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. Author manuscript; available in PMC: 2020 Nov 1.
Published in final edited form as: J Cell Physiol. 2019 Apr 15;234(11):20420–20431. doi: 10.1002/jcp.28643

Hypoxic Regulation of Angiotensin-Converting Enzyme 2 and Mas Receptor in Human CD34+ Cells

Shrinidh Joshi 1, Hannah Wollenzien 1, Estelle Leclerc 1, Yagna PR Jarajapu 1,*
PMCID: PMC6660366  NIHMSID: NIHMS1021681  PMID: 30989646

Abstract

CD34+ hematopoietic stem/progenitor cells (HSPCs) are vasculogenic and hypoxia is a strong stimulus for the vasoreparative functions of these cells. Angiotensin converting enzyme-2 (ACE2)/Angiotensin-(1-7)/Mas receptor (MasR) pathway stimulates vasoprotective functions of CD34+ cells. This study tested if ACE2 and MasR are involved in the hypoxic stimulation of CD34+ cells. Cells were isolated from circulating mononuclear cells (MNCs) derived from healthy subjects (n=46) and were exposed to normoxia (20% O2) or hypoxia (1% O2). Luciferase reporter assays were carried out in cells transduced with Lentivirus carrying ACE2- or MasR- or a scramble-3’-UTR gene with firefly luciferase reporter. Expressions or activities of ACE, AT1R, ACE2 and MasR were determined. In vitro observations were verified in HSPCs derived from mice undergoing hindlimb ischemia (HLI). In vitro exposure to hypoxia increased proliferation and migration of CD34+ cells in basal conditions or in response to VEGF or SDF compared to normoxia. Expression of ACE2 or MasR was increased relative to normoxia while ACE or AT1R expressions were unaltered. Luciferase activity was increased by hypoxia in cells transfected with the luciferase reporter plasmids coding for the ACE2- or MasR promoters relatively to the control. The effects of hypoxia were mimicked by VEGF or SDF under normoxia. Hypoxia induced ADAM-17-dependent shedding of functional ACE2 fragments. In mice undergoing HLI, increased expression/activity of ACE2 and MasR were observed in the circulating HSPCs. This study provides compelling evidence for the hypoxic upregulation of ACE2 and MasR in CD34+ cells, which likely contributes to vascular repair.

Keywords: CD34+ cells, Hypoxia, ACE-2, Mas Receptor

Graphical Abstract

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Background

Accumulated evidence based on numerous laboratory and clinical studies strongly support the vasoregenerative functions of bone marrow-derived CD34+ hematopoietic stem/progenitor cells (HSPCs) and their therapeutic potential for the treatment of ischemic vascular disorders. (Mackie & Losordo, 2011; Schachinger et al., 2006) CD34+ cells home to the areas of ischemia and accomplish ischemic vascular repair largely by paracrine mechanisms and by vascular engraftment. (Jarajapu & Grant, 2010; Ziebart et al., 2008)

Sensitivity to the hypoxia-regulated factors derived from ischemic injury such as vascular endothelial growth factor (VEGF) and stromal-derived factor-1α (SDF) stimulate the vasoreparative functions of CD34+ HSPCs including migration, proliferation, vascular incorporation and the release of paracrine factors. (Jarajapu et al., 2014) Optimal expression of CXCR4, VEGFR1, and VEGFR2 enable these cells to respond to the signals of hypoxia. (Semenza, 1999; Ulyatt, Walker, & Ponnambalam, 2011) Conversely, hypoxic preconditioning increases the surface expression of CXCR4, VEGFR1, and VEGFR2, which may promote the vasoreparative functions of progenitor cells. (Tang et al., 2009) In human mesenchymal stem cells, hypoxia stimulated proliferation and tissue formation with increased Oct-4 and connexin-43 expressions and extracellular matrix formation. (Grayson, Zhao, Bunnell, & Ma, 2007) In clinical conditions that are associated with impaired innate vasoreparative potential, hypoxic desensitization of the cells has been observed. (Caballero et al., 2007; Jarajapu et al., 2011; Jarajapu et al., 2014)

Classical renin angiotensin system (RAS) consists of angiotensin converting enzyme (ACE), the octapeptide product angiotensin (Ang) II and AT1 and AT2 receptors. RAS has been expanded with the discovery of novel enzymes and peptide fragments of Ang II. It has now been well documented that ACE2, a monocarboxy peptidase, generates the heptapeptide angiotensin-(1-7) (Ang-(1-7)) from Ang II and that Ang-(1-7) produces cardiovascular protective functions by acting largely on the receptor, Mas (MasR). (Santos et al., 2003) While Ang II produces hypertensive, pro-oxidative, hypertrophic effects and pro-fibrotic effects in cardiovascular system, Ang-(1-7) elicits counter-regulatory effects on ACE/Ang II pathway by producing vasodilatory, antihypertensive, antihypertrophic, antifibrotic and antithrombotic effects. (Ferreira et al., 2010) Previous studies have provided strong evidence for the expression of local RAS in bone marrow and for a regulatory role of angiotensin peptides in the hematopoietic functions in human and murine bone marrow cells. (K. Rodgers, Xiong, & DiZerega, 2003; K. E. Rodgers, Xiong, Steer, & diZerega, 2000) We have previously shown that ACE2/Ang-(1-7)/MasR is expressed in human and murine HSPCs and that the activation of this axis stimulates vasoreparative functions of the cells. (Jarajapu et al., 2013; Singh et al., 2015) Importantly, we have demonstrated that genetic ablation of MasR impairs the migratory prowess in vitro and mobilization from bone marrow into the circulation in vivo in response to ischemic injury. (Vasam et al., 2017) This study tested the hypothesis that the expression of ACE2 and MasR is hypoxia-regulated in human and murine HSPCs, which in turn stimulates vascular repair-relevant functions in human CD34+ HSPCs. Cells were exposed to hypoxia or to the hypoxia-regulated factors, and HIF1α-dependent and -independent regulation of ACE2 or MasR expression were determined. The vascular repair-relevant functions of the cells were evaluated in vitro. Observations in human cells were confirmed in murine cells and in a mouse model of ischemic injury.

Materials and methods

Characteristics of subjects

This study was approved by Institutional Biosafety Committee of North Dakota State University. The work described has been carried out in accordance with The Code of Ethics of the World Medical Association (Declaration of Helsinki) for experiments involving humans. Leucocyte samples were derived from healthy human individuals (n=85) including both males and females of age ranging from 28 to 75 years at the United Blood Services (Fargo, ND). Participants in the study are healthy Caucasian individuals and nonsmokers. Informed consent was obtained from all participants. Leucocytes were collected in Leucoreduction chamber (LRS chambers) following apheresis by using TrimaAccel system (80440). Freshly obtained LRS cones were used for the isolation of CD34+ cells as described before. (Singh et al., 2015)

Isolation of CD34+ cells

Peripheral blood mononuclear cells (MNCs) were isolated by Ficoll-Paque (GE Healthcare Waukesha, WI, USA) density-gradient centrifugation (800 g, 30 min), according to the manufacturer’s protocol. Differentiated MNCs were collected from buffy coats and plasma was excluded by washing the cells three times with working buffer (1X PBS with 2% FBS and 1 mM EDTA) and centrifuged at 120g for 10 min. Cell count was performed by hemocytometer and 100 million MNCs were enriched for lineage negative (Lin−) cells by using a human progenitor cell enrichment kit (19356; StemCell Technologies, Vancouver, BC, Canada) as per supplier’s instructions. Unwanted cells are targeted for removal with tetrameric antibody complexes recognizing CD2, CD3, CD11b, CD11c, CD14, CD16, CD19, CD20, CD24, CD56, CD61, CD66b glycophorin A and dextran-coated magnetic particles. Lin− cells were further enriched for CD34+ cells by using a positive selection kit (18056; StemCell Technologies, Vancouver, BC, Canada) by using a cocktail containing an antibody to human Fc receptor to minimize nonspecific binding and tetrameric antibody complexes recognizing CD34. Antibody labelled cells are targeted with dextran-coated magnetic nanoparticles and separated using EasySep magnet (18000; StemCell Technologies, Vancouver, BC, Canada). The purity of enriched CD34high/CD45low cells was assessed by flow cytometry (Accuri C6, BD Biosciences). Cells were stained by incubating with APC ant-human lineage cocktail (363601; Biolegend, 1:500, Ex/Em: 650/660 nm), PE anti-human CD45 (304039; Biolegend, 1:500, Ex/Em: 565/578 nm) and FITC anti-human CD34 antibodies (343604; Biolegend; 1 in 250, Ex/Em: 490/525 nm) or isotype control antibodies (1 in 500) (Biolegend, SanDiego, CA, USA) in the presence of FcR blocking reagent (130-059-901; Miltenyi Biotech, 1:100) for 45 min at 4°C. Dead cells were excluded using 7-AAD viability staining solution (420404; Biolegend). The purity of the CD34+ cells was 96%, when evaluated randomly during the study (Supplement Figure 1).

Exposure of cells to normoxia or hypoxia

Freshly isolated cells were plated in round-bottom 96-well plates (Nunc) with no more than 30,000 cells per well in StemSpan SFEM (09650; StemCell Technologies). Cells were maintained in a CO2 incubator at 37°C in 20% O2 (normoxia) or were exposed to hypoxic conditions (1% O2-94% N2 and 5% CO2). Hypoxia was accomplished by using hypoxia chamber (ProOx C21, BioSpherix, Lacona, NY, USA). Hypoxia chamber was maintained at 37°C and was sealed and purged with nitrogen for 1 hour to achieve 1% O2, 5% CO2, and -balance N2. Preliminary studies were carried out to determine the optimum level of hypoxia and optimum duration of exposure. Among different levels of hypoxia, 0.1% and 1% hypoxia for 12 hr, showed similar increase in the ACE2 activity. Along similar lines, among different durations of exposure, increase in ACE2 protein expression and activity was similar at 12 and 24-hr exposure (Supplement Figure 2). Therefore, the rest of the study was carried out at 1% hypoxia with 12-hr exposure.

Up to 4×105 cells were exposed to hypoxia or normoxia, and cells were either used freshly for flow cytometry, migration or proliferation, or preserved for western blotting and real-time PCR.

Cell Migration and Proliferation assays

Migration was determined by QCM™ chemotaxis 5 μM, 96-well cell migration assay (ECM512; EMD Millipore) as described before. (Singh et al., 2015) Briefly, 2×104 cells were plated in the cell-migration chamber plate in the serum-free basal medium in the presence or absence of chemoattractant (SDF (100 nM), VEGF (30 nM). Cells were allowed to migrate for five hours in either normoxic or hypoxic conditions. After five hours cells were dislodged from the membrane using cell detachment buffers, lysed and quantified using fluorescent dye at 480/520 nm using Spectramax plate reader (Molecular Devices). Migratory response to SDF, VEGF or hypoxia was expressed as percent increase in migrated cells compares to untreated cells kept in normoxic condition.

Proliferation was determined by colorimetry using cell proliferation BrdU incorporation (Roche Bioscience) as described before. (Singh et al., 2015) The assay was performed using 10,000 cells treated with or without SDF/VEGF under normoxic or hypoxic conditions for 48 hours. Proliferation was evaluated by measuring absorbance at 370 nm (Spectramax plate reader) and expressed as fold increase as compared to mitomycin (1 μM), which is known to inhibit cell proliferation.

Quantitative real-time polymerase chain reaction (qRT-PCR)

RNA was extracted from CD34+cells using Trizol and the concentration and purity of RNAs were determined by absorbance spectroscopy (NanoDrop Technologies). RNA (100 ng) was reverse transcribed using a qScript cDNA synthesis kit (Quantabio) according to manufacturer’s protocol. SYBR green was used to detect DNA synthesis in real-time. Each sample contained 10 ng DNA, 20 μM of forward and reverse primers, and the iQ SYBR green containing supermix (Bio-Rad). The specific primers for ACE2, ACE, AT1 receptor (AT1R), Mas receptor (MasR) primers were synthesized by Invitrogen (list of primer sequences is provided in the Table 1 of the supplementary material). β-actin was used as an internal housekeeping gene. The reactions were run on a Stratagene Mx3000P using the following conditions: 10 min at 95°C, followed by 40 cycles (10 sec at 95°C (denaturation step), 20 sec at 60°C (annealing step), and 30 sec at 72°C (extension step). The Ct values of each gene were normalized to β-actin using the delta Ct method where delta Ct = Ct gene − Ct Actin.

ACE and ACE2 activities

ACE and ACE2 activities were determined in the CD34+ cell-lysates and cell-supernatants by using enzyme-specific fluorogenic substrates (ES005 for ACE and ES007 for ACE2; R&D Systems), respectively, as described before. (Joshi, Balasubramanian, Vasam, & Jarajapu, 2016) Enzyme-specific inhibitors, captopril and MLN-4760, were used to define ACE- and ACE2-specific enzyme activities, respectively, and enzyme-sensitive fluorescence was expressed as arbitrary fluorescence units/μg of protein/hr.

Western blotting

Cells and cell supernatants were collected after exposure to normoxic or hypoxic conditions. Cells were lysed in a radioimmuno precipitation assay (RIPA) buffer (Tris 10 mM pH 7.4, containing 140 mM NaCl, 1 mM EDTA , 1 mM NaF, 0.10% SDS, 0.50% sodium deoxycholate 0.1 % NP-40, 1% Triton X-100) in the presence of protease inhibitors (Thermo Fisher). Protein concentration in cell lysates and cell supernatants was determined using bicinchoninic acid with bovine serum albumin as a standard (Thermo Fisher). Equal amounts of protein (30μg) were loaded and separated by SDS-PAGE using SurePage 10% pre-casted gels (Genescript). Proteins were electroblotted onto nitrocellulose membranes (Bio-Rad). The blots were blocked using 5% (w/v) milk in Tris-buffered saline containing 0.5% (v/v) Tween-20. The membranes were then incubated with different antibodies. The anti-ACE2 antibodies that were used recognized either an epitope within the ectodomain of ACE2 (ab87436 (Abcam); amino acids (AAs) 350-450), an epitope within the cytoplasmic domain of ACE2 (ab15348; AAs 788-805). A third antibody (sc-20998) recognized a non-disclosed epitope located in the region of ACE2 (AAs 631-805) that includes part of the ectodomain (AAs 18-740), the transmembrane domain (AAs 741-761) and the cytoplasmic domain (AAs 762-805). Two distinct antibodies against ACE were used: antibody ab28311, directed against the first peptidase unit of ACE (AAs 30-630) and antibody GTX11737, which epitope was not disclosed by the manufacturer. Antibodies directed against the Mas receptor (sc-135063), or AT1 receptor (ab124734) were also used. β-actin antibody (mab8929; R&D Systems) was used as loading control for cell lysates. To assure that the proteins were transferred on the blot MemCode staining (ThermoFisher) was used. HRP-conjugated goat anti-mouse (405-306; Biolegend) or donkey anti-rabbit (406-401; Biolegend) secondary antibodies were used at 1:20,000 dilution as recommended by the manufacturer. Primary antibodies were diluted in Tris-buffered saline containing (0.5% v/v) Tween-20 (TBS-T) supplemented with 5% milk whereas secondary antibodies were diluted in TBS-T only. The bands were visualized by incubating the blots with enhanced chemiluminescence reagent (ECL, K15045-D50; Advansta) and developed on X-ray films (Phenix research products). Band intensities were quantified by using Image J software (NIH).

Flow cytometry

Expression of MasR and AT1R on CD34+ cells was evaluated by flow cytometry (Accuri C6, BD Biosciences). CD34+ cells were treated with FcR blocking reagent (Miltenyi Biotech) for 5 min at 4°C and washed with cell-staining buffer (420201; Biolegend). The cells were then incubated with fluorescently labeled anti-MasR or-AT1R antibodies. The fluorescence labeling was performed by using the Zenon-Alexa Fluor 488 Rabbit IgG labelling kit (Z-25302) for the MAS antibody (SC-54682) and the Zenon-R-Phycoerythrin Rabbit IgG labeling kit (Z-25355) for the AT1 antibody (ab124734) as per the manufacturer’s protocol.

Lentiviral luciferase reporter vectors

Lentiviral luciferase reporter vectors for ACE2, MasR and their respective control vectors were custom-made by Applied Biological Materials Inc., (Richmond, BC, Canada). The human ACE2 (−1541/−1537) and MasR (−974/+160) promoter regions were cloned into the pLenti-luciferase-UTR (firefly luciferase) reporter vector and were under the control of the cytomegalovirus (CMV) and SV40 enhancer promoters. (Supplement Figure 3) Sequence analysis showed that the promoter region of ACE2 contained five putative HIF-1α binding sites (−1541/−1537, −870/−866, +543/+547, +487/+491 and +149/+153) whereas the promoter region of MasR contained three putative HIF-1α sites (−921/−917, −340/−335 and +187/+191). A control lentiviral vector containing a scrambled gene sequence with CMV-driven expression of firefly luciferase was used as control in all experiments.

Additional luciferase reporter plasmids carrying miRNAs that selectively downregulate either ACE2 or MasR expression in response to hypoxia were obtained (Applied Biological Materials Inc.). The two miRNAs miR-421 and miR-143 were chosen for ACE2 and MasR respectively, based on sequence alignment analysis (blast-NCBI-NIH) using miRNA-binding prediction algorithms. Indeed, putative binding sites for miRNA 421 and miRNA 143 were found in the 3’ -UTR of the ACE2 and MasR transcript, respectively (Supplement Figure 4). Lentiviral vectors that were detailed above were redesigned to contain both pre-miRNA and luciferase inserts under the same CMV promoter with the transcription termination site after pre-miRNA, thus expression of pre-miRNA in response to hypoxia would prevent the expression of either ACE2 or MasR.

Cell transfection with the lentivirus was carried out using Lipofecatmine™ 3000 (Invitrogen), according to manufacturer’s instructions. Following 48-hr transfection, cells were washed with PBS by centrifugation (120g) to remove the excess of lentiviral particles and lipofectamine. The cells were then suspended in StemSpan in 96 well plates and incubated under hypoxia or normoxia as described above.

Luciferase Assay

Luciferase activity was measured by using Luciferase Assay kit (G287; Applied Biological Materials) as per manufacturer’s instructions. Luminescence was measured using SpectraMax luminometer (Molecular Devices) at 480 and 560 nm.

Mouse Hind-limb Ischemia

Male C57Bl/6 NHsd (wild type) (Envigo) were used in this study. All animal studies were approved by the Institutional Animal Care and Use Committee (IACUC) at North Dakota State University. All experiments were carried out in accordance with guidelines and regulations approved by IACUC. Hind-limb ischemia (HLI) surgery was carried out in mice under isoflurane anesthesia as described before.(Vasam, Joshi, & Jarajapu, 2016) Peripheral blood was collected on day 2 following HLI and circulating Lin- cells were isolated by using cell isolation kit (StemCell Technologies). Cells were tested for ACE, ACE2, AT1R and MasR, activities. Expression of ACE2 was detected using Zenon Rabbit IgG Allophycocynin labelling kit (Z-25351; ThermoFisher) for ACE2 primary antibody (ab87436; abcam) and Zenon Alexa Fluor 488 labelling kit (Z-25002; ThermoFisher) for MasR (SC-390453; Santa Cruz Biotechnology). Labelling of primary antibodies with Zenon complexes was performed as described above.

Statistical analysis

Data are presented as mean values ± S.E.M. Number of experiments ‘n’ represent the number of donors used in the experiment or number of mice in the experimental group. Statistical analyses were performed using GraphPad InStat 3.0 (GraphPad Software, San Diego, CA). Statistical differences in the mean were assessed using one-way ANOVA followed by Bonferroni’s post-test for multiple comparisons and were considered significantly different at P<0.05. Power analysis was carried out post hoc by using Minitab software (Minitab 17; PA USA) and all experiments are powered at 80% or higher.

Results

Exposure to hypoxia (1% oxygen) stimulated proliferation of CD34+ cells as determined by BrdU incorporation in basal conditions or in response to hypoxia-regulated factors, stromal-derived factor-1α (SDF) or vascular endothelial growth factor (VEGF) compared to cells exposed to normoxic conditions with 20% oxygen. Compared to normoxic cells in basal conditions, cell proliferation was increased significantly (P<0.05, n=8) in the presence of 100 nM SDF or 25 nM VEGF (Figure 1A). Along similar lines, the migratory response of CD34+ cells was potentiated by hypoxic exposure, as compared to normoxia, (P<0.05, n=8) in basal conditions (untreated) or in response to SDF or VEGF treatments (Figure 1B).

Figure 1. Hypoxia potentiates migration and proliferation of human CD34+ cells:

Figure 1.

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A. Proliferation of cells was expressed as fold-increase compared to the mitomycin (1 μM)-treated cells in normoxic conditions. Hypoxia increased this response in basal, and in stromal-derived factor 1α (SDF)- or vascular endothelial growth factor (VEGF)-stimulated conditions (P<0.05, n=8). B. Migratory response in cells expressed as percent increase over baseline response in untreated cells in normoxic (20% O2) conditions. Exposure to hypoxia (1% O2) increased this response in the untreated (basal) or in the presence of SDF (100 nM) or VEGF (n=25 nM) (n=8).

Exposure to 12-hour hypoxia increased mRNA expression of ACE2 (P<0.002, n=6) and MasR (P<0.01, n=6) but not ACE or AT1R compared to normoxia (Figure 2). In agreement with this, protein levels of ACE2 (P<0.05) and MasR (P<0.01), as determined by western blotting and flow cytometry, respectively, were increased by hypoxia (n=6) compared to normoxia, but protein levels of ACE or AT1R were unchanged. Furthermore, ACE2 activity is significantly increased in lysates of cells exposed to hypoxia compared to the cells exposed to normoxia (P<0.01, n=8).

Figure 2. Hypoxia increased the expression of ACE2 and MasR in human CD34+ cells:

Figure 2.

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A and E. Exposure to hypoxia (1% O2) increased mRNA expression of ACE2 or MasR as indicated by lower ΔCt as compared to cells in normoxia (20% O2) (n=6). B. ACE2 enzyme activity was increased in lysates derived from hypoxia-exposed cells (n=8) compared to that observed in normoxic cells. C and D. ACE2 protein levels were increased in hypoxia-exposed cells compared to normoxic cells (n=6). F, G and H. Surface expression of MasR was significantly increased in hypoxia-exposed cells compared to normoxic cells (n=6). Expressions of ACE and AT1 receptors or ACE activity were not affected by hypoxia.

Then we sought to determine if the observed effects of hypoxia on ACE2 and MasR expressions is HIF1α-dependent. Firstly, we have used a pharmacological inhibitor of HIF1α, 2-methyl estradiol (2ME), during the exposure to hypoxia. Concurrent treatment of cells with 2ME prevented the upregulation of ACE2 protein (P<0.05, n=5) or the enzyme activity (P<0.05, n=5) by hypoxia (Figure 3). Similar inhibitory effect was observed on MasR expression (P<0.05, n=5), suggesting that the observed effects of hypoxia on ACE2 and MasR are indeed HIF1α-dependent.

Figure 3. Hypoxic increase in the expression of ACE2 and MasR in human CD34+ cells is HIF1α-dependent:

Figure 3.

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Pharmacological treatment with 2-methyl estradiol (2-ME) prevented the effects of hypoxia on the protein expression of ACE2 (A and B) or MasR (D and E) (n=5). C. Increased ACE2 activity under hypoxic conditions was not observed in the presence of 2-ME (n=5).

Transcriptional regulation of ACE2 and MasR expressions by HIF1α were then evaluated by using luciferase reporter assay. Neither normoxia nor hypoxia affected the luciferase activity in cells transduced with blank-luciferase (Figure 4). ACE2-luciferase activity is significantly increased by exposure to hypoxia compared to normoxia (P<0.001, n=6), which was abolished by co-expression of miR421 in both normoxic (P<0.05) and hypoxic (P<0.001) conditions (n=6). Along similar lines, MasR-luciferase activity was increased by exposure to hypoxia (P<0.05, n=6) that was abolished by co-expression of miR143 (P<0.001) (Figure 4). It is important to note that miR 421 and miR143 expressions did not affect the expression of MasR and ACE2, respectively, confirming the target-specificity.

Figure 4. Transcriptional regulation of ACE2 and MasR by HIF1α in human CD34+ cells:

Figure 4.

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A. Luciferase activity was not changed in cells transduced with the control luciferase reporter plasmid (Blank-Luc) upon exposure to hypoxia. Luciferase activity in cells transfected with the Luciferase-ACE2 reporter plasmid (ACE2-Luc) was higher in hypoxia compared to that in normoxia (n=6). Co-expression of miR421 but not miR143, prevented the increase in luciferase activity regardless of the exposure. B. Luciferase activity in cells transfected with the Luciferase-MasR reporter plasmid (MasR-Luc) was higher in hypoxia compared to that in normoxia (n=6). Co-expression of miR143 but not miR421, prevented the increase in luciferase activity regardless of the exposure.

Hypoxia-regulated factors, SDF and VEGF, are known to stimulate proliferation and angiogenic properties of HSPCs in the ischemic areas partly via inducing gene transcription by acting on their respective receptors. (Hoeben et al., 2004; Wu & Yoder, 2009) Therefore we checked if the upregulation of ACE2 and MasR expressions is recapitulated by SDF or VEGF. Luciferase reporter assays were carried out in cells treated with either SDF or VEGF. Both SDF (100 nM) and VEGF (100 nM) increased ACE2- and MasR-luciferase activities in a normoxic environment while no activity was observed in cells expressing blank-luciferase (Figure 5). Then, we tested the effect of receptor-selective inhibitors were used to determine the involvement of specific receptor of SDF or VEGF in mediating this response. Effect of SDF was abolished by simultaneous treatment of cells with plerixafor (10 μM) confirming the involvement of CXCR4 in this response. Effects of VEGF were abolished by a nonselective VEGF receptor antagonist, axitinib (30 μM) but partially inhibited by cabozantinib (100 nM), a selective antagonist of VEGFR2 (Figure 5).

Figure 5. Transcriptional regulation of ACE2 and MasR by stromal-derived factor 1α (SDF) and vascular endothelial growth factor (VEGF) in human CD34+ cells:

Figure 5.

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Treatment with SDF or VEGF in the presence or absence of different antagonists did not alter luciferase activity in cells transduced with control luciferase (Blank-Luc). Luciferase activity was increased by treatment with either SDF (100 nM) or VEGF (100 nM) in cells transduced with either ACE2-Luc or MasR-Luc (n=6) compared to untreated cells. SDF-induced increase in luciferase activity was prevented by AMD3100 (10 μM) in both ACE2-Luc and MasR-Luc transfected cells (n=5). Cabozanitib partially reversed VEGF-induced luciferase activity in ACE2-Luc expressing cells (n=5) but did not affect the luciferase activity in MasR-Luc transfected cells (n=4). In contrast, axitinib (30 μM) prevented the effects of VEGF in both ACE-luc and MasR-luc transfected cells (n=5).

In another set of experiments, the supernatants of cells exposed to normoxia or hypoxia were analyzed for ACE and ACE2 activities. Supernatants derived from cells exposed to normoxia showed ACE and ACE2 activities (Figure 6) suggesting constitutive shedding of the ectodomains of the enzymes that retained the activity. Exposure to hypoxia increased the ACE2 activity by 4-fold in the supernatants but not that of ACE suggesting that hypoxia increased shedding of a catalytically active fragment of ACE2 ectodomain (Figure 6). This was further confirmed by Western blotting analysis, which detected ACE2 fragments that were not observed in supernatants obtained from normoxic cells (Figure 6). An antibody that recognizes an epitope located between AAs 350-450 within the ectodomain of ACE2 detected a fragment of 90 kDa. Interestingly, no fragment was detected with an antibody recognizing the cytoplasmic domain of ACE2, suggesting that the shedded ACE2 fragment lacks the cytoplasmic domain. Two fragments, a strongly stained fragment of 90 kDa and a weakly stained fragment of 80 kDa, were detected with a third antibody, raised against amino acids 631-805 of ACE2. The 80 kDa fragment was not detected by the antibody recognizing the cytoplasmic domain of ACE2, suggesting that this fragment consists of parts of the ectodomain. This fragment was not detected when the cells were treated with an inhibitor of ADAM17 (Tapi-2, 50 μM) or MMP (GM6001, 25 μM) suggesting that the 80 kDa band is a fragment of ACE2 and is not due to non-specific binding of the antibody (Figure 6).

Figure 6: Hypoxia stimulates shedding of ACE2 in human CD34+ cells:

Figure 6:

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A and B. An ectodomain-specific antibody of ACE2 detected 92 kDa ACE2 fragment in the cell supernatants, as determined by Western blotting. The levels of ACE2 fragments were higher in the hypoxia-exposed cells compared to normoxic cells (n=6). Equal protein loading in each lane was confirmed by staining of the blot with memcode (B). A cytoplasmic domain specific ACE2 antibody did not detect any protein fragments in the supernatants (n=4). A third distinct ACE2 antibody with an undisclosed epitope also recognized ACE2 fragments in supernatants derived from hypoxic cells but not in those derived from normoxic cells (n=5). C. ACE2 activity was increased in the cell supernatants upon exposure to hypoxia in comparison with normoxia. No change was observed in ACE activity in the supernatant (n=8). D and E. In the presence of TAPI-2 or GM6001, ACE2 activity or the ACE2 fragments were not observed in the supernatants upon exposure to hypoxia (n=5).

Finally, we tested in an in vivo setting, if ACE2/Mas expression is altered in HSPCs that are mobilized from bone marrow in response to ischemic insult. LSK cells were collected from the peripheral blood in mice undergoing hind limb ischemic injury. All mice have experienced 88-95% decrease in the blood flow. As shown before, (Vasam et al., 2017) mobilization of bone marrow LSK cells is maximal at day-2 post-HLI. Cells were tested for ACE, ACE2, AT1R and MasR expression or activities. Cells derived from HLI-mice have higher mRNA and protein expressions of ACE2 and MasR compared to the cells obtained from non-ischemic control mice (P<0.01, n=8) (Figure 7). No change was observed in the expression of ACE or AT1R. Activity of ACE2 is increased in the lysates derived from HLI-cells by 2-fold compared to the lysates of control cells while no change was observed in the ACE activity (Figure 7). Observed changes in ACE2 and MasR expressions or activity could be either due to the exposure of circulating cells to hypoxic environment that was induced by vascular injury or due to the actions of ischemia-induced (hypoxia-regulated factors), SDF or VEGF that are higher in the circulation following vascular injury.

Figure 7: Ischemic injury increases the expression of ACE2 and MasR in the circulating hematopoietic stem/progenitor cells (HSPCs) in mice:

Figure 7:

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A. Lin-Sca-1+cKit+ (LSK) cells were increased on day 2 following hind limb ischemic (HLI) injury (n=7). B. Cells derived from ischemic mice have increased mRNA expression of ACE2 (lower ΔCt) compared to that observed in cells from non-ischemic mice. ACE mRNA expression was unchanged by HLI (n=5). C. ACE2 activity was higher in cells obtained following HLI compared to cells from non-ischemic mice (n=5). D and E. Representative histograms of flow cytometric data determining the surface expression of ACE2. Expression of ACE2 was higher in cells obtained from ischemic compared to non-ischemic mice (n=5). F and G. Representative histograms of flow cytometric data determining the surface expression of MasR. Expression of MasR was increased in cells obtained from ischemic compared to non-ischemic mice (n=5).

Discussion

This study reports several novel findings. Exposure to hypoxia stimulates migration and proliferation in human HSPCs. Hypoxia upregulates the expression of ACE2 and MasR in HIF1α-dependent manner, and this is evidently due to enhanced transcription of ACE2 and MasR genes. SDF and VEGF increases transcription of ACE2 and MasR in hypoxia-independent manner. Exposure to hypoxia increased shedding of ACE2 ectodomain fragments in HSPCs with detectable ACE2 activity in the supernatants. In a mouse model of ischemic insult, LSK cells that were mobilized from bone marrow in response to ischemia have increased expression of ACE2 and MasR.

While a very few studies have reported the hypoxic upregulation of ACE2, this is the first report to show hypoxic regulation of MasR by using pharmacological and molecular tools. In a rat model of middle cerebral occlusion, ACE2 and MasR expressions were increased by six hours following ischemia and remained higher for 24 hours.(Joshi et al., 2016) A recent study by Chang et al observed MasR upregulation in rat embryonic cardiomyoblasts following two-hour hypoxia in vitro but not AT1R. (Chang et al., 2018) Hypoxia was shown to be protective in Ang II-induced hypertrophy via MasR-upregulation in this study. However molecular mechanisms involved in the hypoxic upregulation of MasR were not addressed in these studies.

Hypoxic regulation of ACE2 appears to be cell type-specific. In hepatocellular carcinoma-derived cells, hypoxia induced ACE2 expression in 48 hours not at 24 hours. (Clarke, Belyaev, Lambert, & Turner, 2014) AMPK activator AICAR but not metformin induced ACE2 expression via SIRT1 upregulation, which binds to ACE2 promoter and induces transcription of ACE2. IL-1β treatment also increased ACE2, which did not involve SIRT1. In pulmonary vascular smooth muscle cells, hypoxia was shown to upregulate both ACE and ACE2. (Zhang et al., 2009) ACE upregulation was progressively increased up to 48 hours, whereas ACE2 expression was increased by hypoxia at 12 hours post-exposure that was declined to normal with prolonged exposure. Intriguingly, typical HREs are identified in the promoter region but ACE2 promoter activity was not detected upon exposure to hypoxia in the luciferase assays. This study concluded that ACE2 upregulation at 12 hours of hypoxia is hypoxia-independent.

Limited information is available in regards to the regulation of ACE2 and MasR by miRs. Target sequence alignment analysis using BLAST confirmed ACE2 and MasR as targets for miR421 and miR143, respectively. Evidence for a potential regulation of ACE2 by miR421 has been shown in human cardiac myofibroblasts. (Lambert et al., 2014) Upregulation of miR421 has been implicated in the development of thrombosis. Elevated miR421 in leucocytes was observed in patients with chronic kidney disease. (Trojanowicz, Imdahl, Ulrich, Fiedler, & Girndt, 2018) The current study using luciferase reporter assays provides strong support for the hypoxic regulation of ACE2 and MasR, which was prevented in the presence of miR421 and miR143, respectively.

Our study shows evidence for the constitutive shedding of ACE and ACE2 in CD34+ cells under normoxia. Exposure to hypoxia further enhanced ACE2 expression and ACE2 shedding but not that of ACE. Cell supernatants have functional ACE2 activity, and ACE2 fragments could be detected by using two different antibodies. The presence of shed ACE2 fragments of diverse sizes has been previously reported and was shown to be largely mediated by ADAM17. (Chodavarapu et al., 2013; Pedersen, Chodavarapu, Porretta, Robinson, & Lazartigues, 2015; Xiao et al., 2012) Xiao et al (2014) detected two glycosylated ACE2 fragments shed from murine proximal tubular cells, one of 70 kDa and one of 90 kDa.(Xiao et al., 2014) Current studies show that shedding of ACE2 in the supernatant was also mediated by ADAM17 or nonspecific MMPs as the fragments were not observed in the supernatant in the presence of the respective inhibitors.

Studies with pharmacological inhibitor confirmed that ACE2 shedding is mediated by ADAM17, which is in agreement with previous reports in different cell types. The study by Jia et al characterized ACE2 shedding in human airway epithelia and showed the involvement of ADAM17, and to some extent that of ADAM10, in the shedding process. (Jia et al., 2009) Later studies reported that ADAM17-dependent shedding is modulated by phorbol esters/PKC, (Lambert, Clarke, Hooper, & Turner, 2008; Xiao et al., 2016) and importantly, calmodulin binding within the cytoplasmic tail of ACE2 would prevent shedding. (Lai et al., 2009) Other studies have showed that hypoxia upregulates ADAM17 in different cell types including human lung fibroblasts, (Chen, Lin, & Chen, 2017) tumor cell lines (Rzymski et al., 2012; Wang, Feng, & Li, 2013) and rat brain (Hurtado et al., 2001), which would amplify ACE2 shedding.

Shedding of ACE2 was reported by several studies previously in physiological or pathological settings. Elegant studies by Xia et al in a mouse model of DOCA salt-induced hypertension provided compelling evidence for an important role of ACE2 shedding in the development of neurogenic hypertension. (Xia, Sriramula, Chhabra, & Lazartigues, 2013) Increased ACE2 shedding due to increased ADAM17 expression in brain results in loss of ACE2 and impairs the protective functions in brain regions that regulate hypertension. ACE2 shedding in renal proximal tubular epithelial cells was also reported by and functional ACE2 fragments were detected in urine indicating a significant loss of local ACE2. Interestingly, shedding was enhanced by exposure of cells to D-glucose in vitro or in diabetic conditions in vivo suggesting that the protective effect of ACE2 in proximal tubules is compromised by this process in diabetes. (Chodavarapu et al., 2013; Salem, Grobe, & Elased, 2014) These observations were later confirmed in diabetic individuals undergoing renal transplants, who had increased levels of ACE2 in urine. (Xiao et al., 2012) In agreement with this finding, treatment with insulin or rosiglitazone reversed the loss of ACE2 in urine in diabetic mice, which is most likely by attenuating hyperglycemia. (Chodavarapu et al., 2013; Salem et al., 2014) ACE2 shedding by ADAM17 was also demonstrated in pancreatic islets, however it was found to have no pathophysiological significance in diabetes. (Pedersen et al., 2015) Expression of neither ACE2 nor ADAM17 was increased with the progression of diabetes in db/db mice. Inhibition of ADAM17 did not affect ACE2 levels in diabetic islets suggesting no significant functional impact of this phenomenon in this setting. (Pedersen et al., 2015)

Functional consequence of ACE2 shedding by ADAM17 appears to be dependent on the anatomical location of this interaction. In brain, ACE2 shedding by ADAM17 resulted in the depletion of ACE2, which was hypothesized to contribute to the maintenance of neurogenic hypertension. Down-regulation of ADAM17 indeed significantly alleviated neurogenic hypertension. Along similar lines, ADAM17-dependent ACE2 shedding in proximal tubular cells was shown to deplete local ACE2 levels, which was aggravated in diabetic conditions. Depletion of ACE2 in kidney potentiates the detrimental effects of local ACE/Ang II/AT1R axis.

The current study reports that hypoxic exposure increases the expression as well as shedding of ACE2 in CD34+ cells, which are destined to the areas of ischemia. In this context, ACE2 shedding would increase the local functional ACE2 activity in the peri-ischemic environment, which indeed opposes the detrimental effects of local Ang II. Thus, hypoxic upregulation of ACE2 and MasR expression, and ACE2 shedding would contribute to the ischemic vascular repair. This beneficial effect would further be amplified by concurrent exposure of cells to hypoxia-regulated factors, SDF and VEGF, which also stimulate the expression of ACE2 and MasR.

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Acknowledgments:

Authors acknowledge the support by staff and volunteers at the United Blood Systems, Fargo, ND, for kindly providing leucocyte samples for the study.

Sources of funding:

This study was partly supported by American Heart Association grant (17AIREA33700012) and NIH-National Institute of General Medical Sciences (NIGMS) and National Institute of Aging (NIA) (AG056881).

The Core Biology Facility at North Dakota State University was made possible by National Institute of General Medical Sciences of NIH, P30-GM 103332-01.

Nonstandard Abbreviations

HSPC

Hematopoietic Stem/Progenitor cells

LSK cells

Lineage-negative, Sca-1 positive, cKit-positive Cells

SDF

Stromal-derived factor 1α

VEGF

Vascular Endothelial Growth Factor

CXCR4

C-X-C Chemokine Receptor Type 4

VEGFR

Vascular Endothelial Growth Factor Receptor

RAS

Renin Angiotensin System

ACE

Angiotensin Converting Enzyme

ACE2

Angiotensin Converting Enzyme-2

Ang

Angiotensin

AT1R

Angiotensin Receptor Type 1

AT2R

Angiotensin Receptor Type 2

MasR

Mas Receptor

HIF1α

Hypoxia-Inducible Factor 1α

MNC

Mononuclear Cells

PBS

Phosphate Buffered Saline

FBS

Fetal Bovine Serum

EDTA

Ethylenediaminetetracetic acid

HLI

Hind Limb Ischemia

Footnotes

Conflicts of interest: None.

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