Unacylated Ghrelin Rescues Endothelial Progenitor Cell Function in Individuals With Type 2 Diabetes

Blood was recovered from 14 individuals with type 2 diabetes who arrived in our patient clinic (sex [M/F], 8/6; A1C, 8 ± 1.2%; age, 55.8 ± 9.46 years; BMI, 28.1 ± 3.22 kg/m²; creatinine, 1.06 ± 0.20 mg/dl; waist circumference, 98.8 ± 8.02 cm; total cholesterol, 198 ± 30 mmol/l; HDL cholesterol, 48 ± 12.98 mmol/l; LDL cholesterol, 115 ± 38.94 mmol/l; triglycerides, 159.5 ± 55.23 mg/dl; fasting glucose, 125 ± 18.03 mg/dl; no retinopathy; hypertension in three patients; blood pressure, 142/89 mmHg; cholesterol/apolipoprotein B, 1.3 ± 0.3). All were treated only with diet. No medications were used. A total of 12 blood donors were used as control subjects (sex [M/F], 6/6; age, 47.8 ± 4.85 years; BMI, 21.25 ± 8.07 kg/m²; creatinine, 0.90 ± 0.083 mg/dl; total cholesterol, 164.03 ± 9.68 mmol/l; HDL cholesterol, 50 ± 8.4 mmol/l; LDL cholesterol, 85.26 ± 16.72 mmol/l; triglycerides, 131.14 ± 25.03 mg/dl; no retinopathy; no hypertension; blood pressure, 126/70 mmHg; cholesterol/apolipoprotein B, 1.6 ± 0.2). Ethics approval was obtained from both SIMT (Servizio Immunoematologia e Medicina Trasfusionale) and the Institutional Review Board of S. Giovanni Battista Hospital, Turin, Italy. Informed consent was provided according to the Helsinki Declaration. For the present study, we had no direct contact with human subjects.

Testing sessions were as follows: UAG (3.0 μg · kg⁻¹ · h⁻¹ i.v. as infusion for 12 h, from 0 to 12 h); AG (1.0 μg · kg⁻¹ · h⁻¹ i.v. as infusion for 12 h, from 0 to 12 h); and isotonic saline (infusion from 0 to 12 h).

All tests were performed starting at 8:30–9:00 a.m. after overnight fasting. An indwelling catheter was placed into an forearm vein for slow infusion of isotonic saline. Cells were isolated from blood samples taken at 0, 6, and 12 h.

To isolate CACs, peripheral blood mononuclear cells retrieved from healthy subjects (nCACs) or from individuals with type 2 diabetes (dCACs) were plated on fibronectin-coated dishes as described by Hill et al. (19). Briefly, the cells were cultured for 4 days in EGM-2 medium (Cambrex, Walkersville, MD). To isolate EPCs (nEPCs from healthy subjects, dEPCs from individuals with type 2 diabetes), peripheral blood mononuclear cells were recovered and cultured onto collagen-1–coated dishes for 21 days in EGM-2 medium as described by Yoder et al. (26). In selected experiments, CACs or EPCs recovered from saline-infused healthy subjects were cultured with 1.2 mg/ml AGE, H₂O₂ (100 μmol/l), 5 mmol/l glucose, or 25 mmol/l glucose alone or in combination with 1 μmol/l UAG or 1 μmol/l AG; 1 μmol/l UAG or 1 μmol/l AG was also used alone. Fluorescence-activated cell sorter analysis was used to characterize CAC and EPC surface markers (anti-CD45, anti-CD14, anti-CD34, anti-CD31, anti–Tie-2, anti-KDR, anti-vWF antibodies; see supplementary methods, available in an online appendix at http://diabetes.diabetesjournals.org/cgi/content/full/db09-0858/DC1). eNOS expression was also evaluated.

The plasma glucose and insulin determination for each group of mice (Charles River Lab, Lecco, Italy) are reported: 16 8-week-old ob/ob mice (blood glucose, 296 ± 19.6 mg/dl; insulin, 55 ± 9 ng/ml); 16 8-week-old C57BL/6J wild-type mice (blood glucose, 92 ± 7.2 mg/dl; insulin, 10 ± 0.5 ng/ml). Animal procedures conformed to the Guide for the Care and Use of Laboratory Animals (27). Blood glucose was measured with a One Touch II glucose meter (LifeScan, Mountain View, CA). Serum insulin was measured with a mouse insulin radioimmunoassay kit (Linco Research, St. Charles, MO), following the manufacturer's instructions.

All reagents and antibodies used are reported in the supplementary methods.

Detection of ROS; GTP-Rac1 loading assay; senescence assay; Western blot analysis; silencing of endogenous p53, Akt, and p47^phox by small interfering RNAs (siRNAs); Matrigel plug assay; immunohistochemistry and immunofluorescence; human and mouse mobilization assays; enzyme-linked immunosorbent assay and radioimmunoassay; isolation and culture of bone marrow–derived cells; and evaluation of MMP9 activation, cytofluorimetry analysis, and in vitro migration assays were described in detail in the supplementary methods.

In vitro and in vivo results are representative of at least three independent experiments, performed at least in triplicate. Densitometric analysis using a Bio-Rad GS 250 molecular imager was used to calculate the differences in the fold induction of protein activation or expression. Significance of differences between experimental and control values (*, #, °, and § indicate P < 0.05, statistically significant) was calculated using ANOVA with Newman-Keuls multicomparison test.

Several lines of evidence indicate that the number and function of EPCs are impaired in diabetes (17,19) and that these events rely mainly on Nox-mediated ROS production (15). The effect of in vivo UAG and AG administration in protecting both CACs and EPCs from oxidative damage was first evaluated. Toward this end, cells isolated from UAG- or AG-treated individuals with type 2 diabetes and healthy subjects were characterized for CAC and EPC markers (supplementary Fig. 1) (18) and subjected to dichlorofluorescein diacetate fluorescence assay. The results reported in Fig. 1,A and B demonstrate that UAG, but not AG, treatment (6 h) drastically reduced ROS production in individuals with type 2 diabetes. Moreover, the number of viable cells was significantly higher after UAG treatment compared with before (Fig. 1 C). A protective effect of UAG on ROS production was also demonstrated in cells cultured with AGE or high glucose (supplementary Fig. 2A). Similar results were obtained when H₂O₂ was used (supplementary Fig. 2B). These findings, together with the observation that neither in vitro nor in vivo UAG administration (supplementary Fig. 2C and D) changed AGE receptor (RAGE) expression, suggest that effector(s) downstream of RAGE is the target for protective effect of UAG. Because similar results were obtained after 12 h of treatment (data not shown), data throughout the study relates to 6-h UAG treatment.

Apocynin, known to affect the assembly of Nox subunits (28), was used to investigate the role of Nox in regulating ROS production. Cells recovered from saline-infused individuals with type 2 diabetes were subjected to apocynin treatment. ROS production was prevented in both dCACs and dEPCs (Fig. 1,D). UAG addition could not further enhance apocynin's effects (Fig. 1,D), suggesting that Nox activity might be controlled by UAG. As the assembly of p47^phox and p67^phox subunits is required for Nox enzymatic activity (29), silencing of p47^phox in dCACs and dEPCs (supplementary Fig. 3A) prevented ROS production (Fig. 1,E). Once more, this effect could not be further enhanced by the addition of UAG (Fig. 1,E). Nox2 activity is also dependent on the small GTPase Rac (29). ROS generation in response to AGE (supplementary Fig. 3B and C) was prevented in cells expressing a dominant-negative RacN17 construct. We thus hypothesized that UAG interfered with Rac1 activity. Indeed, Rac1 activation was detected in cells recovered from saline-treated individuals with type 2 diabetes, but not from UAG-infused patients (Fig. 1 F). Similar results were obtained in vitro by culturing cells with AGE (supplementary Fig. 3D). Hence, the modulation of Rac1 activity is a crucial step in the UAG anti-ROS protective effect.

Rac1 membrane localization and function rely on isoprenylation, which has been correlated with AMP-activated protein kinase (AMPK)-dependent hydroxymethylglutaryl CoA reductase activity (30). To rule out the possibility that inhibition of Rac1 activity by UAG depends on this pathway, AMPK phosphorylation was evaluated. Neither short- nor long-term exposure to UAG alone or in combination with AGE affected AMPK activity (supplementary Fig. 4).

p53, p21, and pRb are major regulators of cell senescence (31). The above results prompted us to evaluate whether the increase in ROS production, generally considered as an upstream signal, translates into an accelerated onset of senescence and whether UAG could rescue this effect. Because both senescence and ROS generation (Fig. 2,A and B) were prevented by silencing p53 (supplementary Fig. 5A), we investigated the in vivo effect of UAG on p53 expression. Accordingly, UAG treatment was able to prevent p53 accumulation, p21 expression, and Rb phosphorylation (Fig. 2,C) and to reduce the number of senescence-associated β-galactosidase (SA-β-gal)-positive dCACs and dEPCs (Fig. 2 D) in UAG-challenged patients. Similar results were obtained in AGE-treated cells (supplementary Fig. 5B and C).

To assess whether the protective effect of UAG also resulted in an enhancement of dCAC and dEPC vasculogenic capability, de novo vessel formation was analyzed in severe combined immunodeficient mice injected with cells recovered from UAG-treated individuals with type 2 diabetes. At 15 days after injection, plugs were recovered and analyzed by immunohistochemistry. As shown in Fig. 3 A and B, the number of functional vessels formed by cells recovered from UAG-treated patients was significantly increased with respect to those from saline-treated patients. The origin of neovessels from host vasculogenic cells was excluded because the majority of vessels were lined by human HLA class I–positive cells (supplementary Fig. 6) (32). Thus, these data provide evidence that UAG restores dCAC and dEPC vasculogenic activity.

Defective EPC and CAC mobilization has been reported in diabetes (19,20). To further investigate the potential therapeutic effect of UAG, CACs and EPCs were recovered from 10 normal healthy subjects and 10 individuals with type 2 diabetes, characterized, and counted. UAG treatment led to an increase in the number of recovered cells in individuals with type 2 diabetes compared with that of healthy subjects, and no effect of UAG treatment was detected in healthy subjects (Fig. 4,A). In contrast, no differences between AG- or saline-treated healthy and type 2 diabetic individuals were observed (Fig. 4,A). In addition, as shown in Fig. 4 B, the percentage of circulating senescent dCACs and dEPCs decreased upon UAG treatment.

Circulating stromal derived factor-1 (SDF-1) (33) and vascular endothelial growth factor (VEGF) (34) strictly control progenitor cell mobilization under stress conditions. We herein demonstrate that 6- or 12-h (data not shown) UAG or AG infusion did not change their serum concentrations (supplementary Fig. 7). Similarly, IGF-1 (35) serum concentration was not affected by UAG treatment (supplementary Fig. 7).

For validation and characterization of the molecular mechanisms regulating bone marrow mobilization, a mouse model of type 2 diabetes (ob/ob mice) was used. After treatment with saline, UAG, or AG, recovered cells were subjected to fluorescence-activated cell sorter analysis for surface markers to confirm EPC identity (data not shown). UAG treatment induced a strong increase of recovered EPCs only in ob/ob mice (Fig. 4,C). Finally, the number of senescent cells was significantly lower in UAG-treated ob/ob mice compared with untreated or AG-treated animals (Fig. 4 D), reproducing our findings in human subjects.

As an impairment of eNOS phosphorylation contributes to defective EPC mobilization in the diabetic setting (36), we investigated whether UAG modulated eNOS activity and the activation of its regulatory protein, Akt (37), in bone marrow stromal cells. The stromal origin of the eNOS-expressing cells was confirmed by the presence of mbKitL (Fig. 5,A). In ob/ob mice, UAG treatment restored both eNOS and Akt phosphorylation (Fig. 5,A). Consistent with the pivotal role of a local activation of MMP9 in promoting progenitor cell mobilization (22), gelatin zymography revealed that MMP9 gelatinolytic activity was induced by UAG (Fig. 5,B). The role of MMP9 activation and sKitL release in controlling this event was further confirmed by functional studies using anti-MMP9 and anti-KitL neutralizing antibodies (Fig. 5,C). Accordingly, in parallel with effects on MMP9 activation, expression of the mbKitL was decreased in ob/ob mice subjected to UAG treatment (Fig. 5,A). Although we cannot rule out the possibility that a paracrine effect of UAG occurs in vivo, herein we have shown that in vitro UAG treatment for 40 min elicited Akt and eNOS phosphorylation in stromal cells obtained from ob/ob-derived total bone marrow pools (Fig. 5,D). In agreement with the results measuring EPC mobilization, AG failed to induce Akt and eNOS phosphorylation (Fig. 5,D). The finding that UAG failed to induce eNOS phosphorylation after knocking down Akt (Fig. 5,E) or inhibiting its activation (Akt inhibitor) (Fig. 5 D) indicates that Akt is a key modulator of the UAG effect. The above data were validated by the lack of effect of UAG treatment in NOS3^−/− mice (23) (supplementary Fig. 8).

Based on our collective dataset, we further investigated whether the biological response of CACs to UAG was mediated by specific binding sites localized to the plasma membrane. To this end, double immunofluorescence experiments, using the UAG analog 488-UAG (100 nmol/l) to label putative binding sites and anti–phycoerythrin-CD45 antibody as a membrane marker, were carried out at 4°C. As shown in Fig. 6,A, at 4°C 488-UAG–binding sites colocalized with CD45, indicating a plasma membrane localization. In agreement with the functional data, unlabeled UAG (1 μmol/l) displaced the fluorescent signal from the cell surface, whereas AG (1 μmol/l) did not (Fig. 6,A). In addition, we also monitored receptor activation at 25°C. As shown in Fig. 6,B, after 20-min stimulation, receptor clusters undergoing internalization were visualized as labeled cytoplasmic vesicles. Moreover, increasing concentrations of unlabeled UAG specifically displaced the fluorescent ligand from both the plasma membrane and endocytic vesicles (Fig. 6 C and supplementary Fig. 9). Similar results were obtained using EPCs (data not shown).

The present data first demonstrate that UAG, unlike AG, reverts diabetes-induced EPC damage by inhibiting activation of the Nox regulatory protein Rac1; as a consequence, UAG protects diabetic EPCs from senescence and improves their vasculogenic capability; again, only UAG rescues EPC mobilization under diabetic conditions by restoring eNOS phosphorylation, and specific UAG-binding sites mediate its effects.

UAG is the most abundant circulating form of ghrelin (3) and plays a positive role on glucose metabolism. In contrast, basic and clinical studies have proposed AG as a diabetogenic hormone (8,38). Indeed, clinical conditions of insulin resistance are associated with an alteration in the circulating ghrelin profile with relative AG excess with respect to UAG (12). Thus, it is tempting to speculate that an altered UAG/AG ratio might contribute both to metabolic changes and to diabetes-associated complications.

Among diabetes-associated complications, abnormal vascular remodeling is believed to play a major role in accelerating vascular disease (39). Historically, it has been assumed that new blood vessels originate from sprouting cells and co-opting of neighboring preexisting vessels. However, both physiological and pathologic angiogenesis are also supported by mobilization and recruitment of other cell types, including the bone marrow–derived cells, such as EPCs (16,18,40,41). Interestingly, alterations in the number and function of these cells correlate with the risk factor profile (19,20). Overproduction of ROSs in these pathologic settings seems to contribute to impaired vascular regenerative processes (42). The plasma membrane Nox is recognized as one of the major regulators of ROS generation (15,29). Activation of the enzyme can occur via many upstream signaling pathways converging on phosphorylation of p47^phox and activation of Rac1 leading to the oxidase assembly (29). Our study shows that UAG can protect diabetic EPCs from oxidative stress by affecting the Nox regulatory protein Rac1.

The accelerated onset of senescence contributes to the impaired EPC bioavailability in patients with diabetes (17). The tumor suppressor p53 is a transcription factor involved in DNA damage mechanisms and is recognized as a negative regulator of cell proliferation in human atherosclerotic and restenosis lesions (43). Moreover, the p53-mediated pathway contributes to EPC senescence-like growth arrest in diabetes (44). Accordingly, the present study shows that silencing p53 in EPCs isolated from individuals with type 2 diabetes or cultured with AGE prevents both ROS production and senescence. The finding that both in vitro and in vivo exposure to UAG negatively modulates p53 accumulation identifies the p53-mediated signal as the primary mechanism through which UAG protects EPCs from senescence. We also found that, by preventing oxidative stress, UAG improves de novo vessel formation. The efficacy of cell therapy certainly depends on the number, functional capability, and successful retention of cells in the site of action. Thus, our data strongly suggest that naturally occurring UAG induces improvement of EPC function and survival that may translate into a more efficient response to vascular dysfunction.

Senescence is also associated with impaired mobilization of bone marrow–derived cells (17). The molecular interactions between stem cells and bone marrow stromal cells, and the molecular mechanisms controlling their mobilization in the bone marrow microenvironment, are poorly understood (45). Considerable interest has arisen about agents able to mobilize and augment progenitor cell delivery to sites of vascular injury to enhance revascularization. Among these, SDF-1 and VEGF are recognized as primary regulators of bone marrow cell mobilization during stress conditions (22,46). In addition, under physiological stresses, the activation of matrix proteases within the bone marrow microenvironment results in the release of sKitL, which enables endothelial and hematopoietic progenitor cells to transit from the quiescent to the proliferative niche and facilitates their mobilization into the circulation (22). The delivery of progenitor cells to sites of neovascularization also relies on functional eNOS activity (23). Indeed, in pathologic settings associated with blunted eNOS activity and reduced systemic NO bioavailability, defective EPC mobilization and impaired vascular regenerative processes occur (24,25). eNOS activation through Akt has been reported for AG acting on the Gq-coupled GHS-R1a in cultured endothelial cells (47). We herein demonstrate that UAG can restore eNOS activity via Akt-mediated phosphorylation in a pathologic setting characterized by impaired eNOS phosphorylation. This was particularly true in bone marrow stromal cells from diabetic mice. Such an event was found crucial to EPC bone marrow mobilization, because UAG had no effect in eNOS knockout mice. Gu et al. (48) identified MMP9 as a major target of NO. We also showed that eNOS phosphorylation, occurring in response to UAG, is associated with MMP9 activation and possibly with the release of sKitL, as suggested by the reduced expression of the mbKitL on stromal cells recovered from UAG-treated mice and by functional studies. Although we cannot exclude that UAG may act in a paracrine manner by locally inducing the release of VEGF from bone marrow stromal cells, we demonstrate that short-term treatment with UAG, but not AG, of bone marrow cells in vitro leads to Akt and eNOS phosphorylation and that, in vivo, these events are associated with MMP9 activation and EPC mobilization. Furthermore, UAG, unlike AG, strongly induced EPC mobilization in individuals with type 2 diabetes, but not in nondiabetic subjects. Notably, no change in serum concentrations of primary mobilization factors was detected after UAG or AG systemic administration.

In addition, our results shed light on the earliest molecular events leading to UAG- but not AG-mediated physiological regulation of EPC bioavailability. Indeed, we showed that EPCs possess specific UAG-binding sites, which are not recognized by AG. In addition to GHS-R1a, which is the AG-specific receptor (3), other ghrelin receptor subtypes exist, whose molecular identities have not yet been characterized, but that recognize both AG and UAG (3). Thus, our data provide the first evidence of the existence of UAG-specific binding sites and of their relevance in human-derived EPCs, which could represent a novel target for pharmacological modulation.

Preclinical and clinical studies generally support the therapeutic potential of autologous EPCs in the treatment of cardiovascular diseases, such as tissue ischemia and myocardial infarction (49). However, EPC mobilization may also accelerate atherosclerotic plaque progression (50) and induce tumor (51) or retina (52) neovascularization in individuals with type 2 diabetes. Nonetheless, therapies with statins, the main EPC mobilization mediators (53), revealed no concerns in terms of neovascularization. We now have reason to believe that the significant EPC mobilization potential of UAG, and the lack of its effects on serum levels of SDF-1 and VEGF, may be exploited for clinical treatment of diabetes- and atherosclerosis-induced vascular impairment.

In the presence of cardiovascular risk factors such as diabetes, EPC availability is reduced, restricting the possibility of treating patients in need with directed/cell-based therapies. Besides displaying a positive influence on β-cell viability and glucose homeostasis (3), UAG mobilizes EPCs, protects EPCs from oxidative stress and from senescence, and increases de novo vessel formation (see model in Fig. 7). This suggests that UAG-related peptides or UAG receptor–specific agonists may be further developed into lead compounds from the perspective of a novel pharmacologic intervention to ameliorate both metabolic control and impaired vascular growth in individuals with type 2 diabetes where AG has failed.

M.F.B. was supported by grants from the Italian Association for Cancer Research (AIRC) and Ricerca Finalizzata Regione Piemonte. M.F.B., L.P., and E.G. were supported by MIUR (Ministero dell'Università e Ricerca Scientifica, cofinanziamento MURST and fondi ex-60%).

No potential conflicts of interest relevant to this article were reported.

We thank Dr. Silvia Soddu for kindly providing the vector pSUPER retro containing p53 siRNA and Dr. Silvia Giordano for kindly providing the V12N17Rac construct as well as for fruitful discussion. We thank Dr. Terence Edgar Hébert for his invaluable contribution to final editing of the article.