Research ArticleBone Marrow Transplantation

Diabetes Impairs Hematopoietic Stem Cell Mobilization by Altering Niche Function

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Science Translational Medicine  12 Oct 2011:
Vol. 3, Issue 104, pp. 104ra101
DOI: 10.1126/scitranslmed.3002191

Abstract

Success with transplantation of autologous hematopoietic stem and progenitor cells (HSPCs) in patients depends on adequate collection of these cells after mobilization from the bone marrow niche by the cytokine granulocyte colony-stimulating factor (G-CSF). However, some patients fail to achieve sufficient HSPC mobilization. Retrospective analysis of bone marrow transplant patient records revealed that diabetes correlated with poor mobilization of CD34+ HSPCs. In mouse models of type 1 and type 2 diabetes (streptozotocin-induced and db/db mice, respectively), we found impaired egress of murine HSPCs from the bone marrow after G-CSF treatment. Furthermore, HSPCs were aberrantly localized in the marrow niche of the diabetic mice, and abnormalities in the number and function of sympathetic nerve termini were associated with this mislocalization. Aberrant responses to β-adrenergic stimulation of the bone marrow included an inability of marrow mesenchymal stem cells expressing the marker nestin to down-modulate the chemokine CXCL12 in response to G-CSF treatment (mesenchymal stem cells are reported to be critical for HSPC mobilization). The HSPC mobilization defect was rescued by direct pharmacological inhibition of the interaction of CXCL12 with its receptor CXCR4 using the drug AMD3100. These data suggest that there are diabetes-induced changes in bone marrow physiology and microanatomy and point to a potential intervention to overcome poor HSPC mobilization in diabetic patients.

Introduction

Transplantation of hematopoietic stem and progenitor cells (HSPCs) is a well-established therapy for treating benign and malignant blood diseases (1, 2). Its clinical applications have steadily expanded to include certain solid tumors (3) and autoimmune disorders (4). The success of HSPC transplantation greatly depends on a sufficient yield of HSPCs harvested from the blood after treatment with granulocyte colony-stimulating factor (G-CSF). G-CSF is the mobilizing agent most widely used in clinics and elicits HSPC mobilization through several putative mechanisms (5). These include proteolytic cleavage and down-regulation of adhesion molecules and chemokines, the latter mediated at least in part by the sympathetic nervous system (SNS) (6, 7). Among other effects, G-CSF increases activity of the SNS in the bone marrow microenvironment, where activation of β-adrenergic receptors in stromal cells inhibits synthesis of the chemokine CXCL12 (6, 7). The resulting alteration in the CXCL12 gradient enables hematopoietic stem cells (HSCs) to egress from the bone marrow (5, 8). A significant fraction of transplant-eligible patients, referred to as “poor mobilizers,” do not achieve adequate numbers of HSCs in the peripheral blood (PB) despite appropriate cytokine stimulation (9, 10). With the exception of drug-induced myelotoxicity, it is largely unknown why these patients fail to mobilize their HSCs. Reviewing a caseload of mobilization procedures performed over a 4-year period, we observed elevated serum glucose concentrations and lower CD34+ HSC numbers in “mobilized” blood in patients with diabetes. Using two distinct mouse models of diabetes, we found that diabetes impairs HSC mobilization by altering perivascular neural and mesenchymal cell function and CXCL12 distribution in the bone marrow.

Results

Patients with diabetes mellitus show poor HSC mobilization

Autologous PB stem cell transplantation procedures performed in the University of Parma Bone Marrow Transplantation Unit between 2004 and 2008 were reviewed (table S1). We found an overall 22.6% (14 of 62) prevalence of mobilization failure, defined as <20 CD34+ and <3000 polymorphonuclear cells/μl, <5000 leukocytes/μl, and <50,000 platelets/μl of blood at the beginning of leukapheresis. The frequency of diabetes was 50% (7 of 14) in poor versus 25% (12 of 48) in good mobilizers and was independent of age, gender, and cycles of previous chemotherapy (Fig. 1A). Among the 48 patients who were able to mobilize their HSCs, the amount of CD34+ cells/kg was lower in diabetic compared to nondiabetic individuals (P = 0.014; Fig. 1B). Additionally, glucose concentrations were significantly higher in poor mobilizers (n = 14 poor mobilizers, n = 48 good mobilizers, P = 0.002; Fig. 1C). There was a trend in slower neutrophil recovery (P = 0.146) and significantly reduced platelet recovery (P = 0.006) after PB stem cell transplantation of diabetic patients (n = 12; P = 0.146 and 0.006, respectively; fig. S1, A and B). When poor mobilizers were transplanted with autologous bone marrow–derived cells, the observed delayed engraftment of diabetic patients was more evident (n = 7; P = 0.002 and P = 0.003, respectively; fig. S1, C and D; all data are summarized in fig. S1E).

Fig. 1

Diabetes reduces G-CSF–induced HSPC mobilization. (A) Table shows the actual numbers (and percentages) of patients treated with G-CSF with more than (good mobilizers) or fewer than (poor mobilizers) 20 CD34+ cells/μl. There was an overall rate of mobilization failure of 22.6% (14 of 62). The frequency of diabetes is 50% (7 of 14) in poor mobilizers versus 25% (12 of 48) in good mobilizers (P = 0.102, Fisher’s exact test). (B) Number of CD34+ cells per kilogram in the peripheral blood (PB) of patients mobilized with G-CSF (10 μg/kg per day). Scatter plot showing mean ± SEM. n = 36 (nondiabetic patients) and n = 12 (diabetic patients). Diabetic patients mobilize CD34+ HSCs more poorly [n = 36 (nondiabetic), n = 12 (diabetic); *P = 0.014 Wilcoxon two-sample test] even among good mobilizers. (C) Glucose concentrations (mg/dl) in good mobilizers versus poor mobilizers. Scatter plot shows mean ± SEM. Higher glucose concentrations are found in poor mobilizers [n = 14 (poor mobilizers), n = 48 (good mobilizers); **P = 0.002, Wilcoxon two-sample test]. (D) Number of CFU-C per 50 μl of G-CSF mobilized PB (mPB), obtained from STZ-treated or control mice. Columns represent means ± SEM. n = 12. ***P < 0.001, two-tailed Student’s t test. (E) Percentages of total CD45.2+ donor-derived cells in the PB of lethally irradiated SJL recipient mice transplanted with 150 μl of G-CSF mobilized PB from C57BL/6 diabetic or control mice as assessed by FACS analysis at regular time intervals. Columns represent means ± SEM. n = 18. **P < 0.01; ***P < 0.001, two-tailed Student’s t test. (F and G) STZ-treated and control mice were divided into four categories based on PB glucose concentrations (<150, 150 to 200, 200 to 300, and >300 mg/dl). Histogram plots represent mean ± SEM of (F) number of LSK cells in 4 × 105 total blood cells by flow cytometry (n = 12; *P = 0.016) and (G) number of CFU-C per 50 μl of mobilized PB (n = 12; **P = 0.006, Jonckheere-Terpstra test).

HSPC mobilization by G-CSF is impaired in animal models of diabetes

To investigate the mechanism underlying poor HSC mobilization in diabetic patients, we evaluated HSPC mobilization in both a type I model of diabetes induced by treatment with streptozotocin (STZ) (11) and a type II model of diabetes, the leptin receptor knockout (db/db) mouse. In both settings, G-CSF injections (125 μg/kg every 12 hours for 4 days) were intraperitoneally administered to mobilize HSPCs. Flow cytometry of blood collected 3 hours after the last injection demonstrated significantly reduced percentages of LinSca-1+c-Kit+ (LSK) cells in the circulation of STZ-treated mice (control, 0.072 ± 0.016; diabetic, 0.024 ± 0.008; n = 5; P = 0.03). Mobilized PB from these animals displayed a significant decrease in circulating colony-forming units in methylcellulose culture (CFU-C) (n = 12, P < 0.001; Fig. 1D) and diminished capacity to reconstitute hematopoiesis in lethally irradiated congenic recipients (Fig. 1E). PB chimerism in recipient mice 4 to 16 weeks after transplantation was reduced by ~50% when mobilized blood was from diabetic compared to control animals (n = 18, P < 0.001 at 4 weeks, P < 0.01 at 8, 12, and 16 weeks; Fig. 1E). Similar results were observed when the same transplantation was performed with mobilized PB from db/db animals (fig. S2A). Given that insulin deficiency characterizes type 1 (STZ-induced) diabetes and hyperinsulinemia characterizes type 2 (db/db) diabetes, we sought to determine whether the mobilization defect was associated with blood glucose or insulin concentration. The STZ-treated and db/db mice were stratified into four categories based on glucose concentrations. An inverse correlation between HSPC mobilization and blood glucose in both diabetes models was found, as measured by the numbers of circulating LSK cells and CFU-C (Fig. 1, F and G, and fig. S2B). Specifically, the Jonckheere-Terpstra test was used to assess the correlation between glucose concentrations and mobilization. An exact two-sided P value was computed using StatXact 6.0 (Cytel Software Corp.). Higher blood glucose was associated with lower mobilization in both STZ-treated (n = 12, P = 0.016) and db/db (n = 12, P = 0.006) mice. Plasma insulin concentration varied depending on the mouse model and was either low/absent (0.1 ± 0.2 ng/ml) or high (30 ± 8 ng/ml) in the poorest mobilizer groups. It was within the normal range (1.1 ± 0.3 ng/ml) in the groups with lower glycemic concentrations in either model. These results suggest that the rate of reduction in HSPC mobilization caused by diabetes is proportional to glucose imbalance and is independent of insulin concentration.

Bone marrow HSC numbers are not affected in STZ-treated mice

Given the correspondence of the data between the model systems, only STZ-treated mice were subjected to more detailed analyses. We assayed whether the reduction in circulating HSPCs in STZ-treated diabetic mice was due to a reduced HSC content in steady-state bone marrow. Percentages and absolute numbers of LSK progenitor cells were increased in the bone marrow of diabetic mice (n = 24, P < 0.001; Fig. 2A), whereas total bone marrow cellularity was comparable to that of controls. This was in keeping with higher CFU-C formation per unit volume from diabetic bone marrow–derived cells (n = 6, P < 0.05; Fig. 2B). Both the percentage and absolute numbers of long-term HSCs (LT-HSCs) (LinSca-1+c-Kit+CD150+CD48 and LinSca-1+c-Kit+Flk-2CD34) were also increased in STZ-treated mice (n = 8, P < 0.01 and P < 0.05; Fig. 2, C and D, respectively). This was accompanied by higher numbers of proliferating HSPCs and LT-HSCs (n = 4, P < 0.05; Fig. 2, E and F, and fig. S2C). In competitive repopulation assays, the diabetic-derived bone marrow cells exhibited an advantage in long-term engraftment when injected into irradiated nondiabetic recipients (n = 10, P < 0.001; Fig. 2, G and H). The correspondence between the number of stem cells by flow cytometry and the repopulating ability in competitive transplantation suggests that LT-HSCs have an intact repopulating potential per stem cell. Together, these results indicate that diabetic mice have an increased number of progenitor cells and LT-HSCs within the bone marrow despite exhibiting impaired HSC mobilization after G-CSF administration.

Fig. 2

Characterization of the hematopoietic compartment in the bone marrow of STZ-treated animals. (A) Number of LSK cells per femur defined by flow cytometry in STZ-treated or control mice. Data are means ± SEM. n = 24. ***P < 0.001. (B) CFU-C numbers of bone marrow (BM) cells obtained from STZ-treated and control mice. Data are means ± SEM. n = 6. *P < 0.05. (C and D) Number per femur of LT-HSCs LinSca-1+c-Kit+CD150+CD48 (C) and LinSca-1+c-Kit+Flk-2CD34 (D) in STZ-treated or control mice as assessed by flow cytometry. Data are means ± SEM. n = 8. *P < 0.05; **P < 0.01. (E and F) Histogram plots showing the fraction of LSK cells (E) and LT-HSCs (F) in different phases of the cell cycle in STZ-treated versus control mice. Columns are means ± SEM. n = 4. *P < 0.05. (G) Schematic representation of the transplantation experiment to assess the number/functionality of bone marrow cells from STZ-induced diabetic mice. (H) Donor-derived CD45.2+ cell engraftment 4, 8, 12, and 16 weeks after transplantation of the PB of secondary SJL recipient mice transplanted with bone marrow isolated from STZ-treated or control mice and mixed with an equal amount of CD45.1 competitor cells. Columns represent means ± SEM. n = 10. ***P < 0.001. (I) CFU-C (relative to input) mean number ± SEM of the migrated purified LSK cells from STZ-treated or control mice in response to a CXCL12 chemokine gradient (300 ng/ml). n = 6. ***P < 0.001. (J) FACS-sorted LSK cells from control or STZ-treated mice were plated onto 24-well plates previously coated with fibronectin (10 μg/ml). After 4 hours, the adhesive LSK content was plated into methylcellulose. Data are mean CFU-C numbers relative to input ± SEM. n = 6. ***P < 0.001. (K) Representative FACS plots and bars indicating Δ mean fluorescence intensity (ΔMFI) over controls, showing the expression of CXCR4, α4 integrin, α5 integrin, and L-selectin in LSK cells isolated from STZ-treated and control mice. n = 9. *P < 0.05. All data were analyzed with unpaired, two-tailed Student’s t test.

Diabetes increases adhesion and decreases chemotaxis of mouse HSPCs

To determine whether the defect in HSPC mobilization in diabetic mice was due to changes in their chemotactic or adhesive properties, we subjected purified LSK cells to in vitro migration and cell attachment assays. LSK isolated from STZ-treated mice demonstrated impaired migration in response to a CXCL12 gradient and increased adhesion to fibronectin ex vivo (n = 6, P < 0.001; Fig. 2, I and J). The expression of surface antigens involved in HSPC retention and migration (CXCR4, L-selectin or CD62L, α4 integrin or CD49d, α5 integrin or CD49e) was analyzed by flow cytometry. CXCR4 expression on control LSK bone marrow cells was low with variable and nonsignificant increases in the setting of diabetes. α4 and α5 integrin expression was unchanged, whereas a significant increase in L-selectin expression was noted on diabetic LSK (n = 9, P < 0.05; Fig. 2K). Thus, some changes in HSPC migratory and adhesive activities could contribute to the observed phenotype.

HSPC numbers and mobilization revert to normal when transplanted into a nondiabetic host

To determine whether the observed changes in HSPC migration, adhesion, and antigen expression in STZ-treated mice persisted in a normal host, we administered G-CSF to mice transplanted 16 weeks earlier with CD45.2 bone marrow cells from STZ- or saline-treated mice (Fig. 3A). However, no difference in the mobilization of CFU-C into the blood was observed (Fig. 3B). Also, transplantation of irradiated animals with mobilized blood revealed that the contribution of the chimeric fractions (CD45.1/CD45.2 ratio) was equivalent in the two groups (Fig. 3, C and D). These data indicate that cell-autonomous alterations in function were not persistent and reverted in a nondiabetic host, suggesting a primary role for the bone marrow microenvironment in the diabetes-related mobilization defect. Further, they exclude the possibility that a direct, durable effect of STZ on HSPCs could account for their mobilization abnormality.

Fig. 3

Altered HSPC mobilization in diabetic mice is not cell-autonomous but is microenvironment-dependent. (A) Schematic representation of the procedure followed to assess whether the inhibition of mobilization is cell-autonomous or microenvironment-dependent. Lethally irradiated SJL recipient mice were transplanted with 1 × 106 bone marrow cells from STZ-treated or control mice, along with 1 × 106 CD45.1 support cells. Sixteen weeks after transplantation, the SJL recipient mice from the two groups underwent a G-CSF mobilization regimen. (B) Number of CFU-C in the PB of recipients transplanted with STZ-treated or control cells after the induction of mobilization with G-CSF. Columns represent means ± SEM. n = 8. (C and D) Actual numbers (C) and ratio (D) of CD45.1+ and CD45.2+ cells in the PB of recipient mice transplanted with STZ-treated or control cells before and after mobilization with G-CSF treatment. Columns represent means ± SEM. n = 9 and 8. (E) Schematic representation of the reverse experiment in which 15 STZ-treated and control C57BL/6 mice were transplanted with wild-type (WT) CD45.1 whole bone marrow cells. After 16 weeks, 10 mice were evaluated for LSK and LT-HSC numbers. Five mice underwent G-CSF mobilization therapy followed by blood collection and evaluation of CFU-C number. (F) Graph bars represent number of LSK cells per femur. Data are means ± SEM. n = 10. **P < 0.01. (G and H) Number per femur of LT-HSCs LinSca-1+c-Kit+CD150+CD48 (G) and LinSca-1+c-Kit+Flk-2CD34 (H) as assessed by flow cytometry. Data are means ± SEM. n = 10. (I) Number of CFU-C per 100 μl of G-CSF–mobilized PB from STZ-treated and control mice 16 weeks after transplantation. Data are means ± SEM. n = 5. *P < 0.05. All data were analyzed with unpaired, two-tailed Student’s t test.

To assess whether the bone marrow microenvironment was sufficient to induce the alterations in HSPC numbers and G-CSF response observed in diabetic mice, we transplanted equivalent numbers of wild-type CD45.1 bone marrow cells into lethally irradiated diabetic and control mice (Fig. 3E). Sixteen weeks after engraftment, bone marrow LSK cells were increased (n = 10, P < 0.01; Fig. 3F), with a trend to increased LT-HSCs in diabetic recipients (Fig. 3, G and H). When G-CSF was administered to these animals, the number of circulating CFU-C was lower in diabetic mice transplanted with wild-type bone marrow LSK cells (n = 5, P < 0.05; Fig. 3I). These results indicate that exposure of HSPCs to STZ was not required for their dysfunction. Rather, persistent extrinsic signals from the abnormal diabetic bone marrow microenvironment or a rapidly reversible change in HSPC function are responsible for maintaining the mobilization defect associated with diabetes.

Diabetes alters bone marrow niche cells

After transplantation into irradiated recipients, LT-HSCs can be visualized in close proximity to the endosteal surface often positioned near osteoblastic cells and sinusoidal vessels (1214). They are retained in the bone marrow by interactions with molecules such as kit ligand or stem cell factor (SCF) present on osteoblastic cells (1519), and CXCL12 expressed by a variety of cell types. Modulating CXCL12 expression is thought to be required for HSC mobilization from the bone marrow into the blood (2022).

Consequently, we examined microanatomic relationships, kit ligand expression, and CXCL12 expression in the bone marrow of diabetic mice. Mice carrying a transgene in which green fluorescent protein (GFP) expression was driven by the osteoblastic-specific Col2.3kb promoter (23) or the Nestin promoter (24) (which labels bone marrow mesenchymal stem cells) were used to identify cells that constitute the bone marrow microenvironment.

Diabetes alters LT-HSC function in the bone marrow

High-resolution confocal microscopy and two-photon video imaging were performed after injection of LT-HSCs into Nes-Gfp and Col2.3-Gfp transgenic mice. Briefly, calvarial bone marrow was visualized after the injection of 5000 dye-labeled LT-HSCs. The total number of HSCs and measurements of their distance relative to osteoblastic cells, the endosteal surface, and Nestin-GFP+ cells were quantified in diabetic and control mice. Higher numbers of LT-HSCs were observed 24 hours after injection in diabetic mice compared to their controls (n = 8, P < 0.05; Fig. 4A). Clusters of two, three, or more cells labeled with the DiD (1,1′-dioctadecil-3,3,3′-tetramethylindodicarbocyanine perchlorate) lipophilic cyanine dye were also observed at higher proportions in diabetic mice (Fig. 4B). To assess whether these events could be attributed to an increased chemoattractive ability of the diabetic marrow causing cell accumulation or to HSC proliferation, we injected a 1:1 mixture of FACS (fluorescence-activated cell sorting)–sorted LT-HSCs stained with the lipophilic cyanine dyes DiD or DiI (1,1′-dilinoleyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate). We found that cells in clusters were exclusively stained with one dye, consistent with migration and proliferation of a single cell, rather than colocalization of multiple injected HSCs (Fig. 4C). Further evaluation was conducted in vitro using DsRed-stained LSK CD48 cells cultured over a layer of stromal cells derived from control or diabetic mice. Despite several days of normoglycemic culture conditions (glucose concentration, 5.5 mM), diabetic stroma still promoted a 20% increase in HSPC growth compared to normal stroma (P < 0.01; fig. S2D). Together with the higher fraction of cycling bone marrow cells found in diabetic mice (P = 0.05; fig. S2E) and increased cell number detected by FACS and two-photon video imaging, these data indicate that the diabetic stroma fosters HSPC proliferation.

Fig. 4

Altered LT-HSC function in the bone marrow microenvironment of diabetic mice. (A) Number of WT LT-HSCs found within 24 hours in the calvaria bone marrow cavity of control or STZ-treated recipients using in vivo imaging. n = 8. *P < 0.05. (B) Percentages of one, two, or three or more cell clusters found in the bone marrow cavity of control and STZ-treated recipients. (C) Representative pictures of the bone marrow cavity of STZ-treated mice injected with equal numbers of DiD- and DiI-labeled HSCs. Blue, second-harmonic generation signal (bone); green, Col2.3GFP (osteoblasts); red, DiD-labeled cells (LT-HSCs); white, DiI-labeled cells (LT-HSCs). Scale bars, 75 μm. (D) Distances of LinSca-1+c-Kit+Flk-2CD34 and LinSca-1+c-Kit+CD150+CD48 cells 24 and 48 hours after transplantation relative to the Col2.3GFP cells and bone (measured in micrometers). Dot plots represent means ± SEM. n = 4. **P < 0.01; ***P < 0.001. (E) Percentage changes in SCF (kitl) mRNA levels in sorted Col2.3GFP+ osteoblasts in STZ-treated and control mice normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (ΔΔCT method). Columns represent means ± SEM. n = 6. *P < 0.05. (F) Percentage difference in Cxcl12 mRNA levels during steady-state hematopoiesis among nestin+ mesenchymal cells and osteoblasts. n = 6. **P < 0.01. All data were analyzed with unpaired, two-tailed Student’s t test.

The number of osteoblastic cells was significantly reduced in diabetic mice (P < 0.05), whereas the number of Nestin-GFP+ cells did not change (fig. S2, F and G). The latter result was also observed in human bone marrow biopsies, where the fraction of nestin+ cells was similar in diabetic and nondiabetic patients (fig. S3, A to D).

In addition, LT-HSCs were found in closer proximity to osteoblastic cells and to the endosteal surface in diabetic mice 24 to 48 hours after transplantation (Fig. 4D). No difference was noted in location relative to nestin-expressing perivascular cells. Assessment of mRNA levels of niche-related genes (CXCL12, VCAM1, SCF, and angiopoietin-1) in osteoblastic cells revealed that, under steady-state conditions, the expression of SCF, previously associated with HSC lodgement in the endosteal region (17), was about twofold higher in diabetic mice (n = 6, P < 0.05; Fig. 4E), consistent with the observed closer proximity of HSCs. These data suggest that the diabetic microenvironment alters the lodgement and proliferation of normal HSCs.

Diabetes alters CXCL12 distribution within the bone marrow

Previous studies have shown that G-CSF–induced HSPC mobilization requires down-regulation of CXCL12 in the bone marrow (2527). During steady-state hematopoiesis, Nestin-GFP+ cells are reportedly the major source of CXCL12 in mouse bone marrow (28). In control mice, Cxcl12 mRNA levels were >10-fold higher in Nestin-GFP+ cells compared to osteoblastic cells (n = 6, P < 0.01; Fig. 4F). Cxcl12 mRNA levels in Nestin-GFP+ cells were reduced by about twofold in diabetic animals compared with controls (n = 6, P < 0.05; Fig. 5A). Exposure to G-CSF had minimal impact on Cxcl12 mRNA levels in Nestin-GFP+ cells in the setting of diabetes, whereas Cxcl12 mRNA levels were efficiently down-regulated in control mice (n = 6, P < 0.01; Fig. 5A). Thus, the magnitude of the Cxcl12 mRNA change after G-CSF administration was markedly lower in diabetic animals compared to controls (92% versus 38%). This was the consequence of both lower baseline Cxcl12 levels and failure of G-CSF to induce Cxcl12 mRNA down-regulation. The lower mRNA levels in diabetic mice were accompanied by a decrease in CXCL12 protein in Nestin-GFP+ cells by flow cytometry (n = 6, P < 0.05; Fig. 5B). Despite these changes in cell type–specific mRNA levels, CXCL12 protein content in bone marrow extracellular fluid did not change (Fig. 5C) and PB actually had higher CXCL12 protein concentrations in diabetic mice (P < 0.05, fig. S2I). Together, these results suggest that CXCL12 expression dynamics are changed in the diabetic microenvironment in a cell-specific manner.

Fig. 5

STZ-induced diabetic mice show aberrant expression of niche-related molecules. (A) Percent changes in the expression of mRNA levels of Cxcl12 in FACS-sorted Nestin-GFP + cells before and after G-CSF treatment relative to control Nestin-GFP+ cells before G-CSF treatment and normalized to GAPDH (ΔΔCT method). Columns represent means ± SEM. n = 6. *P < 0.05; **P < 0.01. (B) Representative FACS plots and graph bars showing percentages of CXCL12+ cells among nestin+ cells. Data are means ± SEM. n = 6. *P < 0.05. (C) Concentration (pg/ml) of CXCL12 in bone marrow extracellular fluid from control and STZ-treated mice. Data are mean numbers ± SEM. n = 6. (D) Schematic representation of the experimental design used to selectively disrupt osteoblastic cells (see Materials and Methods). (E) Donor-derived CD45.2+ cell engraftment 4 weeks after transplantation of mobilized PB in lethally irradiated SJL recipient mice. Columns represent means ± SEM. n = 8 in each group. P < 0.05; **, ••, ▿▿, ##P < 0.01; ♦♦♦, □□□, ★★★P < 0.001. C-PBS, nondiabetic mice, PBS-treated; C–G-CSF, nondiabetic mice, G-CSF–treated; STZ-PBS, diabetic mice, PBS-treated; STZ–G-CSF, diabetic mice, G-CSF–treated; OcΔ/Δ-PBS, nondiabetic, osteoblast-deficient mice, PBS-treated; OcΔ/Δ–G-CSF, nondiabetic, osteoblast-deficient mice, G-CSF–treated; OcΔ/Δ-STZ PBS, diabetic, osteoblast-deficient mice, PBS-treated; OcΔ/Δ-STZ G-CSF, diabetic, osteoblast-deficient mice, G-CSF–treated. All data were analyzed with unpaired, two-tailed Student’s t test.

Cell depletion defines the role of osteoblasts in mobilization

To clarify the relative role of osteoblasts in the mobilization defect in diabetic settings, we used the diphtheria toxin (DT) system to selectively ablate these cells from STZ-treated and control mice. Briefly, mice in which expression of Cre recombinase was driven by the osteoblast-specific osteocalcin promoter (29) were crossed with mice harboring an inducible DT receptor transgene (iDTR) (30). In the double-transgenic mice, Cre-mediated removal of a transcriptional STOP cassette allows the expression of the DT receptor. Upon DT administration, efficient ablation of the target population was achieved. Mice with only one transgene were used as controls. Diabetes was induced in both double-transgenic (Oc-CRE–iDTR) and control (Oc-CRE or iDTR) mice. After depletion of osteoblasts, mice were treated with G-CSF or saline. Mobilized PB was transplanted in lethally irradiated congenic recipients along with support cells (Fig. 5D). PB chimerism analysis at 4 weeks was used to define progenitor mobilization.

Loss of osteoblasts in nondiabetic animals increased the number of progenitors in the blood even without G-CSF (Fig. 5E). These data suggest that osteoblasts play a role in HSPC retention in the bone marrow. With osteoblast depletion, G-CSF mobilization was compromised. Therefore, osteoblasts participate in the mobilizing ability of G-CSF; these two findings were in nondiabetic animals. In diabetic mice, the depletion of osteoblasts completely abolished any residual mobilizing potential of G-CSF (Fig. 5E). These data suggest that loss of osteoblasts observed in the bone marrow of diabetic animals may in part account for poor HSPC mobilization. However, other cells must participate, or the defect with osteoblast depletion without diabetes would have the same degree of compromise as the setting of osteoblast depletion with diabetes. Therefore, non-osteoblasts are a part of the STZ-induced mobilization defects.

Given that the mobilization process is proportional to the magnitude of the change of CXCL12 levels in the bone marrow microenvironment, it is reasonable to conclude that the nestin+ mesenchymal cells are a potential candidate for a non-osteoblast participant in G-CSF–mediated progenitor cell release. Thus, we wanted to explore how nestin+ cell function is compromised by diabetes.

The diabetic bone marrow shows increased sympathetic innervation and impaired response to β-adrenergic stimulation

To further assess potential mechanisms for compromised mobilization in diabetic animals, we evaluated SNS cells that had been previously shown to participate in G-CSF–induced HSPC mobilization. Specifically, treatment with G-CSF was noted in previous studies to increase sympathetic activity in the bone marrow, leading to Cxcl12 mRNA down-regulation in perivascular nestin+ mesenchymal stem cells and an increase in HSPC release (6, 7). We performed immunostaining for tyrosine hydroxylase, the rate-limiting enzyme in catecholamine synthesis, in the skulls of diabetic and control mice. Catecholaminergic nerve terminals were increased by more than twofold in the calvarial bone marrow of diabetic mice (n = 5, P < 0.05; Fig. 6, A and B). To determine whether increased catecholaminergic innervation was associated with altered sensitization of β-adrenergic receptors in Nestin-GFP+ cells, we treated diabetic and control Nes-Gfp transgenic mice with the β-adrenergic receptor agonist isoproterenol and measured Cxcl12 mRNA levels 2 hours later (Fig. 6, C and D). Reduced Cxcl12 mRNA levels in diabetic Nestin-GFP+ cells were confirmed under steady-state hematopoiesis. Further, the decrease in Cxcl12 mRNA levels after isoproterenol treatment in control animals was markedly blunted in diabetic mice (n = 6, P < 0.05; Fig. 6C). Similar results were obtained from unsorted bone and trabecular bone marrow cells (Fig. 6D).

Fig. 6

SNS dysautonomia is responsible for deregulation of CXCL12 gradient. (A) Representative pictures from whole mounting of calvaria of control (left panel) and STZ-treated (right panel) mice. Red signal, tyrosine hydroxylase. Scale bars, 250 μm. (B) Nerve terminal quantification in terms of perimeter in calvaria of STZ-treated versus control mice. n = 5. *P < 0.05. (C and D) Fold changes in the expression of CXCL12 mRNA in nestin+ cells (C) and bone marrow (D) from control and STZ-treated mice under baseline conditions and 2 hours after injection of isoproterenol (5 mg/kg, intraperitoneally) relative to control cells at baseline and normalized to GAPDH (ΔΔCT method). n = 6. *P < 0.05. (E) (Left) Western blot showing the amount of phospho-PKA (pPKA) in sorted nestin+ cells from control (+/− isoproterenol) and STZ-treated (+/− isoproterenol) mice. (Right) Histogram plot showing ratio of phospho-PKA to actin. (F) Fold changes in the expression of Cxcl12 mRNA in nestin+ cells from diabetic saline-treated and diabetic β3 blocker–treated (5 mg/kg per day for 10 days) animals relative to nondiabetic controls. Data are normalized to GAPDH (ΔΔCT method). n = 6. *P < 0.05. Percentage of donor cell engraftment 4 weeks after transplantation of 150 μl of mobilized PB from STZ-treated and control mice (along with whole bone marrow support cells) in congenic lethally irradiated recipients. (G and H) Mice were mobilized with a single dose of AMD3100 (G) or a single dose of AMD3100 at the end of the G-CSF mobilization regimen (H). All data were analyzed with unpaired, two-tailed Student’s t test.

Activation of the β-adrenergic receptor is known to induce adenosine 3′,5′-monophosphate (cAMP)–dependent protein kinase A (PKA) phosphorylation of the Thr197 residue in the catalytic subunit of PKA. Therefore, we assessed the amount of phospho-PKA in sorted Nestin-GFP+ cells from control and diabetic mice under baseline conditions and after isoproterenol administration (Fig. 6E). Isoproterenol markedly increased (~2.5-fold) the level of active PKA in control mice by Western blot using anti–phospho-PKAα/β (Thr197) antibody. Conversely, in diabetic mice showing higher baseline levels of phospho-PKA, isoproterenol induced only minor changes. β-Adrenergic signaling in Nestin-GFP+ cells is mediated by binding to and activation of β3-adrenoreceptor, selectively expressed on nestin-positive mesenchymal cells and absent on osteoblastic cells (28). Therefore, we administered a selective β3-adrenergic blocker (SR59230A) to diabetic mice and compared CXCL12 expression to that in diabetic mice injected with saline only. CXCL12 expression in sorted Nestin-GFP+ cells was restored by β3-adrenergic blockade, supporting a model of sympathetic hyperactivity causing aberrant CXCL12 regulation (Fig. 6F). These results are consistent with diabetes-related SNS dysfunction perturbing the molecular axis governing HSPC mobilization by G-CSF.

SNS-independent HSPC mobilization is preserved in diabetes

The impairment in stem cell mobilization in diabetic mice was observed upon G-CSF treatment. G-CSF action relies at least in part on the functional integrity of SNS terminals. Because we could not find any phenotypical or functional alteration of CXCR4 receptor expression by HSCs, we tested whether directly interfering with the interaction of CXCL12 with its cognate receptor, CXCR4, would affect HSPC mobilization. AMD3100 is a clinically used bicyclam antagonist that reversibly blocks the interaction between CXCL12 and CXCR4 (3134), prompting rapid HSPC mobilization. Mice were injected with AMD3100 (5 mg/kg), and equal volumes of blood were transplanted in lethally irradiated nondiabetic recipients, together with supporting cells. Evaluation of chimerism at 4 weeks showed virtually complete mitigation of the diabetic mobilization defect, because no significant differences in engraftment between mobilized diabetic and control HSPCs were observed (Fig. 6, G and H). AMD3100 alone is a weak HSPC mobilizer compared to G-CSF (35, 36). Concomitant administration of G-CSF and AMD3100 resulted in higher levels of donor chimerism and again demonstrated rescue of the mobilization abnormalities previously noted in diabetic mice. These data are consistent with involvement of CXCL12 modulation by cells of the marrow microenvironment in the diabetes-induced mobilization defect.

Discussion

Our results demonstrate that STZ-treated diabetic mice show a similar impairment in the G-CSF–induced mobilization of HSPCs to that observed in humans and that this defect is not the consequence of reduced bone marrow HSPC content or an intrinsic HSPC defect. Rather, the data support a model in which cells of the microenvironment are perturbed in diabetes and lose their ability to modulate CXCL12 expression in response to specific stimuli. Altered CXCL12 expression may blunt HSPC release from the bone marrow and can be overcome by direct pharmacological inhibition of the interaction of CXCL12 with its CXCR4 receptor.

The preservation or increase in HSPC numbers we observed is in contrast to recent reports of reduced LSK cells in diabetic mice (37, 38). These differences may reflect species differences or the long-term consequences of diabetes disease, which we did not assess in this study. All mice used in our experiments were <12 weeks of age. For the STZ model, animals were analyzed 5 to 8 weeks after the induction of diabetes, and we performed both immunophenotypic and functional assays. It should be pointed out that discrepancies in the quantitative estimation of progenitor cells in diabetes have been observed in other tissues, suggesting that the function rather than the quantity of progenitor cells may better define the deleterious effects of the disease (39, 40).

We documented some hematopoietic cell-autonomous changes in diabetes such as reduced migratory ability and increased adherence to extracellular matrix components. Similar findings have been reported for other cell types in diabetic settings (4144). These properties of diabetic HSPCs might participate in the altered mobilization capacity; however, they were not retained after transplantation into nondiabetic hosts, suggesting that the phenotype may be reversible. We did not further explore these features; rather, we focused on whether there were abnormalities in the bone marrow microenvironment. It has been reported that hyperglycemia induces persistent epigenetic changes in stromal (endothelial) cells (45). Correspondingly, we found that HSPC proliferation induced by diabetic bone marrow stromal cells was maintained even when HSPCs were cultured in normal glucose media for several days. Moreover, we showed that the bone marrow microenvironment of diabetic mice per se is sufficient to induce early proliferation of transplanted wild-type LT-HSCs, and this phenomenon is accompanied by changes in their localization. LT-HSCs tend to localize closer to the osteoblastic cells and the endosteal surface in diabetes at early and, most prominently, later time points. Diabetes reduced the number of osteoblastic cells, supporting the notion that bone formation is impaired in patients affected by the disease. By selectively depleting osteoblasts, we showed that they participate in the residual ability of diabetic animals to mobilize HSPCs in response to G-CSF, but that other bone marrow cell types also contribute. We hypothesize that those other cells are the nestin+ mesenchymal stem cells that we show have a defect in CXCL12 expression in response to G-CSF in diabetic mice. HSPC mobilization by G-CSF has been defined by others as involving altered CXCL12 expression (6, 7, 46). This has been reported to occur by G-CSF altering SNS activity, leading indirectly to down-regulation of CXCL12 expression (6, 7). SNS nerve terminals in the bone marrow are mainly located along arterioles and innervate perivascular cells (28, 47). Perivascular nestin+ mesenchymal stem cells have been shown to be innervated by SNS fibers, are known to serve as a major source of CXCL12 production in the mouse bone marrow, and down-regulate Cxcl12 in response to G-CSF or adrenergic stimulation (28). Although the overall content of CXCL12 in the bone marrow parenchyma was unchanged in STZ-treated animals, CXCL12 mRNA and protein levels in steady-state diabetic mice were reduced in stromal nestin+ mesenchymal cells. Furthermore, the profound suppression of CXCL12 synthesis that follows G-CSF administration disappeared in nestin+ cells from diabetic mice, suggesting that the loss of the chemokine directional gradient may be responsible for the defect in mobilization observed in diabetic settings. We also observed increased SNS terminals in diabetic calvaria bone marrow, associated with a blunted response of nestin+ cells to β-adrenergic agonists. It is known that diabetes is associated with increased SNS activation in humans (48, 49). These results strongly suggest that diabetes-related dysfunction of the autonomic nervous system may be responsible for the impaired function of nestin+ cells, leading to defective HSPC mobilization. However, given that G-CSF acts through multiple pathways to elicit HSPC mobilization (46), we cannot exclude the participation of additional mechanisms in the mobilization defect.

The importance of finding efficient mobilization strategies for diabetic patients is particularly evident in light of the delayed engraftment observed when bone marrow–derived stem cells are used as a source for autologous transplantation in these patients. Normal HSPC mobilization in diabetic mice using AMD3100, which directly blocks binding of CXCL12 to CXCR4, demonstrates that the disease does not affect this specific interaction, providing a pathophysiological rationale for the use of AMD3100 in the mobilization of HSPCs for patients with diabetes. Finally, we found that nestin+ cells with morphology and distribution similar to that of the murine nestin+ cells are also present in the human bone marrow, suggesting a possible similar role in human disease. Although the short-term diabetes mouse model we used may not reflect the late consequences of diabetes encountered in many patients, these studies serve as a guide for a more extensive evaluation of diabetic patients and provide a rationale for the use of alternatives to G-CSF for stem cell mobilization in patients with diabetes.

Materials and Methods

Diabetic mouse model

Diabetes was induced in 4- to 6-week-old C57BL/6 (Jackson Laboratory) Nes-Gfp (24) and Col2.3-Gfp (Jackson Laboratory) male mice with five consecutive intraperitoneal injections of STZ dissolved in citrate buffer (pH 4.5) (50 mg/kg per day). Mice were fasted for at least 6 hours before receiving the injection. Mice injected with citrate buffer alone were used as controls. B6.SJL-Ptprca Pep3b/BoyJ mice (SJL) (Jackson Laboratory) were used as transplant recipients. Diabetes onset was confirmed by measuring insulin levels [Ultra Sensitive Mouse Insulin ELISA (enzyme-linked immunosorbent assay), Crystal Chem Inc.] and blood glucose (GlucoMeter OneTouch Ultra). Only animals with glucose values higher than 300 mg/dl in a single measurement and concomitant low/absent level of insulin were further used for the experiments. Leptin receptor knockout (db/db) mice were purchased from the Jackson Laboratory. All mice were housed according to Institutional Animal Care and Use Committee guidelines and used at 12 to 18 weeks of age.

Experimental HSC mobilization and blood collection

Recombinant human G-CSF (Neupogen, Filgrastrim) was administered at the dose of 125 μg/kg every 12 hours for eight consecutive intraperitoneal injections. AMD3100 (Sigma) was used at 50 mg/kg in a single dose. PB samples were obtained through retro-orbital bleeding 3 hours after the last injection of G-CSF and 1 hour after AMD3100 injection. In detail, mice were anesthetized with a mixture of ketamine and xylazine, and blood was collected from the retro-orbital venous plexus through a microcapillary heparinized tube.

Flow cytometry and cell cycle analysis

Hematopoietic stem progenitors were identified on the basis of their expression of lineage markers as well as c-Kit, Sca-1, CD48, CD150, CD34, and CD135 (Flk-2) expression. Lineage staining was performed with a cocktail of biotinylated anti-mouse antibodies to Mac-1α (CD11b), Gr-1 (Ly-6G and Ly-6C), Ter119 (Ly-76), CD3, CD4, CD8a (Ly-2), and B220 (CD45R) (BD Biosciences). For detection, we used lineage-streptavidin conjugated with PacOrange (Invitrogen), c-Kit–APC (allophycocyanin) (CD117), CD48-FITC (fluorescein isothiocyanate) or PeCy5 (CD34), CD150-PE (phycoerythrin) (CD135), and Sca-1–APC–Cy7 or PeCy7 (BD Biosciences). For congenic strain discrimination, anti–CD45.1-APC and anti–CD45.2-FITC antibodies (BD Biosciences) were used. Adhesion molecule expression profile CXCR4, CD49D, CD49E, and CD62L were all PE-conjugated (eBioscience). CXCL12 labeling was performed with monoclonal anti-human/mouse CXCL12 antibody clone 79018 after fixation and permeabilization (Cytofix/Cytoperm kit, BD Biosciences) followed by secondary goat anti-mouse immunoglobulin G (IgG)–PE (both from R&D Systems). For cell cycle staining, cells were fixed overnight in 70% ethanol, washed twice in phosphate-buffered saline (PBS), and stained with a solution containing propidium iodide and ribonuclease (RNase) A in 0.1% Triton X-100. For cell cycle staining in HSPCs, 18 × 106 bone marrow cells were first stained with LSK and SLAMS markers, then fixed and permeabilized (Cytofix/Cytoperm kit, BD Biosciences), and incubated with Ki-67 FITC (BD Biosciences) (1:50) for 20 min at 4°C. After washing, cells were resuspended in 600 μl of washing buffer. Five microliters of 4′,6-diamidino-2-phenylindole (DAPI) (1 mg/ml) solution was added to the cell suspension. Acquisition was conduced with LSRII (BD Biosciences) after 30 min. Compensation and data analysis were performed with FlowJo 8.5.3.

CFU-C assay

CFU-C was performed as previously described (50). Briefly, bone marrow–derived cells or 50 μl of ACK-lysed PB cells was seeded into methylcellulose (Stemcell Technologies, M3434). Overall number of colonies was scored after 10 days.

Experimental PB transplantation

Blood (150 μl) from mobilized STZ-treated and control CD45.2 mice was mixed with 106 CD45.1 support cells and injected into lethally irradiated (9.5 Gy) SJL-45.1 recipients in the tail vein. Engraftment was monitored at 4-week intervals by enumerating donor CD45.1/.2+ cells in the PB by FACS analysis.

Bone marrow cell isolation

Mice were killed via CO2 asphyxiation; tibiae, femurs, and spine were removed and excess soft tissue was eliminated. Using a pestle and mortar, we crushed and washed the bones in PBS with 0.5% fetal bovine serum (FBS) and passed them through a 40-μm filter into a collection tube. Cells were spun at 1500 rpm for 5 min; the supernatant was removed, and cells were then further processed for staining, sorting, or transplantation.

Collagenase treatment of bone

Bones were cracked to remove bone marrow content. Bone fragments were further cut into small fragments and transferred into a prewarmed collagenase solution (Stemcell Technologies). Digestion was performed for 1 hour at 37°C under vigorous shaking. The solution was then filtered through a 40-μm filter. The flow-through was then pelleted and further processed for FACS analysis or RNA extraction.

RNA isolation and quantitative real-time reverse transcription–polymerase chain reaction

Nestin and Col2.3 cells were FACS-sorted for GFP protein directly into lysis buffer, and RNA isolation was performed with the Dynabeads mRNA DIRECT Micro kit (Invitrogen). Primer sequences and reverse transcription–polymerase chain reaction (RT-PCR) procedure used have been previously described (28).

Experimental bone marrow transplantation

All bone marrow transplantations were performed by tail vein injection. For competitive transplantation, 106 whole bone marrow cells from diabetic or control (CD45.2) mice were mixed with 106 CD45.1+ (competitor) wild-type cells and injected into lethally irradiated (9.5 Gy, one dose on the day of transplant) recipient BL6-SJL (CD45.1+) mice. Engraftment efficiency in recipients was monitored by donor contribution of CD45.2+ cells with FACS analysis.

Adhesion and migration assays

For adhesion assays, 2 × 103 FACS-sorted LSK cells were plated in triplicate on 24-well culture dishes precoated with fibronectin. The adherent fraction was dissociated by treatment with cell dissociation buffer, enzyme-free (Invitrogen), collected, and plated in MethoCult media (M3434, Stemcell Technologies). Progenitor adhesion was determined as the percentage of CFU-C in the adherent fractions relative to the frequency of CFU-C in the input sample.

For migration assays, 2 × 103 FACS-sorted LSK cells were suspended in 500 μl of RPMI and seeded in triplicate in the upper chamber of 24-well, 5-μm Transwell plates (Corning). Cells were incubated for 1 hour at 37°C in the presence or absence of SDF-1α (stromal cell–derived factor 1α), after which the supernatant was removed and the wells were washed once with PBS to remove nonadherent cells. Adherent cells were harvested by treatment with cell dissociation buffer, enzyme-free (Invitrogen), and plated in methylcellulose. Progenitor adhesion was determined as mentioned above.

Imaging of the HSC niche

Mice were anesthetized and prepared for in vivo imaging as previously described (51). FACS-sorted HSCs were stained in PBS for 15 min at 37°C with DiD (Invitrogen) or DiI (Invitrogen) using a 1:200 dilution and injected into lethally irradiated recipients. An about 4 × 6–mm area of the calvarium comprising the central sinus and the surrounding bone marrow cavities within the left and right frontal bones was scanned. When using Col2.3GFP and Nestin-GFP reporter mice, three-dimensional models of the osteoblastic or nestin cells and the bone (second-harmonic generation) were developed through Z-stack reconstruction. Using the Pythagoric theorem, we determined the location of stained HSCs injected in relation to the bone and the GFP+ cells. The images were analyzed with ImageJ software (National Institutes of Health; http://rsbweb.nih.gov/ij/).

Immunostaining of mice calvaria

Calvaria from diabetic and control mice were fixed in 4% paraformaldehyde for 1 hour and subsequently washed twice in PBS. Blocking of unspecific binding and permeabilization were achieved with 20% goat serum and 0.5% Triton X-100 in PBS overnight. Endogenous biotin was blocked with Vector Kit (catalog number Sp-2001) following the manufacturer’s instructions. Primary antibody (Chemicon rabbit anti–tyrosine hydroxylase; BD Pharmingen APC-conjugated anti-CD31) was applied at 1:100 in 20% goat serum and 0.1% Triton X-100 in PBS and incubated for 2 to 3 days. Samples were washed with 0.1% Triton X-100 in PBS and incubated overnight with a secondary biotinylated goat anti-rabbit antibody (1:200) in 20% goat serum and 0.1% Triton X-100 in PBS. After washing with 0.1% Triton X-100 in PBS, samples were incubated for 2 hours with ABC kit (Vector PK-6100) prepared 30 min before use. After washes with PBS, samples were incubated with Cy3-tyramide (Perkin Elmer) (1:200) in diluent reagent for 30 min, washed with PBS, and visualized by fluorescence microscopy. Projections of Z-stacks (100 to 200 μm) of calvaria bone marrow were analyzed with the SlideBook software (Intelligent Imaging Innovations).

Drug administration

A single injection of isoproterenol (Sigma; 5 mg/kg, intraperitoneally) was administered to diabetic and control mice. SR59230A (Sigma; 5 mg/kg, intraperitoneally) was administered daily for 10 days to diabetic mice.

Mice treated with saline were used as a control. Animals were killed 2 hours after injection by CO2 asphyxiation, and the femora were flushed with 200 μl of PBS. TRIzol (Invitrogen) was used for total RNA extraction.

Osteoblast deletion

Mice in which expression of Cre recombinase was driven by osteocalcin promoter (29) were crossed with mice harboring an iDTR (30). In the double-transgenic mice, Cre-mediated removal of a transcriptional STOP cassette allows the expression of DT receptor. Upon DT administration, efficient ablation of the target population was achieved. Mice with only one transgene were used as controls. Both double- and single-transgenic mice were treated either with STZ for diabetes induction or with saline. After 5 weeks, DT was administered at the dose of 100 ng per mouse twice a day for 14 days. Nine days after the toxin administration was started, G-CSF was administered (eight injections of 125 μg/kg every 12 hours). At day 14, blood was collected and used for flow and transplantation assays.

Enzyme-linked immunosorbent assay

CXCL12 protein levels were assessed in bone marrow supernatant and PB plasma. Briefly, femurs from diabetic and control mice were flushed four times with 100 μl of PBS (total volume) in Eppendorf tubes. PB was collected via retro-orbital bleeding in Eppendorf tubes without adding anticoagulant. The samples were then incubated at 37°C for 1 hour and spun at 4.6 rpm for 5 min. Plasma was then collected and used for CXCL12 evaluation with ELISA kit (Ray Biosystem) following the manufacturer’s instructions.

Insulin levels were assessed in PB plasma with ELISA kit (Crystal Chem Inc.) following the manufacturer’s instructions.

Western blot

Nestin-GFP cells were sorted with FACSAria. Western blot was performed with anti–phospho-PKA antibody (Invitrogen) following the manufacturer’s instruction.

Statistical analysis

Mice. Unless otherwise specified, unpaired, two-tailed Student’s t test was used and data have been plotted as average ± SEM. Statistical significance is indicated as follows: *P < 0.05; **P < 0.01; ***P < 0.001.

Human. Fisher’s exact test was used to compare the frequency of diabetes between poor and good mobilizers. The Wilcoxon two-sample test was used to compare the distribution of CD34+ cells per kilogram and glucose levels between patient groups. Neutrophil recovery and platelet engraftment were measured from the day of transplantation with PB stem cells or bone marrow–harvested cells. The cumulative incidence of engraftment was estimated with the Kaplan-Meier method, and the difference between diabetic and nondiabetic patients was assessed by the log-rank test. All P values were based on a two-sided hypothesis and computed with SAS 9.2 (SAS Institute).

Supplementary Material

www.sciencetranslationalmedicine.org/cgi/content/full/3/104/104ra101/DC1

Materials and Methods

Fig. S1. Recovery of patients after stem cell transplantation.

Fig. S2. Diabetes-induced changes in stem cells and their microenvironment.

Fig. S3. Detection of nestin+ cells in human bone marrow biopsies.

Table S1. Details of the human caseload included in the study.

Footnotes

  • * These authors contributed equally to this work.

References and Notes

  1. Acknowledgments: We thank D. Dombkowski, L. B. Prickett-Rice, K. Folz-Donahue, and S. Lahiri for cell sorting expertise and E. Corradini for technical support. We are grateful to C. Lo Celso for help and advice. Funding: F.F. was supported by fellowships from Collegio Ghislieri, Associazione Italiana Leucemie, Associazione Cristina Bassi, and National Heart, Lung, and Blood Institute (NHLBI) U01HL100402. S.M.-F. is thankful to American Society of Hematology (ASH) for the supporting scholarship. B.S. was supported by the Chamber of Industry and Commerce of the Government of Spain. S.M.S. was supported by NHLBI 5T32HL007623-24. P.S.F. was funded by NIH grants R01HL097819 and R01DK056638. F.Q. was supported by the following grants: FP7-BIOSCENT, NMP-214539 2007, PRIN AL2YNC 2007, and Italian Ministry of Health THEAPPL 2008. D.T.S. is supported by the NIH NHLBI HL097794, HL097748, HL100402 and DK050234, the Ellison Foundation, and the Harvard Stem Cell Institute. Author contributions: F.F. and S.L. designed and performed the experiments, analyzed the data, and wrote the manuscript. S.M.-F. performed the nerve quantitative analysis and RT-PCR on nestin+ cells and osteoblasts, analyzed the data, and edited the manuscript. B.S. performed the experiments and edited the manuscript. J.A.S. helped with in vivo imaging experiments. B.Y.Y. performed the statistical analysis on patient data. E.M., G.G., L.P., E.L.R., M.M., and V.R. performed the staining on human biopsies and analyzed the human data. S.M.S. performed the experiments and edited the manuscript. C.P.L. and P.S.F. contributed reagents and provided advice on the manuscript. F.Q. and D.T.S. supervised the experiments and the overall study, interpreted the data, and wrote the manuscript. Competing interests: D.T.S. is a consultant to Fate Therapeutics, Genzyme, Hospira, and Bone Therapeutics and a shareholder in Fate Therapeutics. P.S.F. is a paid consultant for Glycomimetics.
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