Insufficient stromal support in MDS results from molecular and functional deficits of mesenchymal stromal cells

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Ineffective hematopoiesis is a major characteristic of myelodysplastic syndromes (MDS) causing relevant morbidity and mortality. Mesenchymal stromal cells (MSC) have been shown to physiologically support hematopoiesis, but their contribution to the pathogenesis of MDS remains elusive. We show that MSC from patients across all MDS subtypes (n=106) exhibit significantly reduced growth and proliferative capacities accompanied by premature replicative senescence. Osteogenic differentiation was significantly reduced in MDS-derived MSC, indicated by cytochemical stainings and reduced expressions of Osterix and Osteocalcin. This was associated with specific methylation patterns that clearly separated MDS–MSC from healthy controls and showed a strong enrichment for biological processes associated with cellular phenotypes and transcriptional regulation. Furthermore, in MDS–MSC, we detected altered expression of key molecules involved in the interaction with hematopoietic stem and progenitor cells (HSPC), in particular Osteopontin, Jagged1, Kit-ligand and Angiopoietin as well as several chemokines. Functionally, this translated into a significantly diminished ability of MDS-derived MSC to support CD34+ HSPC in long-term culture-initiating cell assays associated with a reduced cell cycle activity. Taken together, our comprehensive analysis shows that MSC from all MDS subtypes are structurally, epigenetically and functionally altered, which leads to impaired stromal support and seems to contribute to deficient hematopoiesis in MDS.


Cytopenias of varying degrees are the hallmark of myelodysplastic syndromes (MDS) and substantially impact the quality of life and cause relevant morbidity and mortality.1, 2 As MDS are considered clonal disorders of hematopoietic stem and progenitor cells (HSPC), the etiology of ineffective hematopoiesis has been mainly attributed to molecular alterations of these CD34+HSPC.3, 4, 5 The bone marrow (BM) microenvironment, in which MDS-derived HSPC reside, has also been considered to contribute to MDS pathogenesis. However, its role has been investigated in considerably less detail.6, 7

Mesenchymal stromal cells (MSC) are an integral component of the BM microenvironment and have an indispensable role in the regulation and support of HSPC.8, 9 Through this interaction, MSC are involved in orchestrating the balance of HSPC self-renewal and differentiation, thereby helping to ensure the life-long supply of mature blood cells. In addition, MSC exhibit immunoregulatory properties and can differentiate into osteoblasts.9, 10 Although it has been demonstrated that the genetic disruption of cells of the mesenchymal/osteoblastic lineage can induce an MDS-like phenotype in mice,11 knowledge about MSC in humans with MDS is limited. This has created controversies regarding their contribution to MDS pathogenesis. Genetic aberrations have been reproducibly found in about one quarter of MDS-derived MSC.12, 13, 14, 15 Furthermore, several cytokines, adhesion molecules and transcription factors have been described to be altered in MDS-derived MSC,16, 17, 18 but it is unclear whether and how these abnormalities influence the pathogenesis of MDS. Conflicting results have also been reported with regard to the biological behavior of MDS–MSC. Although their clonogenic potential and hematopoietic support capacities were shown to be reduced in some MDS patients,15, 19, 20, 21 other data suggested a proliferative advantage and normal hematopoietic support by MDS-derived MSC.17, 22, 23

In order to investigate whether and how MSC contribute to ineffective hematopoiesis in MDS, we conducted a detailed analysis of molecular and functional properties of MDS-derived MSC. Our analysis is the first to include a sufficient number of samples from patients with early MDS, advanced MDS and acute myeloid leukemia with MDS-related changes (sAML), and should thus overcome difficulties in interpretation that may arise from the marked heterogeneity of MDS.

Patients and methods

Patients and samples

BM specimens were obtained from 121 patients with newly diagnosed MDS (median age 66 years, range: 26–91 years). According to the WHO 2008 classification of MDS, 45 patients (37%) had refractory cytopenia with multilineage dysplasia, 29 patients (24%) had refractory anemia with excess blasts, 13 patients (11%) had MDS associated with isolated del5q (MDS del5q), 4 patients (3%) had chronic myelomonocytic leukemia, 2 patients (1%) had MDS unclassified and 28 patients (24%) had sAML (Table 1).

Table 1 Patients demographics

From 72 of these patients only MSC were generated, whereas in 15 patients only CD34+ HSPC were isolated. In 34 patients, paired samples of MSC and CD34+ HSPC were available (Supplementary Table S1, Supplementary Figure S1).

BM samples from 67 age- and sex-matched healthy individuals (median age 63 years, range: 31–85 years, P=0.3324) served as controls. Signed informed consent was obtained from all patients and donors, and the study was performed in accordance with the Declaration of Helsinki and approved by our local institutional review board.

Isolation, expansion and characterization of MSC

Following the isolation by density centrifugation, 2–5 × 107 mononuclear cells were cultured in Dulbecco’s modified Eagle’s Medium low glucose containing 30% fetal bovine serum and 1% Penicillin/Streptomycin/L-Glutamine (Sigma-Aldrich, St Louis, MO, USA) at 37 °C, 5% CO2 in a humidified atmosphere as previously described.24 Non-adherent cells were removed and medium was changed weekly. After an incubation period of 14–21 days, colonies>50 cells (colony-forming unit fibroblast, CFU-F) were counted by light microscopy.25 Cumulative population doublings and doubling time were calculated as described before.26 All experiments were carried out using MSC derived from passages 2–4. To fulfill the criteria of the International Society for Cellular Therapy and to exclude contamination of MSC cultures by hematopoietic cells MSC were analyzed for the expression of CD34, CD45, CD73, CD90, and CD105, by flow cytometry.

MSC differentiation

In addition to the analysis of surface markers, differentiation assays were performed in passage 3. Osteogenic differentiation was induced by adding dexamethasone (10−7 M), ascorbic acid (50 μg/ml) and β-glycerolphosphate (10 mM, all Sigma-Aldrich) and subsequently visualized by Alizarin red, van Kossa and alkaline phosphatase staining. Adipogenic and chondrogenic differentiation were performed as previously described (for details see also Supplementary Information).24, 27 All images were captured using an Axiovert 25 microscope (Zeiss, Jena, Germany) using either a × 2.5 objective (Zeiss EC Plan-Neofluar × 2.5x/0.075, for chondrogenic differentiation), × 5 objective (Zeiss CP-Achromat × 5 Ph0, for native and osteogenic differentiated MSC) or a × 10 objective (Zeiss CP-Achromat × 10 Ph1, for adipogenic differentiation) and digitalized with Spot Software (Diagnostic Instruments Inc., Sterling Heights, MI, USA). For the purpose of quantification, the differentiation was graded according to microscopic analysis of staining intensity as follows: 0=absent; 1=weak; 2=moderate; 3=intensive.

Furthermore, mRNA expression levels of Osterix and Osteocalcin (OC), both established markers of early and late osteogenesis, were analyzed by quantitative real-time PCR in MSC during osteogenic induction as well as under steady-state conditions.

OC enzyme-linked immunosorbent assay

OC serum levels were measured on a Modular Analytics E 170 analyzer (Roche Diagnostics, Mannheim, Germany) using Elecsys N-MID OC immunoassay according to the manufacturers’ instructions.

Isolation of CD34+ cells

Mononuclear cells were obtained from BM aspirates by density gradient separation and then subjected to immunomagnetic enrichment of CD34+ cells (Miltenyi Biotec, Bergisch Gladbach, Germany) as published.28, 29

Flow cytometry

All flow cytometric analyses were performed using a FACSCalibur (BD Biosciences, Heidelberg, Germany) and data were analyzed with FCS Express V3 software (De Novo Software, Los Angeles, CA, USA). A list of antibodies is provided in the supplement (Supplementary Table S2).

Quantitative real-time PCR

RNA from MSC was purified using the RNeasy mini kit in combination with the RNAse-free DNase kit (both Qiagen, Hilden, Germany) according the manufacturer’s instructions. All experiments were carried out in duplicate on a StepOne Plus Real-time PCR Cycler using SYBR Green PCR Master Mix (both Applied Biosystems, Life Technologies, Carlsbad, CA, USA). All primer sequences can be provided on request. GAPDH served as reference control, and differences in mRNA expression levels were calculated as fold-changes by the ΔΔCt method.

Analysis of cellular senescence in MSC by β-galactosidase staining

β-Galactosidase activity as an indicator of senescence was determined in MSC from hematopoietic cell and MDS patients using the Cellular Senescence Detection Kit (Biolabs, San Diego, CA, USA) as recommended by the manufacturer. Cells were counted by light microscopy and the fraction of senescent cells (β-galactosidase positive) was assessed.

Methylation analysis

Genome-scale methylation profiles were obtained as described previously30 using Infinium methylation 450 K arrays. Details about the statistical analysis of the methylation data sets are provided in the Supplementary Information. Specific gene methylation patterns were validated by 454 (Roche) bisulfite sequencing, as described previously.31 Primer sequences are provided in Supplementary Table S3.

Angiopoietin-1 enzyme-linked immunosorbent assay

Culture supernatants from MSC were collected by standardized methods. Levels of Angiopoietin-1 were measured using the Angiopoietin-1 Immunoassay (R&D Systems GmbH, Wiesbaden, Germany) according to the manufacturer’s protocol. Data were analyzed using a Perkin Elmer Wallac Victor2 1420 Microplate Reader.

Long-term culture-initiating cells

Initially, 1.2 × 106 MSC were cultivated on 96-well plates (Costar, Corning Incorporated, Corning, NY, USA) until at least 80% confluence was reached and then irradiated with 30 Gray using Gulmay RS225 X-ray equipment. Afterwards, 6 × 103 CD34+ cells were plated on these MSC feeder layers and cultivated in 5 ml long-term bone marrow culture (LTB-MC) medium for 5 weeks with weekly changes of culture medium. At the end of that period, the medium was replaced by clonogenic methylcellulose medium H4534 (Stem Cell Technologies, Vancouver, Canada) containing 10 IU/ml erythropoietin (NeoRecormon 1000 IU; Roche), and cells were cultured for another 2 weeks at 37 °C, 5% CO2 in a humidified atmosphere. Finally, colonies were counted under a light microscope.

Cell cycle analysis of CD34+ HSPC after coculture with MSC

1.6 × 104 MSC/cm2 were seeded on six-well plates Dulbecco’s modified Eagle’s medium low glucose (Sigma-Aldrich) to build a feeder layer and were washed after 1 day with PBS. On these MSC feeder layers, 2.1 × 104/cm2 CD34+ HSPC were cultured in IMDM containing 20% fetal bovine serum, 1% Penicillin/Streptomycin/L-Glutamine and cytokines (IL3, IL6, FLT3 and SCF 10 ng/ml). After 72 h of coculture, CD34+ cells were harvested and stained with Hoechst 33 342, intracellular Ki-67 and a fluorescein isothiocyanate-conjugated anti-CD34 antibody for 30 min as previously described.24

Statistical analysis

Statistical analyses were performed using Prism 5.01 (GraphPad Software Inc., La Jolla, CA, USA). For interindividual comparison the two-sided unpaired Student’s t-test was used, whereas for intraindividual analysis the Wilcoxon signed rank test was used. For all experiments means and s.e.m. are given. Statistical significance was established at P0.05.


Impaired growth kinetics and increased cellular senescence of MSC from patients with MDS

Purified MSC from healthy controls and from patients with MDS were positive for MSC markers CD73, CD90 and CD105 (>95% of cells) and did not express detectable levels of hematopoietic antigens CD34 and CD45 (<2% of cells) (Figure 1a). MSC layers were assessed using light microscopy and showed the characteristic fibroblastoid appearance for MSC from healthy controls, whereas MSC from MDS patients were larger and appeared disorganized (Figure 1b).

Figure 1

Phenotype, growth properties and senescence of MDS-derived MSC. (a) Mean fluorescence intensity (MFI) of CD34, CD45, CD73, CD90 and CD105 was determined on MSC of hematopoietic cell (HC) and MDS by flow cytometric analysis. Representative histograms are shown. Red line represents the respective isotype control and black line represents MDS-derived MSC. (b) Representative micrographs depict morphology of MDS- and HC-derived MSC. Scale bars indicate 100 μm. Bar charts show CFU-F normalized to 1 × 107 plated BM mononuclear cells (MNC) (HC n=58; all MDS n=58; refractory cytopenia with multilineage dysplasia (RCMD) n=29; refractory anemia with excess blasts (RAEB) n=15; sAML n=10) (c), maximum number of passages (HC n=31; all MDS n=36; RCMD n=19; RAEB n=10; sAML n=8) (d), and cumulative population doublings (CPD) (HC n=31; all MDS n=36; RCMD n=19; RAEB n=10; sAML n=8) (e). Results are shown for healthy controls (white bars), all MDS patients (black bars), patients with RCMD (diagonal striped bars), RAEB (longitudinal striped bars) and sAML (dotted bar). Cellular senescence was assessed by β-galactosidase staining in MSC (passage 3) of HC (n=12, white bar) and MDS patients (n=12, black bar). One-hundred MSC per individual sample were counted using light microscopy and the percentage of senescent cells (β-galactosidase positive=blue) was determined. One representative experiment is shown and scale bars indicate 100 μm (f). All bars indicate mean and all error bars indicate s.e.m. Unpaired Student’s t-test was used to detect statistically significant differences between HC and MDS subtypes. Asterisks display P-values *P<0.05, **P<0.01, ***P<0.001.

To assess the clonogenic potential of MSC derived from healthy controls and patients with MDS and sAML,CFU-F assays were performed. CFU-F counts of patients with MDS and sAML were significantly reduced in comparison with healthy controls (Figure 1c). Accordingly, MDS-derived MSC could be maintained in culture for a significantly lower number of passages (MDS mean±s.e.m.: 5±0.41; healthy controls mean±s.e.m.: 12±0.58, Figure 1d) and proliferated slower, as indicated by lower cumulative population doublings (MDS mean±s.e.m.: 20.47±0.59; healthy controls mean±s.e.m.: 28.12±0.97, Figure 1e) and a longer time to achieve an equivalent cumulative population doublings (passage 3: MDS mean±s.e.m.: 18.42±0.26, 36 days; healthy controls mean±s.e.m.: 18.56±0.28, 32 days, not shown). This finding remained significant when we looked at MDS subtypes separately.

In line with these findings, MSC from patients with MDS were more prone to cellular senescence, as shown by a significantly higher number of SA-β-gal-positive cells, (mean±s.e.m.: 46.08%±6.95%) compared with healthy controls (mean±s.e.m.: 26.92%±4.84%, Figure 1f). This premature cellular replicative exhaustion might represent a mechanism responsible for the impaired growth of MSC observed across all MDS subtypes.

Specific methylation pattern of MDS-derived MSC

Senescence of MSC has been previously associated with DNA methylation changes.32 To explore the possibility that changes in DNA methylation could underpin the observed phenotypic alterations, we used Infinium 450 K arrays to investigate the methylation patterns of 450 000 CpG dinucleotides in a set of 12 MSC samples. DNA samples were prepared from four healthy donors, four patients with refractory cytopenia with multilineage dysplasia and four patients with refractory anemia with excess blasts. To minimize the effects of epigenetic variations unrelated to MDS, all samples were obtained from male donors and similar age groups. High-quality methylation profiles were obtained for all 12 samples and showed a very high similarity between samples (Supplementary Figure S2A). Furthermore, all profiles showed a characteristic bimodal distribution of methylation values (Supplementary Figure S2B), with similar shapes and ranges, which further confirmed the high quality of the methylation data set.

Further statistical analysis of the methylation data (see Supplementary Materials and Methods for details) identified 252 differentially methylated CpGs. Interestingly, these CpGs consistently showed pronounced methylation differences between healthy controls and MDS samples (Figure 2a), indicating highly specific methylation changes in the MSC from MDS patients. We also used principal component analysis of the 252 differentially methylated probes to explore clustering of samples and to identify outliers. The results showed specific clustering, and healthy control samples were clearly separated from the two MDS populations (Figure 2a). These data indicate for the first time the presence of specific methylation patterns for the three cell populations analyzed and suggest that a comparably small set of methylation markers can be used to separate MSC from patients and healthy controls.

Figure 2

Methylation analysis of MSC from patients with MDS and from healthy controls. (a) Cluster analysis of 252 differentially methylated CpGs shows highly consistent differences between MDS patients (green: RAEB, purple: RCMD) and healthy controls (orange). Upper panel: cluster dendrogram, lower panel: principal component analysis. (b) Validation of array-predicted methylation differences by 454 bisulfite sequencing of sample pools. Arrowheads highlight CpGs represented on the array, blue scale bars indicate 20 bp. Vertical lines indicate individual CpG dinucleotides. Methylation maps show sequencing reads (rows) and the methylation of individual CpGs (columns). Methylated CpGs are shown in red, unmethylated CpGs in green and gaps in white. Numbers indicate the number of sequencing reads. (c) 454 bisulfite sequencing of independent, individual samples. Methylation maps show the average methylation ratios for individual CpGs, sequencing coverages ranged from 160–2626 reads. (d) mRNA expression for the three hypermethylated candidate genes TBX15, HOXB1 and PITX2 was quantified by real-time PCR (RT-PCR) in MSC of eight healthy controls (white bars) and 15 MDS patients (black bars). Means and s.e.m. are given and Student’s t-test was used for comparison of healthy controls and MDS patients. Asterisks display P-values *P<0.05, **P<0.01.

Subsequent data analysis revealed 65 differentially methylated genes, with the majority containing a single differentially methylated probe. However, a subset of genes from the HOXB cluster contained up to 8 differentially methylated genes and the PITX2 and TBX15 genes contained 11 and 13 differentially methylated CpGs, respectively (Supplementary Figure S2C), which suggests that these genes are particularly affected by methylation changes. To validate and further analyze the array-predicted methylation differences for these genes, we used deep bisulfite sequencing. Fragments were PCR amplified from the same sample pools and sequenced using 454 technology, which routinely generated sequencing coverages of 100X and more. The sequencing results were in excellent agreement with the array-based predictions and demonstrated consistent hypermethylation for all three genes in MSC from MDS patients (Figure 2b). Furthermore, we also used 454 bisulfite sequencing to analyze methylation patterns of HOXB1, PITX2 and TBX15 in MSC from an additional set of 10 independent MDS and healthy control samples. The results again showed MDS-dependent hypermethylation for all genes and for all samples analyzed (Figure 2c). For TBX15 and HOXB1, hypermethylation was located in the promoter regions of these genes (Supplementary Figure S3) and resulted in significant downregulation of mRNA expression (Figure 2d), consistent with the well-established gene-silencing function of promoter methylation.33 For PITX2, sequencing revealed a pronounced hypermethylation of the gene body, which resulted in significant overexpression of PITX2 mRNA in MDS-derived MSC (Figure 2d). The latter finding is in agreement with the positive correlation between gene body methylation and gene expression that has been described in several previous studies.33, 34

Finally, we performed pathway analysis to explore the biological relevance of our MDS-dependent hypermethylation pattern. Gene ontology analysis of the 65 differentially methylated genes showed a strong enrichment for biological processes associated with cellular phenotypes and for molecular functions associated with transcriptional regulation (Supplementary Figure S2D), thus suggesting that MDS-related DNA methylation changes might affect the functional properties of MSC.

MSC from MDS patients exhibit decreased osteogenic differentiation potential

To further characterize the phenotype of MSC, we investigated their trilineage differentiation potential.35 The results showed that MSC from MDS patients exhibited no significant differences with regard to adipogenic or chondrogenic differentiation (Supplementary Figure S4). In contrast, osteogenic differentiation potential of MDS-derived MSC was significantly reduced as indicated by the respective cytochemical stainings (Figures 3a and b). Consistently, the mRNA level of Osterix, a transcription factor critically involved in the early differentiation process toward osteoblasts, was significantly reduced in MSC from all MDS subtypes under steady-state conditions and during osteogenic differentiation (Figures 3c and e). The mRNA expression of OC, an established marker of mature osteoblasts,36 appeared moderately reduced under steady-state conditions in sAML and MDS cases with an elevated blast count (Figure 3d), and was significantly diminished when osteogenic differentiation was induced (Figure 3f). In agreement with this finding, serum OC levels were significantly lower in patients with MDS with an elevated blast count (5%) and in patients with sAML in comparison with healthy controls (Figure 3g). As the deletion of Dicer1 was recently shown to result in an impaired osteogenic differentiation of MSC associated with MDS evolution in mice,11 we investigated Dicer1 expression in human MSC and found a significant downregulation of Dicer1 mRNA in MDS and sAML (Figure 3h). These findings suggest that the impaired osteogenic differentiation of MSC also contributes to the pathophysiology of MDS in humans.

Figure 3

Impaired osteogenic differentiation potential of MSC from patients with MDS. (a) After 14 days of differentiation, Alizarin red, van Kossa and alkaline phosphatase (ALP) staining was used to visualize osteogenic differentiation. Representative micrographs of MSC derived from HC and different MDS subtypes are shown. Scale bars indicate 100 μm. (b) Differences with regard to osteogenic differentiation between HC (n=11, white bar), MDS (n=10, black bar) and sAML (n=3, dotted bar) were quantified by the score described in the Methods section. mRNA expression of Osterix (c) was determined using RT-PCR without osteogenic induction in MSC of HC (n=12, white bar), MDS (n=20, black bar) and sAML (n=5, dotted bar). (d) displays OC mRNA under steady-state condition in HC (n=12, white bars), MDS of various subtypes (n=24, black bars) and sAML (n=6, dotted bar). In addition, results are shown separately RCMD (n=11, diagonal striped bars) and RAEB (n=7, longitudinal striped bars). Following osteogenic induction Osterix (e) and OC (f) mRNA was quantified by RT-PCR at given time points in MDS (n=8, pointed line) and HC (n=6, line with squares). In an additional cohort of 20 newly diagnosed MDS patients (black bar) and in five untreated patients with sAML (dotted bar). OC serum levels were measured and also depicted separately for patients with BM blasts < 5%, (n=11, diagonal striped bars) and>5% (n=9, longitudinal striped bars) (g). (h) mRNA expression of Dicer1 was determined using RT-PCR without osteogenic induction in MSC of HC (n=33, white bar), MDS (n=31, black bar), RCMD (n=15, diagonal striped bars), RAEB (n=8, longitudinal striped bars) and sAML (n=7, dotted bar). For all experiments means and s.e.m. are given and Student’s t-test was used for comparison of HC and MDS subgroups. Asterisks display P-values *P<0.05, **P<0.01, ***P<0.001.

Altered chemokine and cytokine expression of MDS-derived MSC

Within the BM microenvironment, MSC physiologically interact with CD34+ HSPC through several soluble factors as well as surface molecules.9, 37 We suspected alterations of such molecules to be present in MDS-derived MSC and investigated them systematically. We found that the mRNA expression of Osteopontin was elevated in MDS-derived MSC, whereas the expression of Kit-ligand was significantly diminished across all MDS subtypes (Figure 4a). Furthermore, mRNA expression of Angiopoietin-1 was significantly decreased in MSC from MDS patients, consistent with a reduced Angiopoietin-1 concentration in supernatants of MDS-derived MSC (Figure 4b). On the other hand, Jagged1 was significantly overexpressed in MDS–MSC on mRNA level as well as on protein level (Figure 4c).

Figure 4

Expression of key molecules in MSC involved in the interaction with CD34+ HSPC. (a) mRNA expression of Osteopontin and Kit-ligand (Kitlg) in MSC measured by quantitative RT-PCR is shown (HC n=21; MDS n=27; sAML n=7). (b) mRNA expression of Angiopoietin-1 (Angpt1) was measured in MSC (HC n=21; MDS n=19; sAML n=7) by quantitative RT-PCR. Angpt1 concentrations in supernatants of MSC cultures (HC n=8; MDS n=15) were assessed by enzyme-linked immunosorbent assay (ELISA). Bar charts display mean Angpt1 concentrations. (c) mRNA expression of Jagged1 was measured in MSC (HC n=23; MDS n=16; sAML n=6) by quantitative RT-PCR. MFI of Jagged1 was determined on MSC of HC (n=29) and MDS (n=23) by flow cytometry. Representative histogram plots as well as bar charts are shown. (d) mRNA expression of seven chemokines was measured in MSC (HC n=26; MDS n=30; sAML n=6) by quantitative RT-PCR. All RT-PCR experiments were performed in duplicate and fold-changes were calculated by the ΔΔCt method. For all experiments means and s.e.m. are given and Student’s t-test was used for comparison of HC and MDS subgroups. Asterisks display P-values *P<0.05, **P<0.01, ***P<0.001.

In addition, mRNA expression of 11 chemokines was investigated, showing significant downregulation of five chemokines (CXCL1, CCL2, CXCL14, CX3CL1 and midkine) in MDS-derived MSC (Figure 4d). In contrast, CCL26 was significantly upregulated in MDS-derived MSC, whereas CXCL7 mRNA expression was overexpressed only in MDS and downregulated in sAML. No differences of mRNA expression were found for CXCL8, CXCL10, CXCL12 and CCL23 (Supplementary Figure S5). These findings suggest complex alterations in chemokine and cytokine production by stromal cells in the BM microenvironment of MDS patients.

MDS-derived MSC have a diminished stem cell supporting capacity

Given these substantial alterations of MDS-derived MSC, we were interested whether this functionally translated into an impaired ability to support CD34+ HSPC. We found that MDS-derived MSC have a 19-fold reduced ability to support healthy CD34+ HSPC in long-term culture assays in comparison with healthy MSC (LTC-IC frequency on healthy MSC mean±s.e.: 1.01%±0.42%; on MDS–MSC mean±s.e.: 0.05%±0.013%) (Figure 5ai). When MDS CD34+ HSPC were cultured on MDS-derived MSC, their LTC-IC frequency (on MDS–MSC mean±s.e. 0.02%±0.011%) was also low comparable to that observed in healthy CD34+ HSPC cultured on MDS–MSC. However, when MDS HSPC of the same patients were cultured on healthy MSC, their LTC-IC frequency was partially restored (on healthy MSC mean±s.e. 0.342%,±0.09%, P=0.0076) (Figure 5aii).

Figure 5

MDS-derived MSC have a diminished stem cell supporting capacity. CD34+ HSPC of 11 healthy controls were cultured on MSC feeder layer derived from five healthy persons (white bar) and derived from 19 patients with MDS (black bar) for 7 weeks, and LTC-IC frequency was determined (ai). CD34+ HSPC of six MDS patients were pairwise cultured on MSC derived from nine healthy controls (white bar) and derived from nine MDS patients (black bar) (aii). Means and s.e.m. are displayed. Unpaired Student’s t-test was used for experiments as shown in (ai) and the Wilcoxon signed rank test was used for paired experiments as shown in (aii). Asterisks display **P<0.01, ***P<0.001. Cell cycle analysis of CD 34+ HSPC patients were performed after 3 days of cultivation either in the presence or without MSC by Ki-67 and Hoechst 33 342 staining and flow cytometric analysis. (bi) shows representative plots of a healthy control (upper plot) and of a MDS patient (lower plot). (bii) shows cell cycle distribution of healthy CD34+ HSPC (n=26)cultivated without feeder layer support (control) on healthy MSC (n=14) as well as on MDS-derived MSC (n=26). (biii) shows cell cycle distribution of MDS CD34+ HSPC (n=26) cultivated without feeder layer support (control) on healthy MSC (n=26) as well as on MDS-derived MSC (n=21). MDS CD34+ cells are also given for early (<5% blasts) and advanced MDS (>5% blasts). In all experiments, white bars indicate G0-phase, black bars G1-phase and gray bars G2/S/M phases. Unpaired Student’s t-test was used to detect statistically significant differences. Asterisk displays *P<0.05.

MDS-derived MSC influence the cell cycle activity of CD34+ HSPC

In a next step we investigated the influence of MSC derived from healthy controls and MDS on the cell cycle state of CD34+ HSPC employing Ki-67 and Hoechst 33 342 staining. After 3 days of cultivation without MSC support a significantly larger proportion of CD34+ HSPC from patients with MDS were in G0-phase (61 vs 32%, P=0.028) and a lower fraction in G1- (25 vs 46%, P=0.026) and in S/G2/M-phase (10 vs 17%, P=0.11) in comparison with CD34+ HSPC from healthy controls (Figures 5bi–biii).

When CD34+ HSPC from healthy controls were cultured on healthy MSC, their distribution among the distinct cell cycle phases remained unchanged with the largest proportion of cells in the G1-phase. In contrast, after 3 days of coculture with MDS-derived MSC we observed a clear shift in the cell cycle activity of healthy CD34+ HSPC with the largest fraction of cells being now in G0-phase (P=0.07, Figure 5bii). This suggests that MDS-derived MSC direct the cell cycle behavior of CD34+ HSPC towards a resting phenotype comparable to the one observed in MDS CD34+ HSPC.

When CD34+ HSPC of MDS patients were cultured together with MDS-derived MSC their cell cycle state remained comparable to that in culture without feeder layer (Figure 5biii). By coculturing CD34+ HSPC from patients with various MDS subtypes on healthy MSC we already observed a reduction of the cell fraction in the G0-phase (P=0.09) and an increase of cells in the G1- (P=0.09) and in the S/G2/M-phase. We then separately analyzed CD34+ HSPC from patients with early (<5% BM blasts) or with advanced MDS (>5% BM blasts). This revealed that coculture with healthy MSC induced a significant shift in the cell cycle activity of CD34+ HSPC from early MDS. The greatest fraction of cells was now in G1-phase reflecting a phenotype previously observed only in healthy CD34+ HSPC. In contrast, no change of cell cycle distribution was found when CD34+ HSPC from advanced MDS patients were cultured together with healthy MSC (Figure 5biii). This implies that the influence on cell cycle activity of MDS-derived MSC is at least partially reversible in early MDS. In advanced MDS, the cell cycle behavior of CD34+ HSPC seems more autonomous and not reversible when transferred into a healthy microenvironment.


Ineffective hematopoiesis is a major characteristic of MDS, but comparatively few studies have addressed the pathophysiological contribution of the BM microenvironment, and in particular MSC.

We here show that MSC from patients with MDS were functionally impaired with regard to their growth capacities and osteogenic differentiation, which was also reflected by specific methylation changes in MDS-derived MSC. Furthermore, we detected an altered repertoire of cytokines and chemokines in the BM stroma of patients with MDS. Of particular interest, these alterations were present across all MDS subtypes and functionally translated into a significantly diminished ability of MDS-derived MSC to support CD34+ HSPC.

As first result we found that MDS-derived MSC have a reduced clonogenic ability and proliferated more slowly. This was associated with an altered morphology and disrupted colony architecture. Previous studies addressing the growth capacities of MDS-derived MSC had revealed controversial findings. Although most small studies also suggested growth deficits of MDS-derived MSC,15, 19, 20, 38 two other groups reported on normal growth and morphology of MDS-derived MSC.39, 40 This previous controversy was probably related to the small sample numbers, different methods of MSC cultivation and the fact that these reports mostly included only one method such as CFU-F to assess the growth potential. By assessing CFU-F, cumulative population doublings, doubling time and the number of passages in a large sample number, our data univocally demonstrate the impaired growth capacities of MSC across all common MDS subtypes. Corresponding to this, our data suggest for the first time that MDS-derived MSC are more senescent in comparison with MSC from an age-matched control group. As neither increased apoptosis nor cell cycle defects have been observed in MDS–MSC,15 increased senescence might reflect a mechanism responsible for their impaired proliferative capacity and altered morphology.41

Next, we were especially interested in the osteogenic differentiation potential of MDS-derived MSC. Previous studies had studied osteogenic differentiation only in a limited number of patients. These studies generally aimed to demonstrate the presence of osteogenic differentiation to fulfill the definition criteria of MSC and therefore mostly used only one cytochemical staining. This did not permit an exact quantification of the osteogenic differentiation.15, 20, 38, 39 In contrast, our analysis revealed that osteogenic differentiation potential of MSC was significantly reduced, affecting all MDS subtypes. This was not only indicated by three different cytochemical stainings, but also consistently by reduced mRNA expression levels of the osteoblastic transcription factor Osterix and the bone formation marker OC. In agreement with this notion, we also detected decreased OC concentrations in the serum of MDS patients. Furthermore, the methylation signature of MDS-derived MSC included a strong and consistent hypermethylation of the T-box transcription factor TBX15, which resulted in reduced mRNA expression of TBX15. Expression of TBX15 in mesenchymal precursor cells has been shown to be required for normal murine bone development,42 which suggests that hypermethylation-induced silencing of TBX15 could be a causal factor for the reduced osteogenic differentiation of MDS-derived MSC. In this context, it is also worth noting that Mellibovsky et al.43 have previously reported on an adynamic bone remodeling associated with decreased mineral deposition in patients with MDS. Lastly, Raaijmakers et al.11 have recently shown that deletion of Dicer1, an RNAse III endonuclease essential for microRNA biogenesis, leads to an impaired osteogenic differentiation of MSC and induces an MDS phenotype in mice . Interestingly, we also found downregulation of Dicer1 in MDS-derived MSC in agreement with earlier findings.44

In the light of these findings and the pivotal role of mesenchymal osteolineage cells in regulating hematopoiesis,9 it did not come as a surprise that the impaired growth and osteogenic differentiation potential of MDS-derived MSC translated functionally into a significantly reduced ability to support CD34+ HSPC in LTC-IC assays. Previous studies using LTC-IC assays to investigate the hematopoietic support by MDS-derived MSC had revealed controversial results. However, a direct comparison with our results is hampered by different experimental designs. In contrast to our approach, where we used BM-derived CD34+ HSPC, the other investigators used either cord blood-derived CD34+ cells, cytokine-expanded CD34+ cells or mononuclear cells. Furthermore, in the study of Soenen-Cornu et al.,18, 21, 22, 45, 46 the murine MS-5 cell layer was used as control for MDS-derived MSC. We also observed a diminished cell cycle activity of CD34+ HSPC when cultured on MDS–MSC. Again, this strengthens the conclusion that impaired stromal support contributes to ineffective hematopoiesis and peripheral cytopenias observed in patients with MDS.

As a potential mechanistic link between the phenotypic alterations of MDS-derived MSC and their insufficient hematopoietic support reported here, we found altered expression levels of Osteopontin, Angiopoietin-1, Jagged1 and Kit-ligand in MDS-derived MSC. These molecules are involved in the regulation of hematopoiesis,9 suggesting that their deregulation contributes to the insufficient hematopoietic support by MDS-derived MSC. In detail, we found a diminished expression of Kit-ligand, which was previously reported to be reduced in the BM plasma of MDS patients.47 Mice lacking Kit-ligand expression on BM stromal cells present with macrocytic anemia,48 which is also a common symptom in MDS. Similarly, overexpression of Jagged1 on MDS-derived MSC, as shown here, has been reported to result in a reduced rate of cobblestone-forming cells in MDS stromal cocultures.38

Besides deregulation of established cytokines, we found the expression levels of several chemokines such as CXCL1, CCL2, CXCL14, CX3CL1, CXCL7, midkine and CCL26 to be altered in MDS-derived MSC. None of these have been previously mentioned in the context of MDS pathogenesis. As chemokines have an important role in the trafficking of immune cells, it is conceivable that the altered immunoregulatory capacities of MDS-derived MSC previously reported by Zhao et al.40 are related to the chemokine imbalances that we observed .

Given the profound alterations of MDS-derived MSC, the question arises whether these cells carry primary defects contributing to hematopoietic failure, or whether the observed changes represent secondary adaptations. Although the reduced Dicer1 expression reported here might be comparable to Dicer1 deletion in mice where a primary stromal defect caused MDS,11 we favor the latter theory based on the following findings:

In MDS, the clonal HSPC expand at the expense of normal HSPC independently from the disease-state and become the dominant cell population in the BM,3, 5 probably allowing them to dominate the stromal cells as well. The alterations of MSC reported here were consistently found across all MDS subtypes without particular relationship to a distinct MDS subtype. This was also true for the DNA methylation changes in MDS-derived MSC, suggesting that the majority of changes might represent a more general reaction to the clonal HSPC irrespective of MDS subtype. This interpretation is further supported by recent findings from our group as well as other research groups showing deficits of osteogenic differentiation and growth in AML and multiple myeloma, comparable to those we observed in MDS-derived MSC. In these studies, the invading myeloid blasts or plasma cells suppressed osteoblastic/MSCs, thereby contributing to hematopoietic insufficiency.24, 36, 49 In multiple myeloma, the decreased stem cell supporting capacity was related to the overexpression of TGF-beta in the BM microenvironment.24 Zhou et al.50, 51 showed that overactivation of TGF-beta signaling is also important in MDS resulting in erythroid impairment, thus providing a rationale for pharmacological inhibition of TGF-beta signaling as treatment of anemia in patients with MDS. Interpreting MDS–MSC alterations as secondary adaptations is also strongly supported by the clinical observation that the eradication of not only AML or myeloma cells but also MDS precursors from the BM through chemotherapy or novel agents leads to the restoration of hematopoiesis. This would not be expected if the deranged microenvironment were the primary and dominant cause of hematopoietic failure. The same reasoning applies to the situation after allogeneic transplantation, where the deranged BM stroma of the host can usually be ‘reprogrammed’ rather quickly to become supportive of the incoming healthy donor cells. Correspondingly, we observed that host-derived MSC of MDS patients, who were in complete hematological remission after allogeneic transplantation, regained a significant greater ability (5.9-fold) to support healthy CD34+ HSPC in LTC-IC assays compared with MSC derived from untreated MDS patients (data not shown).

In summary, our comprehensive analysis showed that MSC from all MDS subtypes are structurally, epigenetically and functionally altered, thus causing impaired stromal support of HSPC. This seems to be an important mechanism in MDS pathophysiology, which might be targeted by novel pharmacological approaches.


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This work was supported by the Leukämie Lymphom Liga e. V., Duesseldorf, Germany. We thank Annemarie Koch, Anke Boeckmann and Tanja Musch for their excellent technical assistance. We thank Johannes C. Fischer and Katharina Raba for their substantial technical support with the FACS analyses. SÖ is a PhD student of the HBIGS graduate school and supported by the PhD program ‘Disease Models and Drugs’ between the University of Heidelberg and the Mannheim University of Applied Sciences.

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Correspondence to T Schroeder.

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The authors declare no conflict of interest.

Additional information

Parts of this study have been presented at the 53rd American Society of Hematology (ASH) Annual Meeting, San Diego, CA, December 10–13, 2011 and at the Annual Meeting of the German-Austrian-Suisse Society of Hematology and Oncology (DGHO), October 19–23, 2012.

Author contributions

Conception and design: TS, RH, FL, SG. Provision of patients samples: UG, CZ, AK, GK, RF, TS. Experiments, collection and assembly of data: SG, TS, JF, R-PC, IB, SÖ. Data analysis and interpretation: TS, RH, SG, UG, NG, DH, FL, BB, JF, R-PC. Manuscript writing: TS, SG, RH, FL, BB. Final approval of the manuscript was given by all authors.

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Geyh, S., Öz, S., Cadeddu, R. et al. Insufficient stromal support in MDS results from molecular and functional deficits of mesenchymal stromal cells. Leukemia 27, 1841–1851 (2013) doi:10.1038/leu.2013.193

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  • MDS
  • MSC
  • methylation
  • bone marrow microenvironment
  • hematopoietic insufficiency
  • CD34+ HSPC

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