 5F). In contrast, wounds treated with prECPs from Leprdb mice showed a dramatic decrease in both vascular volume density and vessel density (p < .01) (Figs. 5D, 5E) relative to controls, but the mean vessel size was not changed (Fig. 5F).
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). Flow was improved by nondiabetic prECPs relative to limbs treated with Leprdb prECPs throughout the entire 11-day time period (p < .05). No significant effect of injection of Leprdb prECPs on flow restoration relative to vehicle-treated controls was observed, although there was a trend toward an inhibition of flow (Fig. 6). Moreover, injection of Leprdb-derived cells resulted in severe limb necrosis in five of six mice, whereas limb necrosis in control mice was much less severe, with only one mouse exhibiting severe necrosis.
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DISCUSSION |
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We have yet to fully understand the nonhematopoietic functions of BMCs, but it is clear that there are at least two types that in some way promote neovascularization. prECPs, such as some Sca1+lin– BMCs, can function as hemangioblasts [33] and also promote vascular growth, possibly by secreting proangiogenic factors. Furthermore, because prECPs represent a large source of hematopoietic stem cells [34, 35], many if not all monocytic EC progenitors (i.e., mECPs) are likely derived from them. The less well-defined monocytic progenitors also differentiate into ECs in vivo and promote vascularization, also probably by secreting proangiogenic molecules [11, 19]. Interestingly, prECPs seem to modulate mECP function in vivo [19]. Thus, we considered it important to understand potential diabetes-induced changes in both primitive and monocytic EC progenitors.
Consistent with our findings for circulating CD34+ EC progenitors in humans [13, 19], type 2 diabetes does not seem to alter the ability of mouse prECPs or mECPs to produce EC under basal conditions in culture. However, because diabetes is associated with elevated levels of intracellular oxidative stress, the constitutive exposure to this stress might affect the ability of diabetic BMCs to withstand additional oxidative stress such as would be expected to increase during tissue injury. Our data indicate that H2O2-induced increases in oxidative stress leads to cell loss in both nondiabetic and Leprdb mECPs, but the effects are similar in the two groups. In contrast, Leprdb prECPs are more sensitive to H2O2-induced cell death than their non-diabetic counterparts. It remains to be determined why sensitivity of primitive but not monocytic EC progenitors increases. Perhaps they are equally sensitive but uptake or quenching of oxidants by other cells in the mECP cultures limits damage to the monocytic progenitors. Also, it is not known if sensitivity of the prECPs represents an increase in production or uptake of reactive oxygen species by the cells or decreased antioxidant enzyme production or activity, although preliminary data suggest the latter (G. Schatteman, unpublished data).
BMCs are active in tissue repair at times of tissue ischemia. Thus, if they are to differentiate into ECs, they need to be able to do so effectively in the context of hypoxia. We previously found that the ability of nondiabetic human mECPs to produce ECs in culture is unaffected by hypoxia [22]. However, the response of diabetic cells to hypoxia has not been examined. Our data show that as with nondiabetic human cells, hypoxia had no effect on type 2 diabetic or monocytic EC progenitors. In contrast, hypoxia greatly stimulated nondiabetic prECP growth, but Leprdb prECPs failed to respond to the stimulus.
Taken together, our in vitro data suggest that under basal conditions, EC progenitors can withstand the stress of obesity and diabetes. However, when coupled with additional stress, such as elevated oxidative stress, EC progenitor function is compromised. It seems that this sensitivity is confined to primitive cells, because even two additional stresses did not affect mECP function in obese diabetic mouse cells in our assays. Still, in vivo, when the mECPs would be subjected to multiple stresses simultaneously, the picture might be different. Furthermore, after longstanding hyperlipidemia and diabetes, mECP sensitivity might be increased. If the inability of diabetic BMCs to produce ECs in vitro reflects their in vivo functional abilities, a reduction in the integration of BMCs into the vascular endothelium could contribute to diabetes-associated impaired angiogenesis.
BMCs can promote vascular growth not only by integrating into the endothelium but also by secreting factors or in some way interacting with endogenous cells to create a more proangiogenic environment. Thus, diabetes has the potential to modulate this function of BMCs as well. Our earlier data demonstrated that, unlike their nondiabetic counterparts, prECPs from Leprdb mice can inhibit neovascularization in diabetic mouse wounds. This suggested intrinsic dysfunction in the Leprdb-derived cells but left open the question of whether an interaction between an unhealthy environment and dysfunctional cells was required for the inhibition. It was also unclear as to whether this inhibitory function was unique to skin wound healing or was true in other models of neovascularization.
Our finding that Leprdb prECPs inhibit revascularization of nondiabetic skin wounds clearly demonstrates that intrinsic changes in the obese diabetic mouse–derived prECPs are responsible for this inhibition. Moreover, the data suggest that these prECPs secrete factors that change the local milieu from proangiogenic to antiangiogenic. Because the number of vessels rather than the size of vessels is decreased, it is likely that it is the initial phase of vessel growth that is inhibited. Data from prECPs injected into ischemic limbs are consistent with, albeit less dramatic than, findings in skin wounds. Although there was no clear inhibition of neovascularization, there was a trend toward reduced flow in limbs treated with diabetic cells. We suspect that the same inhibitory effect is likely present in the ischemic limb but that the assay is less sensitive, perhaps because of volume dilution of the cells (i.e., the limb is much bigger than the skin wound). A more sensitive indicator of flow restoration may be limb functional improvement, and here we saw a marked deterioration in the limbs treated with Leprdb-derived cells relative to controls, because toe and foot autoamputation and limb necrosis were markedly increased.
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CONCLUSIONS |
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ACKNOWLEDGMENTS |
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REFERENCES |
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