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Am J Physiol Renal Physiol 295: F1017-F1022, 2008. First published July 23, 2008; doi:10.1152/ajprenal.90218.2008
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Erythropoietin expands a stromal cell population that can mediate renoprotection

Baoyuan Bi, Jiankan Guo, Arnaud Marlier, Shin Ru Lin, and Lloyd G. Cantley

Section of Nephrology, Yale University School of Medicine, New Haven, Connecticut

Submitted 27 March 2008 ; accepted in final form 21 July 2008


   ABSTRACT
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 ABSTRACT
MATERIALS AND METHODS
 RESULTS AND DISCUSSION
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 REFERENCES
 
Recent studies have demonstrated that erythropoietin (EPO) receptors are expressed on tubular epithelial cells and that EPO can protect tubular cells from injury in vitro and in vivo. Separate studies have demonstrated that marrow stromal cells (MSCs) exert a renoprotective effect in ischemia-reperfusion and cisplatin tubular injury via the secretion of factors that reduce apoptosis and increase proliferation of tubular epithelial cells. In the present study we demonstrate that MSCs express EPO receptors and that EPO can protect MSCs from serum deprivation-induced cell death and can stimulate MSC proliferation in vitro. The administration of EPO to mice resulted in the expansion of CD45-Flk1-CD105+ MSCs in the bone marrow and in the spleen and mobilized these cells as well as CD45-Flk1+ endothelial progenitor cells into the peripheral circulation. Consistent with previous reports, the administration of EPO diminished the decline in renal function associated with cisplatin administration. This effect was partially reproduced by intraperitoneal injection of cultured EPO-mobilized cells in cisplatin-treated mice. Thus the in vivo expansion and/or activation of these cells may contribute to the renoprotective effects of EPO to protect tubular cells from toxic injury.

renal tubular cell; erythropoietin receptor; cisplatin; mesenchymal stem cell

ERYTHROPOIETIN (EPO) is a glycoprotein hormone that is well known for its ability to stimulate the formation and differentiation of erythroid precursor cells in the bone marrow. However, it has become increasingly clear that other cell types express the EPO receptor (EpoR) and can respond to EPO treatment. Recent observations have demonstrated that EpoRs are expressed on cells outside of the bone including neurons, endothelial cells, cardiomyocytes, and renal tubular cells (21, 34, 36), expanding the potential biological role of EPO beyond erythropoiesis. Interestingly, animal studies have suggested that EPO can act on these cells to protect organs such as the brain, heart, and kidney against acute injury (12, 17, 25, 29, 35–37). The majority of these experiments support a model in which EPO can activate signaling pathways to prevent apoptosis and/or stimulate reparative proliferation of the injured cells (Refs. 9, 19, 34; reviewed in Ref. 3).

Recently, bone marrow-derived cells other than erythroblasts, including human marrow stromal cells (MSCs) and endothelial progenitor cells (EPCs), have also been shown to express the EpoR and/or proliferate in response to EPO treatment (26, 27, 38, 39). MSCs represent a mixed cell population that were originally identified by their ability to adhere to plastic culture dishes and to survive in primary culture for many passages (8). A similar population of stromal cells has been isolated from adipose tissue, termed adipose-derived stromal cells (ADSCs; Ref. 18). Because some MSCs can be induced to differentiate to become adipocytes, chondrocytes, or osteoclasts, they have also been named mesenchymal stem cells, and a series of surface marker proteins, including CD105, CD90, and Stro-1, have been used to isolate and better characterize these cells (6, 16). Multiple studies have demonstrated that injury of heart, nerves, or kidney by either tissue ischemia-reperfusion or toxin exposure can be prevented or diminished by the infusion of MSCs that have been expanded in vitro and reinfused in large numbers (1, 4, 11, 14, 20, 31, 33). Although some studies have suggested that infused MSCs provide this protection by differentiation into functional cells of the injured organ, the majority of studies support a differentiation-independent paracrine and/or autocrine effect of the infused cells (Refs. 4, 11, 14, 31; reviewed in Ref. 7). Therefore, the identification of factors that can act to expand this cell population in vivo and thereby increase the tissue-protective effect is of significant therapeutic interest.

In this report, we demonstrate that mouse MSCs express EpoRs, and that EPO can prevent apoptotic cell death of MSCs in culture and increase the numbers of MSCs present in both bone marrow and spleen after in vivo administration. EPO treatment also results in the mobilization of a small number of stromal cells into the circulation (erythropoietin-mobilized stromal cells, EmSCs). Cultured EmSCs are shown to protect against cisplatin-induced acute tubular injury when reintroduced into the mouse.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS AND DISCUSSION
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In vivo administration of erythropoietin and cisplatin. Unless otherwise noted, in vivo studies were performed with female 8- to 12-wk-old C57BL/6J mice weighing 18–20 g. Mice were maintained on a standard diet, and water was freely available. EPO (1,000 IU/kg sc, Amgen) or saline (vehicle control) was administered daily for up to 3 days, followed by bone marrow isolation, peripheral blood culture, or cisplatin-induced acute tubular injury. Cisplatin (10 mg·kg–1·day–1) was dissolved in saline at a concentration of 1 mg/ml and administered intraperitoneally on two successive days to induce acute tubular injury as previously described (4). Renal function was monitored by serum blood urea nitrogen (BUN) with the COBAS Integra system (Roche). After 6 days animals were killed, and kidneys were perfused with saline, fixed in formaldehyde, sectioned, and stained with hematoxylin and eosin. All animal protocols were approved by the Institutional Animal Care and Use Committee of Yale University.

Stromal cell isolation and generation of conditioned medium. MSCs were isolated from mouse femurs and cultured as described previously (4). Isolation of EmSCs was performed after 3 days of EPO treatment as described above. Whole blood was collected on day 4, followed by red blood cell lysis (BD Pharmingen) and plating of remaining cells in Iscove's modified Dulbecco's medium (IMDM) basal medium + 10% fetal calf serum (FCS). Nonadherent cells were removed by a medium change at 48–72 h and every 4 days thereafter. Cells were passaged before reaching confluence (typically once/3–4 wk) and were used after passage 2 (8–10 wk).

For in vivo experiments, 1 x 105 EmSCs suspended in 200 µl of IMDM were injected intraperitoneally 4 h before the second dose of cisplatin. Both control and EmSC-treated animals received 1 ml of saline intraperitoneally daily from day 0 to day 5. For experiments using conditioned medium rather than cells, EmSC-conditioned medium (EmSC-CM) was generated by culturing 1 x 106 stromal cells at passage 2 in serum-free IMDM for 3 days, the resultant conditioned medium was centrifuged to remove cell debris, and the supernatant was used immediately. Mice were given either 1 ml of EmSC-CM or fresh IMDM intraperitoneally daily for 5 days (days 0–4) and 1 ml of saline on day 5. The control group receiving cisplatin with 1 ml of fresh IMDM daily was indistinguishable from the control group receiving cisplatin with 1 ml of saline daily, and these groups were combined for statistical analysis. The experiment was repeated on four separate occasions.

Fluorescence-activated cell sorter analysis. For phenotypic analysis of freshly isolated mononuclear cells, 0.1 ml of peripheral blood from mice treated with EPO or saline was utilized after red blood cell lysis. Lineage-negative cells were identified by incubating 1 x 106 cells from each animal [suspended in 1 ml of HBSS (Invitrogen) containing 10% FCS] in the presence of biotinylated monoclonal anti-mouse lineage antibodies (CD3, B220, CD11b, Gr-1, and Ter119; BD Pharmingen, San Diego, CA) followed by avidin-conjugated phycoerythrin (PE; BD Pharmingen). For analysis of stromal cell populations, samples of whole blood, whole bone marrow [isolated as described previously (4)], or splenic cells [isolated by the method of Patschan et al. (22)] were subjected to red blood cell lysis followed by labeling with anti-mouse CD105-fluorescein isothiocyanate (FITC), Flk-1-PE, and CD45-APC antibodies (BD Pharmingen). After washing with HBSS, propidium iodide (PI, 2 µg/ml; Sigma) was added to label dead cells, followed by fluorescence-activated cell sorter (FACS) analysis with a FACSCalibur (BD Pharmingen). At least 100,000 events were analyzed per sample, all staining was normalized to isotype-matched control antibodies (BD Pharmingen), and data were analyzed with CellQuest software (Becton Dickinson).

For phenotypic analysis of in vitro cultured bone marrow or peripheral blood stromal cells, cells were detached with trypsin-ethylenediaminetetraacetic acid (EDTA) for 5 min, immediately washed with phosphate-buffered saline (PBS) to remove trypsin, and resuspended at 106/ml and analyzed as described above. No difference in surface marker expression (including MSC-related and hematopoietic molecules) was observed by gently detaching the cells with a cell scraper and then mixing the cell suspension with a syringe to disaggregate cell clusters.

Stromal cell survival and proliferation assays. For proliferation assays, MSCs were seeded onto six-well plates (1 x 105 cells/well) in IMDM + 10% serum and allowed to attach for 24 h at 37°C, and then the medium was replaced with IMDM ± EPO (10 IU/ml) in the absence of serum. Cells were incubated for 24 h, followed by addition of bromodeoxyuridine (BrdU, 10 µM) and quantification of BrdU-positive cells at 48 h performed via immunocytochemistry as previously described (4). For analysis of cell survival, MSCs were treated with or without EPO in 0.2% serum for 6 days, followed by determination of the number of surviving adherent cells by washing to remove dead cells and counting the number of remaining adherent cells/high-power field in 10 randomly selected fields/well. To determine the number of dead cells, all cells were collected after 6 days (both the detached cells in the culture supernatant and the adherent cells), resuspended in 400 µl of saline containing PI, and immediately analyzed by flow cytometry to identify PI-positive (dead) cells.

Analysis of EPO receptor expression. For protein analysis, 40 µg of total protein obtained from MSC lysates was resolved on a 12% SDS-polyacrylamide gel, transferred to a nitrocellulose membrane (Amersham Biosciences), and blocked with 5% nonfat dry milk in PBS containing 0.1% Tween 20 for 1 h at room temperature. Lysates from K562 cells (an erythroleukemia cell line) were used as a positive control for EpoR expression. The EpoR was detected with anti-EpoR (1:100, Santa Cruz Biotechnology) followed by incubation with rabbit IgG conjugated to horseradish peroxidase (1:3,000; Amersham Biosciences) for 1 h at room temperature. The anti-EpoR antibody detects either a single band at ~64 kDa or a doublet depending on the cell type (5). Antigen-antibody complexes were visualized with enhanced chemiluminescence (ECL, Amersham). For PCR analysis, total RNA was isolated from either MSCs or mouse proximal tubule cells (MPTs, Ref. 30) as previously described (28). Two-step quantitative real-time RT-PCR was performed with power SYBR Green mix (Applied Biosystems) with a 7300 AB Real-Time PCR machine (Applied Biosystems). EpoR expression levels were determined by the comparative threshold cycle ({Delta}Ct) method, with hypoxanthine guanine phosphoribosyl transferase 1 (Hprt1) as the reference gene. Primers used were EpoR: GGACACCTACTTGGTATTGG (forward) and GACGTTGTAGGCTGG (reverse) and Hprt1: CAGTACAGCCCCAAAATGGT (forward) and CAAGGGCATATCCA (reverse).

Adipocyte differentiation. MSCs and EmSCs were seeded at a concentration of 2.5 x 104/cm2 in a six-well plate. Differentiation medium containing 10 µM insulin (Sigma), 0.5 mM isobutylmethylxanthine (Aldrich), 1 µM dexamethasone (Sigma), and 200 µM indomethacin (Sigma) was added 24 h after seeding as per the method of Rebelatto et al. (24). The cells were grown for 3 wk, with medium replacement twice a week. Adipocyte differentiation (accumulation of lipid droplets) was detected by oil red O staining.

Statistical analysis. All results are expressed as means ± SE. Comparisons between groups were analyzed with Microsoft Excel by t-test (2-sided) or ANOVA where appropriate.


   RESULTS AND DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
GRANTS
 REFERENCES
 
Erythropoietin pretreatment protects against cisplatin-induced kidney injury. To determine whether injection of EPO affords protection against acute kidney injury resulting from toxic exposure to cisplatin, mice were injected with EPO or vehicle intraperitoneally for 3 days, followed by cisplatin injection (Fig. 1A). Three days after the first injection of cisplatin, control animals exhibited a fourfold elevation in BUN (94.3 ± 19.7 mg/dl), whereas those pretreated with EPO exhibited a less than twofold increase (32 ± 4.9 mg/dl) (Fig. 1B), similar to the effects reported by Bagnis and coworkers (2). Examination of renal histology 3 days after cisplatin treatment demonstrated severe tubular damage in the vehicle-treated animals with multiple tubules containing obstructing casts (Fig. 1C, arrows). In EPO-pretreated animals, tubular damage was also present; however, overt cell loss and cast formation were substantially reduced compared with the vehicle-treated group.


Figure 1
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  		Fig. 1. Erythropoietin (EPO) protects against acute kidney injury. A: mice received EPO (1,000 IU/kg sc) or saline vehicle on 3 successive days, followed by cisplatin injection (CIS; 10 mg/kg ip on 2 successive days) to induce acute injury (experiment repeated twice with a total of 8 mice in each group). B: blood urea nitrogen (BUN) values 24 and 72 h after the first cisplatin injection reveal a significant attenuation of the BUN rise following cisplatin injection in mice receiving EPO pretreatment; n = 8. *P < 0.01 vs. control. C: renal histology on day 6 after cisplatin injection reveals less tubular injury and cast formation in EPO-treated animals compared with controls (arrows).

 
MSCs express EPO receptors and proliferate in response to EPO. Since MSCs have been shown to protect against acute kidney injury when injected in large numbers, we examined the possibility that EPO-mediated renoprotection might result in part from increased MSC numbers or mobilization in the mouse. Examination of cultured mouse MSCs demonstrated the presence of the EpoR by both PCR and Western blot analysis (Fig. 2, A and B). To determine whether EPO can act directly on MSCs to prevent cell death and/or to increase cell proliferation, MSCs were cultured in serum-free medium in the presence of EPO or vehicle control (PBS) for 48 h. Determination of BrdU uptake revealed a modest but significant increase in the number of dividing cells after EPO treatment (Fig. 2C). Prolonged periods of serum deprivation led to detachment and death of a large percentage of MSCs in the vehicle-treated group, whereas EPO treatment diminished cell death (Fig. 2D, quantified in Fig. 2E) and resulted in a greater number of surviving, adherent cells (Fig. 2F, quantified in Fig. 2G). Thus EPO can increase MSC cell numbers by directly inhibiting cell death and stimulating proliferation.


Figure 2
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  		Fig. 2. EPO stimulates stromal cell expansion. A: PCR of mRNA from marrow stromal cells (MSCs) or cultured mouse proximal tubule cells (MPTs) using primers specific for the EPO receptor (EpoR) with control primers for Hprt. Comparative threshold cycle ({Delta}Ct) = 0.0031 for MSC and 0.0003 for MPT. B: immunoblotting of lysates from MSCs and K562 cells with anti-EpoR antibody reveals the presence of the EpoR in both cell types. C: quantitation of bromodeoxyuridine (BrdU) uptake by MSCs cultured ±EPO for 48 h; n = 3. *P < 0.01. D: fluorescence-activated cell sorter (FACS) analysis of propidium iodide (PI) uptake by MSCs following 6 days of culture in 0.2% serum ±EPO. E: quantitation of 5 experiments as shown in D. *P < 0.01. F: representative photograph of surviving adherent MSCs 6 days after culture in 0.2% serum ±EPO. G: quantitation of 5 separate experiments as shown in F. *P < 0.01. hpf, High-power field.

 
Erythropoietin increases the number of stromal cells in the marrow and circulation. To determine whether EPO treatment can increase MSC cell numbers in vivo, nucleated cells were harvested from spleen and bone marrow before and after 3 days of EPO treatment and analyzed for the presence of CD45 (a leukocyte marker), Flk1 (an endothelial cell marker), and endoglin (CD105). CD105 is a component of the transforming growth factor-β receptor signaling complex and is expressed on several cell types including hematopoietic stem cells, vascular progenitor cells, and MSCs (10, 13, 23). The presence of CD105 on adherent marrow-derived cells that lack expression of CD45 and Flk1 is commonly used for identification of MSCs (6, 13, 16). At baseline, CD45-Flk1-CD105+ cells made up <1% of nucleated bone marrow and spleen-derived cells. After treatment with EPO, the number of these cells significantly increased in both bone marrow and spleen (Fig. 3A).


Figure 3
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  		Fig. 3. EPO expands and mobilizes stromal cells in vivo. A: quantitation of FACS analysis of CD45-Flk1-CD105+ cells freshly isolated from bone marrow and spleen after 3 days of treatment with either saline or EPO. n = 5 mice/group. *P < 0.05 vs. PBS control. B: mice were treated with EPO or vehicle, followed by FACS analysis of peripheral blood for the presence of lineage-negative cells; n = 19 mice/group. *P < 0.05. C: cells in peripheral blood from mice treated with PBS or EPO were analyzed for the presence of CD45 and Flk1. CD45-Flk1– cells are at bottom left. D: CD45– cells were analyzed for expression of Flk1 and CD105. CD45-Flk1-CD105+ cells are at top left. E: quantitation of CD45-Flk1-CD105+ cells from 3 independent experiments as shown in C and D. Values are expressed as % of all nucleated cells present in peripheral blood. *P < 0.05 vs. PBS control. F: quantitation of adherent peripheral blood stromal cells derived from PBS- or EPO-treated animals after 14 days in culture; n = 10. *P < 0.05. G: representative image of oil red O staining of MSCs and EmSCs after 3 wk of culture under adipocyte-differentiation conditions showing extensive collection of fat droplets in MSCs but not EmSCs (x900 magnification).

 
To determine whether EPO treatment mobilized nonerythroid cells into the circulation, mice were injected with EPO for 3 days, followed 24 h later by FACS analysis of peripheral blood to identify circulating lineage-negative cells (nucleated cells lacking surface marker expression of erythroid-, monocyte-, leukocyte- or lymphocyte-specific proteins). At baseline, ~1% of circulating nucleated cells were lineage negative, whereas EPO treatment increased this to ~2% (Fig. 3B). Examination of surface marker expression revealed that EPO treatment increased the numbers of both CD45-Flk1+ cells (potential endothelial progenitor cells) and CD45-Flk1-CD105+ cells (mobilized stromal cells; Fig. 3, C and D, quantified in Fig. 3E).

In vitro culture of whole blood obtained from mice treated with EPO revealed an increased number of adherent, proliferating fibroblast-like cells compared with control mice (Fig. 3F). These cells continued to proliferate for >60 days in vitro, and FACS analysis at 45 days of culture revealed that their expression profile was similar to that of bone marrow-derived MSCs as well as the cells present in the circulation of control mice (Table 1). The EPO-mobilized cells did not express the EPC marker Flk1 or the hematopoietic stem cell markers CD34 or c-Kit but did express higher levels of CD45 than is typical of bone marrow MSCs.


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  		Table 1. EmSCs express markers typical of bone marrow-derived MSCs

 
To determine whether EPO-mobilized cells exhibit the stem cell characteristics typical of some MSCs, the cells were maintained for 3 wk under conditions designed to induce adipocyte differentiation. Whereas a high percentage of bone marrow-derived MSCs acquired adipocyte characteristics under these conditions, EPO-mobilized cells failed to differentiate (Fig. 3G). This result suggests either that mobilization induces changes in MSCs that prevent subsequent differentiation or that EPO-mobilized cells represent a specific subset of MSCs that lack stem cell properties. We have named this distinct population of cells EPO-mobilized stromal cells (EmSCs).

EmSCs protect against cisplatin-induced kidney injury. Previous studies suggested that the protective effect of MSCs in the injured kidney occurs primarily via an endocrine or paracrine effect rather than transdifferentiation or fusion. Therefore, we examined the possibility that EmSCs may provide a renoprotective effect even though they appear to be stromal rather than stem cells. Mice were treated with cisplatin with or without administration of either a single injection of EmSC or daily injections of EmSC-CM. Control mice received the cisplatin with an equal volume of either saline (control for EmSC injection) or fresh IMDM (control for EmSC-CM injection). Renal function (as judged by BUN values) was improved in mice receiving EmSC compared with control mice (Fig. 4A), with a similar trend in those receiving EmSC-CM, although this did not reach statistical significance. Consistent with the functional effects of EmSC injection, renal histology 6 days after injury revealed less residual tubular injury and greater preservation of tubular architecture in the EmSC- and EmSC-CM-treated animals (Fig. 4B).


Figure 4
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  		Fig. 4. EmSCs can partially protect mice from cisplatin-induced acute kidney injury. Mice were injected with cisplatin and divided into 3 groups: group 1: control, receiving 1 ml of either saline (S, n = 10) or control Iscove's modified Dulbecco's medium (M, n = 10) daily; group 2: EmSC treatment, receiving 1 x 105 EmSCs in 1 dose as well as 1 ml of saline daily (n = 15); and group 3: EmSC-conditioned medium (EmSC-CM) treatment, receiving 1 ml of EmSC-CM daily (n = 10). A: BUN values before cisplatin treatment and 3 and 6 days after cisplatin treatment. P < 0.01 for EmSC vs. control; P = 0.06 for EmSC-CM vs. control. B: histology of the renal cortex 6 days after cisplatin treatment reveals marked tubular injury in control mice, with less injury in EmSC-treated and EmSC-CM-treated mice (magnification x400).

 
Conclusions. Studies by several groups have now demonstrated that MSCs can be isolated and expanded in vitro and then injected in large numbers to induce a renoprotective effect (4, 11, 15, 20, 31, 32). However, to our knowledge there have been no studies demonstrating the in vivo activation or expansion of these cells. In the present study we demonstrate that MSCs express the EpoR and are increased in numbers in both marrow and spleen as well as mobilized into the circulation by EPO. The EPO-mobilized cells fail to differentiate in vitro yet retain renoprotective effects when injected into mice subjected to cisplatin-induced injury. These studies leave unanswered the question of whether it is primarily the increase in MSC numbers seen after EPO treatment or the mobilization of some of these cells into the circulation that provides the protective effect. Furthermore, the ability of EPO to increase the number of circulating CD45-Flk1+ cells suggests that the protective effects of EPO might also include accelerated vascular repair following kidney injury. Combined with the previously described direct effects of EPO on renal tubular cells, these findings provide further support for randomized trials to determine whether early EPO treatment can improve the outcome of patients who develop acute kidney injury.


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This work was supported by an R01 award from the National Institute of Diabetes and Digestive and Kidney Diseases (DK-66216) and an Established Investigator Award from the American Heart Association to L. G. Cantley.


   FOOTNOTES
 

Address for reprint requests and other correspondence: L. G. Cantley, Yale Univ. School of Medicine, 333 Cedar St., PO Box 208029, New Haven, CT 06510 (e-mail: lloyd.cantley{at}yale.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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