[go: up one dir, main page]

Skip to main content
Proceedings of the National Academy of Sciences of the United States of America logoLink to Proceedings of the National Academy of Sciences of the United States of America
. 2006 Aug 18;103(35):13156–13161. doi: 10.1073/pnas.0604203103

Myeloid lineage progenitors give rise to vascular endothelium

Alexis S Bailey *,†,, Holger Willenbring *,§,‡,, Shuguang Jiang *,†, Daniel A Anderson *,†, David A Schroeder *,, Melissa H Wong *,**, Markus Grompe *,§, William H Fleming *,†,††
PMCID: PMC1559769  PMID: 16920790

Abstract

Despite an important role in vascular development and repair, the origin of endothelial progenitors remains unknown. Accumulating evidence indicates that cells derived from the hematopoietic system participate in angiogenesis. However, the identity and functional role of these cells remain controversial. Here we show that vascular endothelial cells can differentiate from common myeloid progenitors and granulocyte/macrophage progenitors. Endothelial cells derived from transplanted bone marrow-derived myeloid lineage progenitors expressed CD31, von Willebrand factor, and Tie2 but did not express the hematopoietic markers CD45 and F4/80 or the pericyte markers desmin and smooth muscle actin. Lineage tracing analysis in combination with a Tie2-driven Cre/lox reporter system revealed that, in contrast to bone marrow-derived hepatocytes, bone marrow-derived endothelial cells are not the products of cell fusion. The establishment of both hematopoietic and endothelial cell chimerism after parabiosis demonstrates that circulating cells can give rise to vascular endothelium in the absence of acute radiation injury. Our findings indicate that endothelial cells are an intrinsic component of myeloid lineage differentiation and underscore the close functional relationship between the hematopoietic and vascular systems.

Keywords: cell fusion, differentiation, endothelial cells, hematopoietic stem cells


The identification and isolation of adult endothelial progenitor cells are of significant interest because of their potential use in targeted therapies of the vascular system (15). Human circulating and bone marrow-derived endothelial progenitor activity has previously been reported in transplant recipients (6, 7). However, functional studies of human endothelial progenitors (811) have mainly relied on in vitro assays. Although expression of the myelomonocytic marker CD14 appears to be a common feature of these circulating progenitors, it has been challenging to determine both the phenotype and origin of these cells. By contrast, marker gene expression in genetically engineered mouse models permits the tracking of donor progenitor cell fates in vivo. Furthermore, if donor and recipient mice with complementary transgenes are used, this provides a means to distinguish cell differentiation from cell fusion (12, 13).

Recently we (14) and others (1517) demonstrated that a single hematopoietic stem cell (HSC) can give rise to both blood cells and vascular endothelial cells. This finding raises the fundamental question of whether endothelial cells are the differentiated progeny of an established hematopoietic lineage or represent a novel HSC-derived progenitor population (18). Hence, we sought to determine whether there was a prospectively identifiable subset of bone marrow cells outside the HSC population that could give rise to vascular endothelium. Here we report that vascular endothelial cells can arise from common myeloid progenitors (CMPs) and the more mature granulocyte/macrophage progenitors (GMPs). Importantly, neither radiation-induced vascular injury nor cell fusion is required to generate endothelial cell outcomes.

Results

To quantify endothelial progenitor activity in distinct bone marrow populations defined by lineage and progenitor status, genetically marked cells derived from adult mice were fractionated based on cell surface marker expression and transplanted into irradiated wild-type recipients. The potential problem of variable expression of transgenic donor cell markers (19) was prospectively addressed by using two independent sources of donor bone marrow cells that were genetically marked with either β-gal (ROSA26) (20) or EGFP (21). In addition to their characteristic tissue distribution and morphology, endothelial cells were identified based on the expression of CD31 and Tie2 and the absence of hematopoietic (CD45) macrophage (F4/80) and pericyte (desmin and smooth muscle actin) marker expression. The functional status of these endothelial cells was confirmed by the production of the procoagulant von Willebrand factor (VWF). Colocalization of donor and endothelial cell markers in individual cells was evaluated by deconvolution microscopy.

To determine whether a specific hematopoietic cell surface marker was informative for identifying cells with endothelial cell potential, bone marrow cells were sorted into lineage marker-positive and lineage marker-negative subpopulations. These cells were transplanted into lethally irradiated recipients at a dose based on their frequency in 1 × 106 bone marrow cells. For example, the B cell lineage marker B220 is expressed on 20% of bone marrow cells; therefore, recipients were injected with either 2 × 105 B220-positive cells or 8 × 105 B220-negative cells (Fig. 1a). When the incorporation of either ROSA26- or EGFP-marked donor cells into liver vascular endothelium was assayed, all endothelial cells arose from the B220-negative subset of bone marrow, indicating that cells committed to the B lymphocyte lineage (B220+) do not have significant endothelial cell potential (Fig. 1b). By contrast, 100% of endothelial cell activity was segregated to the c-kit-positive bone marrow subset despite a 24-fold higher input dose of c-kit-negative cells. Further evaluation revealed that both stem cell antigen-1 (Sca-1)-negative cells and cells that expressed mature hematopoietic lineage markers (Lin+) including Ter119 (erythrocytes) and Mac-1 and Gr-1 (myelomonocytic cells) gave rise to endothelium (Fig. 1b). Thus, bone marrow-derived endothelial cell progenitors reside outside of the stem cell population, consistent with the hypothesis that endothelial cells can arise from the differentiated progeny of HSCs. These findings, in combination with the absence of endothelial cell outcomes derived from lymphoid progenitors (data not shown), prompted us to investigate whether endothelial cell progenitors represented the normal developmental potential of myeloid lineage precursors.

Fig. 1.

Fig. 1.

Bone marrow-derived progenitor cells have endothelial cell potential. Based on the expression of hematopoietic cell markers, sorted populations of bone marrow progenitors were injected into lethally irradiated recipient mice at a dose equivalent to their frequency in 1 × 106 bone marrow cells. (a) Fractionation of bone marrow cells based on the expression of the B cell marker B220. (b) Portal vein branches were assayed for donor-derived endothelial cells after transplanting either marker-positive (filled bars) or marker-negative (open bars) fractions of bone marrow. The percentage of vessel cross-sections containing EGFP or ROSA26 donor-derived endothelial cells is indicated. Both Sca-1-negative and lineage-positive (Lin+) subpopulations of bone marrow gave rise to endothelial cells, indicating the existence of endothelial progenitors distinct from the HSC phenotype. ∗, below level of detection of 0.05%. Error bars indicate SEM. (c) Isolation of HSCs [c-kit+Sca-1+Lin (KSL)], CMPs, and GMPs. Lineage-negative cells (Left) were sorted for c-kit and Sca-1 (Center). Sca-1, c-kit+ cells were further fractionated into CMP and GMP populations based on CD34 and FcγR expression (Right). ∗∗, KSL lineage marker mixture differs from CMP/GMP lineage markers as indicated in Methods. (d) Transplantation schema for detecting donor-derived endothelial cells after transplantation of CMPs and GMPs.

Akashi et al. (22) identified CMPs and GMPs in adult bone marrow. GMPs are known to be further differentiated progeny of CMPs and have lost the capacity to produce erythrocytes and platelets. Both GMPs and CMPs have limited self-renewal capacity, and their short-lived, mature progeny typically are not detected in the peripheral blood for >3–4 weeks after transplantation (22, 23). To test the endothelial cell potential of these myeloid lineage progenitors, 5–10 × 103 GMPs or CMPs isolated from either ROSA26- or EGFP-expressing bone marrow were transplanted into lethally irradiated recipients along with a radioprotective dose of host-type bone marrow cells (Fig. 1 c and d). Before evaluating the recipient blood vessels for donor-derived endothelium, the peripheral blood was analyzed to ensure that neither the sorted CMP or GMP populations contained any contaminating multilineage stem/progenitor cells. No donor multilineage hematopoiesis was observed in four independent experiments evaluating a total of 25 transplant recipients.

To confirm that any long-lived, donor-derived hematopoietic progeny of CMPs and GMPs in the perivascular space were distinguished from bone marrow-derived endothelial cells, a detailed phenotypic analysis was performed (Fig. 2). The endothelial cell phenotype (CD31+, VWF+, CD45, F4/80, and DAPI+) was distinct from that of platelets (CD31+, VWF+, CD45, F4/80, and DAPI), macrophages including Kupffer cells (CD31+, VWF, CD45+ or CD45, and F4/80+), granulocytes (CD31+, VWF, CD45+, and F4/80), lymphocytes (CD31 or CD31+, VWF, CD45+, and F4/80), and pericytes (CD31, VWF, smooth muscle actin+, and desmin+). Colocalization of these markers in individual ROSA26- or EGFP-positive donor-derived cells was confirmed by deconvolution microscopy (Fig. 2 d and e and Fig. 5, which is published as supporting information on the PNAS web site). Hematopoietic cells, tissue-resident macrophages, and pericytes were readily detected and easily distinguished from endothelium in the liver parenchyma (Fig. 2f and Fig. 6, which is published as supporting information on the PNAS web site). Liver tissue was evaluated as early as 2–3 weeks after transplantation, and the frequency of portal vein cross-sections that contained donor endothelial cells was determined. Of a total of 7,300 portal vein cross-sections from recipients transplanted with CMPs, a mean of 1.3% contained donor-derived endothelial cells. Analysis of 6,060 portal veins in the GMP transplant recipients revealed that the frequency of portal veins containing donor endothelial cells was 0.8% (Fig. 2g). GMPs are the more differentiated progeny of CMPs (24); therefore, our results indicate that endothelial cell potential persists at least until the GMP stage of myeloid differentiation. When endothelial cell outcomes were examined at 6 months after transplant, no donor-derived endothelium was detected in recipients of either ROSA26- or EGFP-marked myeloid progenitors (data not shown). This finding is consistent with the limited lifespan and the modest self-renewal potential of both the CMP and GMP populations (22). In contrast, long-lived HSCs, with a high self-renewal potential, produce a modestly increased frequency of endothelial cells over this time period (14, 17).

Fig. 2.

Fig. 2.

Myeloid lineage progenitor cells give rise to vascular endothelial cells. After transplantation of either CMPs or GMPs, recipient livers were analyzed for the presence of donor-derived endothelial cells. (a) X-Gal detection (blue) of a ROSA26-marked GMP-derived endothelial cell expressing VWF (red). (Inset) Higher magnification. (b) X-Gal detection of a CMP-derived endothelial cell. (c) An EGFP+ (green) CMP-derived VWF+ (red)-expressing endothelial cell. (Inset) Higher magnification. (d) Deconvoluted images of a CMP-derived β-gal+ (green), VWF+ (red), DAPI+ (blue) endothelial cell (arrow in Left) that is negative for CD45 (magenta) expression (Center). A single host-derived CD45+ (magenta), VWF blood cell is present next to the vessel wall (arrowhead). (Right) Merged image. (e) Z-stack images of a β-gal+ (green), VWF+ (red), DAPI+ (blue) endothelial cell taken at 0.5-μm intervals demonstrating colocalization of β-gal and VWF expression in a single cell. (f) A host-derived F4/80+, CD45+ cell (magenta, arrowhead) in the liver parenchyma of a GMP transplant recipient. (Inset) A donor-derived F4/80+, CD45+ hematopoietic cell (green). The same cell is indicated with an arrow (magenta). (g) Frequency of portal vessels containing donor-derived endothelial cells after transplantation of either CMPs or GMPs. (Scale bars: a and c, 20 μm; b, d, and f, 5 μm.) L, vessel lumen.

Although our data are consistent with the differentiation of CMPs and GMPs into endothelial cells, the results of recent studies led us to evaluate cell fusion between the transplanted myeloid cells and host endothelial cells as an alternative mechanism. We (13) and others (25) have shown that correction of the metabolic liver disease fumarylacetoacetate hydrolase (Fah) deficiency by transplantation of HSCs is the result of fusion between donor hematopoietic cells and host cells. Moreover, when the progeny of HSCs were evaluated, we found that myelomonocytic cells fused with the diseased hepatocytes (23). To exclude cell fusion as the mechanism responsible for the generation of donor-marked endothelium, we used experimental strategies that have recently been used to detect fusion-derived hepatocytes, cardiomyocytes, and Purkinje neurons (12). Briefly, HSCs isolated from Cre recombinase transgenic mice were transplanted into mice harboring a β-gal gene inactivated by a floxed stop codon (R26R) (20). Thus, only host cells that fuse with donor cells will undergo DNA recombination and express β-gal. To ensure successful recombination in endothelial cell fusion products, the Tie2 promoter was used to drive Cre expression (26). Identification of donor-derived cells that were not the product of cell fusion was facilitated by using HSCs derived from mice that ubiquitously express EGFP (19). Moreover, to simplify the detection of hepatocyte fusion events, R26R recipient mice were backcrossed to render them Fah-deficient (13).

After transplantation, both donor-derived hepatocytes and endothelial cells were observed at the expected frequencies (14, 23). None of the EGFP-positive endothelial cells expressed the β-gal reporter gene, consistent with direct endothelial cell differentiation of the transplanted cells (Fig. 3). However, EGFP-positive hepatocytes expressed β-gal, as was evident from both X-Gal reactivity and immunostaining, a finding indicative of cell fusion (Fig. 3 ae). Because wild-type as well as Fah-deficient hepatocytes lack Tie2 expression (data not shown), this result could only be attributed to Tie2 activity in the hepatocyte’s hematopoietic fusion partner. Apart from endothelial cells, HSCs (27) and, more recently, angiogenic myeloid cells (28) have been isolated from mouse bone marrow and peripheral blood based on Tie2 expression. Fusion with these cells could result in β-gal activation in the host hepatocytes and explain this result. Thus, Tie2 expression might distinguish the hepatocyte’s hematopoietic fusion partner from other myelomonocytic cells (23).

Fig. 3.

Fig. 3.

Donor-derived endothelial cells are not products of cell fusion. (ae) Donor hematopoietic cells fuse with host hepatocytes. (b) X-Gal detection of fusion-derived hepatocytes (blue) after transplantation of EGFP, Tie2-Cre HSCs into a R26R, Fah recipient [portal vein (PV)]. (Inset) Higher magnification. (ce) Fusion-derived hepatocytes are both EGFP+ (green channel in c) and β-gal+ (red channel in d). (e) Merged image of c and d. (fi) Donor-derived endothelial cells do not arise through cell fusion. (g) A donor-derived, EGFP+ (green) endothelial cell (arrowhead) expresses the endothelial cell marker CD31 (magenta). (h) The same cell does not express the β-gal marker (red). (i) A Tie2 (red)-expressing EGFP+ (green) endothelial cell showing activation of the Tie2 promoter. (jm) The Cre recombinase experimental approach in the reverse direction was performed by transplanting EGFP, R26R HSCs into Tie2-Cre recipients. (k and l) An endothelial cell (arrowhead) that expresses both the donor marker (green) and CD31 (magenta) (k) and is negative for β-gal expression (not red) (l). (m) Tie2 expression (red) in a donor-marked endothelial cell (green). (nr) To evaluate cell fusion without the requirement of Cre-mediated recombination, EGFP+ HSCs were transplanted into ROSA26 mice. Recipient livers were analyzed for the presence of fusion-derived EGFP+ (green), β-gal+ (red) endothelial cells and hepatocytes. (o) EGFP+ (green) donor-derived endothelial cells were negative for β-gal staining (box), whereas host endothelial cells were β-gal-positive (red, arrow). (p) High magnification of the EGFP+, β-gal endothelial cell shown in o. (q and r) An EGFP+ donor-derived hepatocyte (q) that coexpresses β-gal (r), indicating that the hepatocyte is fusion-derived. (Scale bars: b, 20 μm; ce, gi, km, and pr, 5 μm; o, 10 μm.) L, vessel lumen.

Although Tie2 expression was readily detected in donor-derived endothelial cells (Fig. 3i), the reverse experiment was performed to rule out lack of Tie2 expression in the donor cell nucleus as the cause for failed R26R activation. HSCs expressing both EGFP and R26R were transplanted into Tie2-Cre transgenic recipients (Fig. 3 jm). In support of our initial findings, none of the donor-derived, EGFP-positive endothelial cells expressed the β-gal reporter gene, indicating that these cells were not fusion products. Successful recombination depends not only on the activity of the promoter driving Cre, but also on accessibility of the R26R locus. Thus, recombination efficiencies might differ between hepatocytes and endothelial and hematopoietic cells. Therefore, by transplanting EGFP-expressing donor cells into ROSA26 recipients (29), a simple and independent labeling approach was additionally used to exclude cell fusion (Fig. 3 nr). As expected, the vast majority of EGFP-positive hepatocytes also expressed β-gal, whereas none of the EGFP-positive endothelial cells were β-gal-positive. The combined results from these three experimental strategies indicate that endothelial cells derived from hematopoietic progenitors are not products of cell fusion but that these cells arise by differentiation of bone marrow progenitors.

To facilitate hematopoietic engraftment, transplantation models typically require the preconditioning of recipients with ionizing radiation. In addition to ablating stem and progenitor cells in the bone marrow, radiation exposure is toxic to vascular endothelial cells (30). This issue raises the important question of whether injury is required for the contribution of circulating progenitor cells to the endothelium. Consequently, we used parabiosis as a noninjury model to determine the contribution of circulating endothelial cell progenitors under steady-state conditions (31, 32). EGFP-expressing and wild-type mice were surgically joined for 12 weeks, separated, and analyzed 4–6 months later. Each member of the parabiotic pair showed a mean of 17% donor-derived hematopoietic cells in the peripheral blood (Fig. 4a). Circulating, donor-derived T cells, B cells, and myelomonocytic cells were found at normal frequencies (Fig. 4b). The successful transfer of self-renewing HSCs was confirmed by serial transplantation (data not shown). When the portal veins were examined in detail, the mean frequency of vessels demonstrating engraftment of EGFP+, CD45, CD31+, or VWF+ endothelial cells was 3.9%. Donor-derived endothelial cells were also detected in other tissues including the skin and kidney (Fig. 7, which is published as supporting information on the PNAS web site). In corroboration of our findings in irradiated recipients, additional experiments with parabiotic pairs of EGFP and ROSA26 transgenic mice did not reveal any evidence for cell fusion in the vascular endothelium (data not shown). These results demonstrate that circulating endothelial cell progenitors contribute to the vascular endothelium during steady-state conditions outside of the context of radiation-induced vascular injury.

Fig. 4.

Fig. 4.

Donor-derived endothelial cells contribute to the vascular endothelium after parabiosis. Mice were separated and analyzed for donor-derived cells in the blood and in blood vessels. (a) Flow cytometric analysis of the peripheral blood of a wild-type mouse after parabiosis with a transgenic, EGFP-expressing mouse reveals the presence of donor-derived blood cells. (b) The percentage of B cells (B220), T cells (CD3), and myeloid cells (Mac-1/Gr-1) of host (open bars) and donor (filled bars) origin; n = 5, error bars indicate SEM. (c) The frequency of donor-derived endothelial cells in the liver vessels of individual parabiotic recipients.

Discussion

Our results demonstrate that vascular endothelial cells can arise from a well defined population of myeloid lineage-restricted bone marrow progenitors, suggesting that endothelial cells represent a previously unrecognized differentiation potential of the myeloid lineage. This finding is consistent with results from several laboratories that have reported circulating cells with myeloid features contributing to human angiogenesis (811). Importantly, we have also shown that donor-derived endothelial cells differentiate from myeloid progenitors and are not products of cell fusion. Whereas transplantation of HSCs results in long-term contribution to the hematopoietic and the vascular systems, transplantation of myeloid progenitors provides only short-term engraftment of hematopoiesis and endothelial cells. This result was anticipated given the limited self-renewal capacity of these progenitor cells (22). Remarkably, the development of endothelial cells from circulating hematopoietic progenitors occurs in the absence of acute radiation injury, suggesting a role for these cells during steady-state conditions.

A notable feature of recent studies in both mice and humans is the similar frequency of long-term incorporation of bone marrow-derived cells into vascular endothelium. Studies evaluating sex-mismatched human bone marrow transplant recipients from our laboratory (6) and others (7) indicate that 2–5% of endothelial cells are donor-derived. This frequency of donor-derived endothelial cells is indistinguishable from that observed in our previously reported (14) mouse transplant model (3–4%) and is similar to recent reports by Larrivee et al. (17). Taken together, these results suggest that bone marrow-derived endothelial cells contribute to 1–5% of the endothelium in the liver, gut, and skin.

Several groups have used innovative approaches to demonstrate that growth factors and cytokines can recruit large numbers of myelomonocytic cells from the bone marrow to a variety of tissues, where they secrete factors that promote endothelial cell proliferation and differentiation (3335). Using a tissue-specific angiogenesis model based on VEGF overexpression, Keshet and colleagues (34) showed a massive recruitment of proangiogenic myeloid cells and a relatively rare frequency of incorporation of these cells into the endothelium. These studies further extend the concept that myelomonocytic cells can serve as an important source of proangiogenic cytokines in the setting of acute ischemic injury (9).

The coexistence of myeloid progenitors capable of endothelial differentiation and proangiogenic myeloid accessory cells likely represents two complementary mechanisms by which myeloid lineage progenitors contribute to angiogenesis. The relative importance of each mechanism may depend on the type of repair or regeneration being studied. For example, our finding of endothelial cell differentiation in a parabiotic model is consistent with circulating progenitors contributing to vascular homeostasis. Our current study clearly identifies myeloid progenitors as a source of vascular endothelium. Because myelomonocytic cells are known to effectively infiltrate both normal and diseased tissues, future studies on the contribution of myeloid lineage-derived endothelial progenitors to vascular regeneration and repair are needed.

Methods

Mice.

All donor and recipient mice were on a C57BL/6 background. B6.129S7-Gt(ROSA)26Sor/J (ROSA26), C57BL/6-TgN(ACTB-EGFP)Osb-YO-1 (EGFP), B6.Cg-Tg(Tek-cre)12Flv/J (Tie2-Cre), and B6.129S4-Gt(ROSA)26Sortm1Sor/J (R26R) mice were used as donors for Ka-Thy1.1-Ly5.2, R26R, FahΔexon5 (Fah−/−), and Tie2-Cre recipients. All procedures were approved by the institutional animal care and use committee of the Oregon Health & Science University.

Isolation of HSCs and Myeloid Progenitors.

For HSC isolation (14), single-cell suspensions of bone marrow were labeled with the antibodies c-kit-allophycocyanin (APC), Ly6AE-B SA-APC-Cy7 (Sca-1), and a phycoerythrin (PE)-conjugated lineage mixture (B220, CD3, CD5, CD4, CD8, Mac-1, Gr-1, and Ter119). HSCs were enriched to homogeneity by double-sorting c-kit+Sca-1+Lin cells by using a FACSVantage (Becton Dickinson, San Jose, CA). For myeloid progenitor isolation (23), cells were labeled with a PE-conjugated lineage mixture [B220, CD3, CD4, CD8, Gr-1, CD19, IgM (e-biosciences, San Diego, CA), and Ter119] and the antibodies IL-7Rα-PE (e-biosciences), Ly6AE-PE, c-kit-APC-Cy7 (e-biosciences), CD34-FITC, and FcγRII/III-APC (e-biosciences). Purified progenitors were obtained by double-sorting of IL-7RαLinSca-1c-kit+CD34+FcγRII/IIIlo (CMP) and IL-7RαLinSca-1c-kit+CD34+FcγRII/IIIhi (GMP) populations. Dead cells were excluded by a combination of scatter gates and propidium iodide staining. All antibodies were purchased from BD Pharmingen (San Diego, CA) unless otherwise indicated.

Transplantation.

Myeloablative conditioning was performed with two doses of 600 cGy each (delivered 3 h apart) of total body irradiation by using a 137Cs γ-source. HSCs and CMPs/GMPs were injected into the retroorbital venous plexus. Recipients received antibiotic water for 1 month after transplantation (neomycin sulfate at 1.1 g/liter and polymyxin B sulfate at 167 mg/liter).

Hematopoietic Engraftment.

Peripheral blood leukocytes were obtained after erythrocyte depletion by sedimentation in 3% dextran (Amersham Pharmacia, Uppsala, Sweden) and hypotonic lysis. Multilineage hematopoietic engraftment was analyzed with the antibodies CD45.2-FITC (donor), CD45.1-PE (host), and the lineage markers Mac-1-APC, Gr-1-APC, B220-APC, and CD3-APC.

Immunohistochemistry.

Recipient tissues harvested at the indicated time points after transplantation were fixed in 4% paraformaldehyde, washed, dehydrated in 30% sucrose, then cryopreserved in OCT medium. Tissue sections (5 μm) were incubated with blocking buffer, then incubated overnight at 4°C with the primary antibodies anti-VWF (1:400; DakoCytomation, Glostrup, Denmark), anti-CD31 (1:100; BD Pharmingen), anti-Tie2 (1:25; Chemicon, Temecula, CA), anti-smooth muscle actin (1:400; Sigma, St. Louis, MO), anti-desmin (1:400; DakoCytomation), or anti-β-gal (1:400; Immunology Consultants Laboratory, Newburg, OR) followed by incubation with Cy3-conjugated secondary antibody (Chemicon). The primary antibodies anti-F4/80 (SEROTEC, Raleigh, NC) and anti-CD45 (1:25; BD Pharmingen) were followed by incubation with Alexa Fluor 647-conjugated secondary antibody (Molecular Probes, Eugene, OR). Nuclei were counterstained with DAPI. Sections were examined and photographed with an Axiovert 200 microscope by using a true-color AxioCam camera and standard epifluorescence filters for FITC, Cy3, Cy5, and DAPI (Zeiss, Jena, Germany). Images were digitally combined by using AxioVision software (Zeiss). Z-stack images were obtained with a Bio-Rad 1024 ES laser scanning microscope (Bio-Rad, Hercules, CA) and analyzed by using DeltaVision deconvolution software (Bio-Rad).

Detection of ROSA26 Donor Progeny.

Recipient tissues were analyzed for β-gal activity in PLP (periodate/lysine/2% paraformaldehyde)-fixed frozen sections. Sections were incubated at 37°C with 1 mg/ml X-Gal (Sigma), 2 mM magnesium chloride, 5 mM potassium ferrocyanide, and 5 mM potassium ferricyanide. Tissue sections were either counterstained with Nuclear Fast Red (Biomeda, Hayward, CA) or stained with the primary antibody anti-VWF (1:400) followed by alkaline phosphatase-conjugated swine anti-rabbit (1:25; DakoCytomation) and detected with the Fast Red Substrate System (DakoCytomation).

Parabiosis.

Pairs of 6- to 12-week-old age- and weight-matched female mice were housed together for at least 1 week before the parabiotic surgery. Mice were anesthetized with a xylazine and ketamine HCl mixture (100 mg/kg ketamine and 10 mg/kg xylazine) and treated with one dose of ketoprofen at 5 mg/kg s.c. before surgery. The surgical site was shaved and sterilized, a small lateral incision was made from the hip to the shoulder, and the skin was freed of attached tissue. The mice were aligned, and the opposing shoulder and thigh muscles were sutured with two discontinuous sutures by using Polysorb 5-0 material. The suture was passed underneath the paired humeri and femur to prevent torsion. A continuous suture was also passed snugly through the peritoneum of each animal. The skin was aligned and joined with 9-mm wound clips. The remainder of the wound was closed with 4-0 silk suture. Buprenex (0.05–0.1 mg/kg s.c.) was administered twice daily and ketoprofen (5 mg/kg s.c.) was administered once daily for the first 2 days after the operation. Each mouse also received a daily dose of 0.5–1.0 ml of warmed saline to ensure proper hydration. Half of the wound clips were removed after 1 week, and the remaining clips were removed after 2 weeks. Each parabiotic recipient (average weight 20 g) was given recombinant human granulocyte colony-stimulating factor (250 μg/kg s.c.) for 4 days at 2–3 weeks after surgery (32). Peripheral blood was collected and analyzed by flow cytometry as previously described to evaluate donor chimerism through cross-circulation. Mice were separated at 12 weeks after surgery, and peripheral blood, liver, kidney, and skin were analyzed 4–6 months later.

Supplementary Material

Supporting Figures

Acknowledgments

We thank Aurelie Snyder for deconvolution microscopy, Mandy Boyd for cell sorting, Dana Pfaffle for assistance with immunofluorescence staining, and Devorah Goldman for helpful comments on the manuscript. This work was supported by National Institutes of Health Grants HL069133 and HL077818 (to W.H.F.), DK068326 and CA118235 (to M.H.W.), and DK-51592 (to M.G.).

Abbreviations

GMP

granulocyte/macrophage progenitor

HSC

hematopoietic stem cell

CMP

common myeloid progenitor

PE

phycoerythrin

APC

allophycocyanin

VWF

von Willebrand factor

Fah

fumarylacetoacetate hydrolase

Sca-1

stem cell antigen-1.

Footnotes

Conflict of interest statement: No conflicts declared.

This paper was submitted directly (Track II) to the PNAS office.

See Commentary on page 12959.

References

  • 1.Rafii S., Lyden D. Nat. Med. 2003;9:702–712. doi: 10.1038/nm0603-702. [DOI] [PubMed] [Google Scholar]
  • 2.Jain R. K., Duda D. G. Cancer Cell. 2003;3:515–516. doi: 10.1016/s1535-6108(03)00138-7. [DOI] [PubMed] [Google Scholar]
  • 3.Coultas L., Chawengsaksophak K., Rossant J. Nature. 2005;438:937–945. doi: 10.1038/nature04479. [DOI] [PubMed] [Google Scholar]
  • 4.Gariano R. F., Gardner T. W. Nature. 2005;438:960–966. doi: 10.1038/nature04482. [DOI] [PubMed] [Google Scholar]
  • 5.Ferrara N., Kerbel R. S. Nature. 2005;438:967–974. doi: 10.1038/nature04483. [DOI] [PubMed] [Google Scholar]
  • 6.Jiang S., Walker L., Afentoulis M., Anderson D. A., Jauron-Mills L., Corless C. L., Fleming W. H. Proc. Natl. Acad. Sci. USA. 2004;101:16891–16896. doi: 10.1073/pnas.0404398101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Peters B. A., Diaz L. A., Polyak K., Meszler L., Romans K., Guinan E. C., Antin J. H., Myerson D., Hamilton S. R., Vogelstein B., et al. Nat. Med. 2005;11:261–262. doi: 10.1038/nm1200. [DOI] [PubMed] [Google Scholar]
  • 8.Fujiyama S., Amano K., Uehira K., Yoshida M., Nishiwaki Y., Nozawa Y., Jin D., Takai S., Miyazaki M., Egashira K., et al. Circ. Res. 2003;93:980–989. doi: 10.1161/01.RES.0000099245.08637.CE. [DOI] [PubMed] [Google Scholar]
  • 9.Rehman J., Li J., Orschell C. M., March K. L. Circulation. 2003;107:1164–1169. doi: 10.1161/01.cir.0000058702.69484.a0. [DOI] [PubMed] [Google Scholar]
  • 10.Zhang R., Yang H., Li M., Yao Q., Chen C. Exp. Hematol. 2005;33:1554–1563. doi: 10.1016/j.exphem.2005.08.011. [DOI] [PubMed] [Google Scholar]
  • 11.Elsheikh E., Uzunel M., He Z., Holgersson J., Nowak G., Sumitran-Holgersson S. Blood. 2005;106:2347–2355. doi: 10.1182/blood-2005-04-1407. [DOI] [PubMed] [Google Scholar]
  • 12.Alvarez-Dolado M., Pardal R., Garcia-Verdugo J. M., Fike J. R., Lee H. O., Pfeffer K., Lois C., Morrison S. J., Alvarez-Buylla A. Nature. 2003;425:968–973. doi: 10.1038/nature02069. [DOI] [PubMed] [Google Scholar]
  • 13.Wang X., Willenbring H., Akkari Y., Torimaru Y., Foster M., Al Dhalimy M., Lagasse E., Finegold M., Olson S., Grompe M. Nature. 2003;422:897–901. doi: 10.1038/nature01531. [DOI] [PubMed] [Google Scholar]
  • 14.Bailey A. S., Jiang S., Afentoulis M., Baumann C. I., Schroeder D. A., Olson S. B., Wong M. H., Fleming W. H. Blood. 2004;103:13–19. doi: 10.1182/blood-2003-05-1684. [DOI] [PubMed] [Google Scholar]
  • 15.Grant M. B., May W. S., Caballero S., Brown G. A., Guthrie S. M., Mames R. N., Byrne B. J., Vaught T., Spoerri P. E., Peck A. B., et al. Nat. Med. 2002;8:607–612. doi: 10.1038/nm0602-607. [DOI] [PubMed] [Google Scholar]
  • 16.Pelosi E., Valtieri M., Coppola S., Botta R., Gabbianelli M., Lulli V., Marziali G., Masella B., Muller R., Sgadari C., et al. Blood. 2002;100:3203–3208. doi: 10.1182/blood-2002-05-1511. [DOI] [PubMed] [Google Scholar]
  • 17.Larrivee B., Niessen K., Pollet I., Corbel S. Y., Long M., Rossi F. M., Olive P. L., Karsan A. J. Immunol. 2005;175:2890–2899. doi: 10.4049/jimmunol.175.5.2890. [DOI] [PubMed] [Google Scholar]
  • 18.Bailey A. S., Fleming W. H. Exp. Hematol. 2003;31:987–993. doi: 10.1016/j.exphem.2003.07.001. [DOI] [PubMed] [Google Scholar]
  • 19.Anderson D. A., Wu Y., Jiang S., Zhang X., Streeter P. R., Spangrude G. J., Archer D. R., Fleming W. H. Stem Cells. 2005;23:638–643. doi: 10.1634/stemcells.2004-0325. [DOI] [PubMed] [Google Scholar]
  • 20.Soriano P. Nat. Genet. 1999;21:70–71. doi: 10.1038/5007. [DOI] [PubMed] [Google Scholar]
  • 21.Okabe M., Ikawa M., Kominami K., Nakanishi T., Nishimune Y. FEBS Lett. 1997;407:313–319. doi: 10.1016/s0014-5793(97)00313-x. [DOI] [PubMed] [Google Scholar]
  • 22.Akashi K., Traver D., Miyamoto T., Weissman I. L. Nature. 2000;404:193–197. doi: 10.1038/35004599. [DOI] [PubMed] [Google Scholar]
  • 23.Willenbring H., Bailey A. S., Foster M., Akkari Y., Dorrell C., Olson S., Finegold M., Fleming W. H., Grompe M. Nat. Med. 2004;10:744–748. doi: 10.1038/nm1062. [DOI] [PubMed] [Google Scholar]
  • 24.So C. W., Karsunky H., Wong P., Weissman I. L., Cleary M. L. Blood. 2004;103:3192–3199. doi: 10.1182/blood-2003-10-3722. [DOI] [PubMed] [Google Scholar]
  • 25.Vassilopoulos G., Wang P. R., Russell D. W. Nature. 2003;422:901–904. doi: 10.1038/nature01539. [DOI] [PubMed] [Google Scholar]
  • 26.Koni P. A., Joshi S. K., Temann U. A., Olson D., Burkly L., Flavell R. A. J. Exp. Med. 2001;193:741–754. doi: 10.1084/jem.193.6.741. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Yano M., Iwama A., Nishio H., Suda J., Takada G., Suda T. Blood. 1997;89:4317–4326. [PubMed] [Google Scholar]
  • 28.De Palma M., Venneri M. A., Galli R., Sergi L. S., Politi L. S., Sampaolesi M., Naldini L. Cancer Cell. 2005;8:211–226. doi: 10.1016/j.ccr.2005.08.002. [DOI] [PubMed] [Google Scholar]
  • 29.Rizvi A. Z., Swain J. R., Davies P. S., Bailey A. S., Decker A. D., Willenbring H., Grompe M., Fleming W. H., Wong M. H. Proc. Natl. Acad. Sci. USA. 2006;103:6321–6325. doi: 10.1073/pnas.0508593103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Paris F., Fuks Z., Kang A., Capodieci P., Juan G., Ehleiter D., Haimovitz-Friedman A., Cordon-Cardo C., Kolesnick R. Science. 2001;293:293–297. doi: 10.1126/science.1060191. [DOI] [PubMed] [Google Scholar]
  • 31.Wagers A. J., Sherwood R. I., Christensen J. L., Weissman I. L. Science. 2002;297:2256–2259. doi: 10.1126/science.1074807. [DOI] [PubMed] [Google Scholar]
  • 32.Abkowitz J. L., Robinson A. E., Kale S., Long M. W., Chen J. Blood. 2003;102:1249–1253. doi: 10.1182/blood-2003-01-0318. [DOI] [PubMed] [Google Scholar]
  • 33.Cursiefen C., Chen L., Borges L. P., Jackson D., Cao J., Radziejewski C., D’Amore P. A., Dana M. R., Wiegand S. J., Streilein J. W. J. Clin. Invest. 2004;113:1040–1050. doi: 10.1172/JCI20465. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Grunewald M., Avraham I., Dor Y., Bachar-Lustig E., Itin A., Yung S., Chimenti S., Landsman L., Abramovitch R., Keshet E. Cell. 2006;124:175–189. doi: 10.1016/j.cell.2005.10.036. [DOI] [PubMed] [Google Scholar]
  • 35.Jin D. K., Shido K., Kopp H. G., Petit I., Shmelkov S. V., Young L. M., Hooper A. T., Amano H., Avecilla S. T., Heissig B., et al. Nat. Med. 2006;12:557–567. doi: 10.1038/nm1400. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supporting Figures

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences

RESOURCES