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CORRESPONDENCE Stefanie Dimmeler: dimmeler{at}em.uni-frankfurt.de
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L. Rössig, C. Urbich, and T. Brühl contributed equally to this work.
Endothelial progenitor cells (EPCs) can originate from bone marrow–derived progenitor cells, circulate with the blood on mobilization from the bone marrow, and home to sites of active vessel growth, a process termed "adult vasculogenesis" (1, 2). The recruitment of EPCs is involved in tumor vascularization (3, 4) and contributes to ischemia-triggered neovascularization (5–8). Although the exact characterization of the EPCs is not entirely clear, various studies suggest that EPC develop from common endothelial and hematopoietic precursor cells, so-called adult hemangioblasts (1, 2, 9). The molecular mechanisms directing endothelial differentiation from stem or progenitor cells, however, are incompletely understood.
As a vehicle for modulating gene expression, chromatin structure remodeling plays a central role in normal development, the physiological differentiation of cells, and both embryonic and adult stem cell functions (10, 11). Indeed, the acetylation of histones is part of the complex epigenetic regulatory process determining lineage-specific gene expression and cell fate decisions by altering the local structure of chromatin (12). Previous reports suggest that the global deacetylation of histones is necessary for in vitro differentiation of embryonic stem (ES) cells (13) and oligodendrocyte lineage progression (14). The interplay between histone acetyltransferases and histone deacetylases (HDACs) is a key regulator in the dynamics of chromatin structure and function. The family of HDACs comprises at least 17 genes that are classified into three groups. Class I HDACs (HDAC1, HDAC2, HDAC3, and HDAC8) and class II HDACs (HDAC4, HDAC5, HDAC6, HDAC7, HDAC9, and HDAC10) can deacetylate histone tails and target other cellular proteins (15). Class III HDACs (Sirtuins) were identified on the basis of their similarity with sir2, a yeast transcription repressor requiring NAD+ as a cofactor (16). Importantly, the inhibition of HDAC was shown not only to block postnatal vessel growth in an animal model of tumor vascularization (17), but also to down-regulate the endothelial nitric oxide synthase (eNOS; reference 18). Thus, we hypothesized that HDAC activity may be required for endothelial differentiation of progenitor cells. Because HDAC inhibitors blocked endothelial differentiation, we further explored the down-stream mechanisms, thereby focusing on the acetylation-dependent regulation of homeobox genes (Hox's).
Hox's encode transcriptional regulatory proteins, which are characterized by a common 60-amino acid DNA-binding motif and regulate differentiation during embryonic development and tissue morphogenesis (19). Members of the Hox family of homeodomain transcription factors play important roles in the embryonic development of the cardiovascular system and also regulate angiogenesis in the adult organism (for review see reference 20). Several Hox transcription factors (e.g., HoxD3, HoxC6, and HoxC8) modulate the expression of integrins, adhesion molecules, and extracellular matrix proteins in mature endothelial cells (21, 22), whereas HoxB5 appears to be involved in the in vitro differentiation of embryonic precursor cells toward endothelial lineage (23). HoxA9, which is important for myeloid, erythroid, and lymphoid hematopoiesis (24, 25) and stem cell expansion (26), is also particularly essential for the migration and tube-forming capacity of mature endothelial cells (27) and, thus, could serve as a switch toward endothelial commitment during progenitor cell maturation.
Our present data demonstrate that HDAC inhibition abrogates the endothelial differentiation of progenitor cells and reduces the expression of the homeobox transcription factor HoxA9. Knockdown and overexpression studies revealed that HoxA9 is a critical regulator of postnatal neovascularization and acts as a master switch to direct expression of the endothelial-committed genes.
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HDAC inhibition down-regulates HoxA9 expression and EPC formation
Because homeobox transcription factors play important roles in the embryonic development of the cardiovascular system and in neovascularization in the adult organism (20, 27), we postulated that the expression of Hox proteins might be altered by HDAC inhibition. BuA or MS-275 time- and dose-dependently reduced mRNA expression (Fig. 2 a) and protein levels (17 ± 3% of control; Fig. 2 b) of HoxA9. Moreover, the down-regulation of HDAC1 by small interfering RNA (siRNA) reduced HoxA9 expression (Fig. 2 c). In contrast, the homeodomain transcription factor HoxD9 was not regulated (Fig. 2 b) and HDAC inhibitors caused only a minor reduction in mRNA levels of HoxB5 (77 ± 11%; Fig. 2 a), indicating that HoxA9 is rather specifically regulated by HDAC.
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HoxA9 regulates eNOS, VEGF-R2, and VE-cadherin gene expression
To further substantiate that HoxA9 plays a critical role in the expression of endothelial genes, we determined whether HoxA9 is required for the maintenance of endothelial marker gene expression in human umbilical vein endothelial cells (HUVECs) as a model for mature endothelial cells. The inhibition of HoxA9 expression by siRNA reduced the mRNA expression of the endothelial marker genes eNOS, VEGF-R2, and VE-cadherin in mature endothelial cells (Fig. 5, a and b). Moreover, HoxA9 siRNA inhibited eNOS protein expression (Fig. 5 c) and the generation of nitric oxide, as measured by DAF-2DA staining, from 46 ± 6 arbitrary units in scrambled oligonucleotide-treated to 11 ± 3 arbitrary units in siRNA-treated endothelial cells (n = 4, P < 0.001). HoxA9-deficient mice consistently showed a reduced eNOS protein expression in the heart (Fig. 5 d). In contrast, overexpression of HoxA9 enhanced the expression of eNOS (Fig. 5 e). These data demonstrate that HoxA9 regulates the expression of endothelial marker genes in vitro and in vivo.
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206-272 [reference 27]; Fig. 6 b). Overexpressed WT HoxA9 bound to the eNOS and VEGF-R2 promoters, whereas the HoxA9 mt showed no binding (Fig. 6 c), confirming the specificity of the interaction. To determine the promoter activation by HoxA9, we used reporter gene constructs driven by the eNOS, VEGF-R2, or VE-cadherin promoters. HoxA9 significantly increased eNOS, VEGF-R2, and VE-cadherin promoter activation (Fig. 6, d–f). Thus, HoxA9 regulates the transcription of prototypic endothelial marker genes in endothelial cells.
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HDACs comprise at least 17 genes, of which HDAC1, HDAC3, and SIRT1 are expressed in human peripheral blood–derived EPCs (unpublished data). The pharmacological inhibition of class I and II HDACs by structurally different pharmacological HDAC inhibitors abrogated HoxA9 expression and the endothelial commitment of progenitor cells. Furthermore, a specific down-regulation of HDAC1 by siRNA reduced the expression of HoxA9. In contrast, HoxD9 and HoxB5 were not significantly regulated by HDAC inhibitors, implicating a specific dependency of HoxA9 transcription on HDAC activity. HDACs are a component of the ALL-1 supercomplex, which binds to the HoxA9 promoter and is required for HoxA9 transcription (37). Thus, one may speculate that HDAC activity is necessary for the ALL-1 supercomplex to allow for transcription of HoxA9 (37).
Our data demonstrate that HoxA9 is required for postnatal neovascularization after ischemia. Previous studies additionally documented that the inhibition of HDAC blocks tumor angiogenesis (17). However, the importance of HDAC and HoxA9 for the embryonic development of vascular structures in vivo is unclear. In fact, several lines of evidence indicate that HDAC1 and HoxA9 are not essential for embryonic vessel formation. In contrast to VEGF gene deficiency, which is lethal because of impaired vessel formation even at the level of haploinsufficiency (38), or VEGF-R2 knockout mice, which die between embryonic days 8.5 and 9.5 from the lack of any vascular formation (39), no abnormalities with respect to embryonic vascularization were obvious in HDAC1–/– or HoxA9–/– mice (40, 41). In our experiments, however, HDAC inhibitors abrogated embryonic angiogenesis and vasculogenesis in the allantois ex vivo assay. This discrepancy might well be rationalized by the broad spectrum inhibitory effect of the pharmacological HDAC inhibitors used in the present study and may indicate that the specific lack of HDAC1 might be compensated for by the various other members of the HDAC family in vivo. Likewise, the lack of a severe embryonic phenotype of HoxA9–/– mice might also be caused by compensation by other members of the Hox family such as HoxB5 during embryonic development in vivo (23). However, our data clearly indicate that HoxA9 is required for adult vasculogenesis. Indeed, HoxA9–/– and HoxA9+/– mice showed a severe impairment of endothelial colony formation, blood flow recovery, and functional regeneration after the induction of ischemia. Thus, one may speculate that mechanisms regulating postnatal neovascularization are not necessarily identical to the mechanisms operational during embryonic vascular development.
Why is HoxA9 important for postnatal neovascularization? In addition to the requirement of HoxA9 for endothelial progenitor colony formation and adult vasculogenesis, our data indicate that HoxA9 directly regulates a variety of key endothelial genes that are involved in the functional maturation and activity of endothelial cells. All three HoxA9 target genes, eNOS (42, 43), VEGF-R2 (39), and VE-cadherin (44), are crucial for angiogenesis. Moreover, the reduction of HoxA9 additionally results in the reduced expression of EphB4 and, thus, inhibits endothelial migration in vitro (27). The reduction of these HoxA9 regulated genes in vivo (as shown for the eNOS) may contribute to the severe neovascularization defect of HoxA9-deficient mice. In addition, HoxA9 regulates myeloid and lymphoid hematopoiesis (25) and, therefore, may affect inflammation-mediated angiogenesis. However, a reduction of myeloid cells was detected in homozygote, but not in heterozygote, mice (25), whereas in our study heterozygote mice showed a severe impairment of neovascularization and endothelial colony forming activity. Furthermore, the overall peripheral white blood cell counts after the induction of ischemia were not notably different between HoxA9-deficient mice and WT littermates (unpublished data). Thus, the impaired neovascularization capacity of HoxA9–/– and HoxA9+/– mice is most likely not related to a general imbalance of inflammatory cells.
Finally, our data provide novel insights into the prototypic physiological mechanism inducing the maturation of endothelial cells. Shear stress is not only one of the most powerful antiatherosclerotic factors, but is also an absolute prerequisite for a functionally intact endothelial monolayer. We demonstrate that the up-regulation of the endothelial signature gene pattern is dependent on HoxA9. Not only did shear stress increase the expression of endothelial marker proteins such as eNOS and VEGF-R2, but it also concomitantly down-regulated the pan-leukocyte marker CD45, which is expressed on hematopoietic progenitor cells. These data clearly indicate that shear stress is capable of promoting the commitment of progenitor cells to an endothelial phenotype. The stimulation of HoxA9 expression and subsequent endothelial maturation by shear stress may considerably contribute to a timely recovery of the endothelial monolayer after injury to prevent atherosclerotic lesion development and restenosis formation, respectively (32, 33, 45, 46).
This study demonstrates that the HDAC-dependent transcription of HoxA9 is necessary for ex vivo maturation of progenitor cells toward the endothelial lineage. These findings not only contribute to a better understanding of postnatal endothelial maturation, but, given the pivotal role of EPCs for neovascularization of ischemic tissue and reendothelialization, may also provide important therapeutic targets. Of note, the inhibition of adult vasculogenesis by HDAC inhibitors may also contribute to the reported profound antitumor activity of HDAC inhibitors (15).
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For the preparation of Sca-1+/lin– BMC-derived EPCs, murine BM was isolated from the hindlimbs of 6–8-wk-old female C57BL/6 mice. Then, cells were filtrated using a 40-µm pore size cell strainer. After blocking with purified anti–mouse CD16/CD32 (Fc
III/II receptor) antibodies (1:100; BD Biosciences), washed cells were incubated with a biotinylated anti–lineage marker antibody cocktail (Becton Dickinson) and antibiotin microbeads (Miltenyi Biotec), and isolated with an automated magnetic cell sorting device (autoMACS; Miltenyi Biotec). Separated lin– BMCs were incubated with microbeads directly conjugated to anti–Sca-1 antibodies, and Sca-1+/lin– BMCs were isolated by a second run through the autoMACS. To assess the capacities of these immature BM stem cells, 0.5–1.0 x 106 pooled Sca-1+/lin– BMCs were plated on 24-well culture dishes coated with human fibronectin and maintained in EBM supplemented with EGM SingleQuots and 20% FCS. After 3 d in culture, adherent Sca-1+/lin– BMCs were stained with Dil-Ac-LDL and were fixed. Then, cells were stained for vWF (Acris Antibodies) and the nuclear marker TO-PRO-3 iodide (Molecular Probes), or against 10 µg/ml lectin by incubating with FITC-labeled Ulex europaeus agglutinin I (Sigma-Aldrich) for 1 h. Staining for both vWF or lectin, and Dil-Ac-LDL, was evaluated by confocal microscopy.
For the isolation of spleen-derived EPCs, murine MNCs were isolated from homogenized splenic tissue derived from HoxA9–/– (provided by H. Jeffrey Lawrence, University of California, San Francisco, Veteran's Administration Medical Center, San Francisco, CA) or WT littermates by density gradient centrifugation with Biocoll separating solution. 4 x 106 MNCs were plated on fibronectin-coated 24-well plates in 0.5 ml EBM supplemented with EGM SingleQuots and 20% FCS and were stained as described above. Outgrowing colonies were detected after 10 d.
Cell culture
Pooled HUVECs were purchased from CellSystems and cultured as previously described (27). HUVECs were exposed to laminar flow in a cone-and-plate apparatus as previously described (47). For shear stress experiments, human CD34+ hematopoietic progenitor cells were purified from MNCs by positive selection with anti-CD34 microbeads (Miltenyi Biotec; reference 28). For the cultivation of macrophages, CD14+ monocytes were purified from MNCs by positive selection with anti-CD14 microbeads (Miltenyi Biotec) using a magnetic cell sorter (Miltenyi Biotec). Purity assessed by FACS analysis was >95%. CD14+ monocytes were incubated in RPMI 1640 with 10% FCS in the presence of 50 ng/ml M-CSF to induce macrophage differentiation (29).
Allantois assay
Allantoides were isolated as previously described (48). In brief, embryos at 8.5 d postcoitum were dissected in PBS at 4°C, and the allantoides were excised and seeded into four-chambered culture slides (Nalgene Nunc International) containing 0.4 ml DMEM with 10% FCS, 2 mmol/liter L-glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin (all reagents obtained from Life Technologies). Explants were cultured at 37°C in a 5% CO2 incubator for 18–20 h in the presence or absence of the HDAC inhibitors. The allantois cultures were fixed for 20 min at 25°C with 4% paraformaldehyde, washed twice in PBS, permeabilized for 15 min at 25°C with 0.02% Triton X-100 in PBS, blocked for 1 h at 25°C with 3% BSA (Sigma-Aldrich) in PBS, and stained with PECAM-1 antibody (MEC13.3; reference 49).
Western blot analysis
Cells were lysed with buffer (20 mmol/liter Tris, pH 7.4, 150 mmol/liter NaCl, 1 mmol/liter EDTA, 1 mmol/liter EGTA, 1% Triton X-100, 2.5 mmol/liter sodium pyrophosphate, 1 mmol/liter ß-glycerophosphate, 1 mmol/liter Na3VO4, 1 µg/ml leupeptin, 1 mmol/liter PMSF) for 15 min on ice. After centrifuging at 20,000 g for 15 min at 4°C, protein content was measured according to the Bradford method. Homogenates (40 µg per lane) were separated on SDS-polyacrylamide gels and transferred to polyvinylidene difluoride membranes, which were then incubated with antibodies to p21 (BD Biosciences), HDAC1 (Abcam), HoxA9, and HoxD9 (Santa Cruz Biotechnology, Inc.), K9-K14-diacetylated histone H3 (Upstate Biotechnology), total histone H3 (Cell Signaling), extracellular signal–related kinase 1/2 (Cell Signaling), eNOS (BD Biosciences), or tubulin (NeoMarkers).
RT-PCR
Total RNA was isolated and either subjected to conventional RT-PCR or quantitative RT-PCR using the oligonucleotide primers summarized in Tables S1 and S2 (available at http://www.jem.org/cgi/content/full/jem.20042097/DC1). Quantification of mRNA was performed in a one-step RT-PCR reaction using the LightCycler (Roche Diagnostics) real-time thermocycler according to the manufacturer's instructions. Amplification was performed with 40 cycles at an annealing temperature of 61°C. Copy numbers were calculated by the instrument software (Roche Diagnostics) from standard curves of an in vitro–transcribed IL-10 cytokine primer mix (Light Cycler control kit RNA; Roche Diagnostics). The specificity of the amplification reaction was determined by a melting curve analysis.
Generation of recombinant adenovirus and adenoviral infection
HoxA9 (transcript variant 1) was amplified by RT-PCR and cloned into a shuttle pAd Track-CMV vector. This plasmid was linearized by digesting with restriction endonuclease PmeI and subsequently cotransformed into Escherichia coli BJ5183 cells with an adenoviral backbone pAdEasy-1 plasmid (all plasmids and E. coli cells were a gift from B. Vogelstein, Howard Hughes Medical Institute, Johns Hopkins Medical Institutions, Baltimore, MD). Recombinants were selected by kanamycin resistance. Finally, recombinants were transfected into HEK293 cells. Recombinant adenoviruses were generated within 7–10 d. The lacZ gene codes for the enzyme ß-galactosidase. The lacZ adenovirus was used as control.
Peripheral blood MNCs (4 x 106 cells/1-cm well) were resuspended in 2.5 ml RPMI 1640 (GIBCO BRL) with 10% FCS and preincubated for 30 min with a mixture of adenovirus, 10 µl Antennapedia peptide (RQIKIWFQNRRMKWKK; 2.5 mM; Biosyntan), and 100 µl Optimem (Life Technologies). After 24 h, 4 x 106 cells were resuspended in 1 ml EBM supplemented with EGM SingleQuots and 20% FCS and plated on fibronectin-coated wells.
Endothelial differentiation of ES cells
CJ7 ES cells, a 129/Sv-derived cell line, were cultivated as previously described (50). To initiate ES cell differentiation and embryonic body formation, ES cells were trypsinized and suspended in IMDM (Life Technologies) with 15% FBS, 10 µg/ml insulin (Sigma-Aldrich), 100 U/ml penicillin, 100 µg/ml streptomycin, 450 µmol/liter monothioglycerol, and endothelial differentiation promoting growth factors including 50 ng/ml recombinant human VEGF (PeproTech), 2 U/ml recombinant human erythropoietin (Cilag AG), 100 ng/ml human basic fibroblast growth factor (Genzyme), and 10 ng/ml murine interleukin 6 (Genzyme). After 7 d, ES cell–derived endothelial cells were collected by anti-CD31 immunomagnetic selection (51).
Reporter gene assay
Reporter gene constructs were previously described (1.6-kb human eNOS promoter fragment [reference 52]; 4-kb human VEGF-R2 promoter [reference 36]). The VE-cadherin promoter (3,032-kb fragment: –2928/+104) was cloned into KpnI–XhoI restriction sites in pGl3 enhancer plasmid (Promega). 3.5 x 105 HUVECs were transiently transfected with 3 µg plasmid DNA using 18 µl Superfect (QIAGEN) as previously described (47). After incubation, cells were lysed with lysis buffer (Promega), and luciferase activity was measured using the Luciferase System (Promega) with a luminometer (model Luminat LB 9501; Berthold).
Murine ischemic hind limb model
The effect of HoxA9 on ischemia-induced neovascularization was investigated in a murine model of hind limb ischemia. The present study was performed with the permission of the State of Hesse (Regierungspräsidium Darmstadt), according to section 8 of the German Law for the Protection of Animals, and conforms to the Guide for the Care and Use of Laboratory Animals measurements. In brief, the proximal portion of the femoral artery, including the superficial and the deep branch, as well as the distal portion of the saphenous artery, was ligated and all arterial branches between the ligations were obliterated using an electrical coagulator. The overlying skin was closed using three surgical staples. 2 wk later, we determined the morphology of the limb and measured the ischemic (right) to normal (left) limb blood flow ratio using a laser Doppler blood flow meter (model moorLDI-Mark 2; Moor Instruments). Before initiating scanning, mice were placed on a heating pad at 37°C to minimize variations in temperature. After the recording of complete scan laser Doppler color images, the perfusion of the ischemic and nonischemic limb was calculated on the basis of colored histogram pixels. To minimize variables, including ambient light and temperature, calculated perfusion was expressed as the ratio of ischemic to nonischemic hind limb perfusion.
For morphological analysis, 8-µm frozen sections of the adductor and semimembranous muscles were used. Myocyte membranes were stained using an antibody to laminin (rabbit) followed by anti–rabbit-Alexa 488. Conductance vessels in the adductor and semimembranous muscles were identified by size (>20 µm) and staining using a Cy3 labeled mouse monoclonal antibody for smooth muscle actin (Sigma-Aldrich).
Statistics
Data are expressed as mean ± SEM or as indicated in the figure legends. Two treatment groups were compared with the independent samples t test, and three or more groups by one-way analysis of variance followed by post-hoc analysis adjusted with a least significant difference correction for multiple comparisons (SPSS Inc.). Results were considered statistically significant when P < 0.05.
Online supplemental material
Fig. S1 shows a representative flow cytometric characterization of endothelial marker expression in adherent human peripheral blood MNC–derived EPCs. Fig. S2 presents a comparison of the effect of structural analoga without HDAC inhibitory capacity (2 mM acetate and 1 mM valpromide) and with HDAC inhibitory capacity (2 mM BuA and 1 mM valproate) on the ex vivo formation of EPCs from human peripheral blood MNCs during 72 h of incubation under endothelial differentiation conditions. Fig. S3 shows cell matrix adhesion. Fig. S4 presents the effect of a 72-h incubation of peripheral blood MNCs with 2 mM BuA, 10 µM MS-275, or 1 mM valproate (VPA) under endothelial differentiation conditions on apoptosis. Table S1 summarizes the sequences of the oligonucleotide primer pairs (forward and reverse) used for RT-PCR. Table S2 summarizes the sequences of the oligonucleotide primer pairs (forward and reverse) used for quantitative RT-PCR (Light Cycler). Online supplemental material is available at http://www.jem.org/cgi/content/full/jem.20042097/DC1 or from the authors by request.
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Acknowledgments |
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This work was supported by grants to S. Dimmeler from the Deutsche Forschungsgemeinschaft (Di600/2-5 and FOR501[Di600/6-1]) and to L. Rössig from the Deutsche Krebshilfe (106177). This work emanates from the European Vascular Genomics Network, a Network of Excellence supported by the European Union's sixth Framework Programme for Research Priority 1, "Life sciences, genomics, and biotechnology for health" (contract LSHM-CT-2003-503254). K.-i. Sasaki was in part supported by the Japan Heart Foundation/Bayer Yakuhin Research Grant Abroad.
The authors have no conflicting financial interests.
Submitted: 11 October 2004
Accepted: 15 April 2005
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4 (CD49d), 
6/group). (b) For morphological analysis, myocytes were identified by staining for laminin (green) in ischemic tissue of WT (left) and homozygote HoxA9–/– mice (right). (c) Conductant vessels were defined by size (>20 µm) and positive staining for 
