Abstract
Mesenchymal stem cells (MSCs) have broad-based therapeutic potential in regenerative medicine. However, a major barrier to their clinical utility is that MSCs from different tissues are highly variable in their regenerative properties. In this study, we defined the molecular and phenotypic identities of different MSC populations from different osseous tissue sites of different patients, and additionally, determined their respective regenerative properties. MSCs from 6 patients were isolated from either bone marrow of the iliac crest (BMSCs) or alveolar bone tissue (aBMSCs) and flow cytometry revealed that regardless of the tissue source, MSC immunotypes had the same expression of MSC markers CD73, CD90, and CD105. However, transcriptomic analyses revealed 589 genes differentially expressed (DE) between BMSCs and aBMSCs, including 8-fold higher levels of bone morphogenetic protein 4 (BMP-4) in aBMSCs. In striking contrast, gene expression of MSCs derived from the same tissue, but between different patients (i.e., BMSCs to BMSCs, aBMSCs to aBMSCs), showed only 38 DE BMSC genes and 51 DE aBMSC genes. A protein array showed that aBMSC and BMSC produced equivalent levels of angiogenic cytokines, however, when placed in angiogenesis model systems, aBMSCs induced significantly more capillaries in vitro and in vivo. Finally, cell transplantation of MSCS into osseous defects showed that the bone regenerative capacity of aBMSCs was significantly greater than that of BMSCs. This study is the first to link the molecular, phenotypic, and regenerative properties of different MSCs from different patients and provides novel insights toward MSC differences based on the osseous tissue origin.
Keywords: Mesenchymal stem cells, Differential gene expression, RNA sequencing, Angiogenesis, Bone regeneration
1. Introduction
Adult-derived mesenchymal stem cells (MSCs) have widespread potential in cell-based therapies due to their regenerative and trophic properties. Historically, MSCs have been characterized by their multipotency in culture and their immunotypes. MSCs were initially isolated from the bone marrow based on their ability to adhere to plastic culture dishes and first described as non-hematopoietic precursor cells with fibroblast morphology and clonogenicity (Friedenstein, et al. 1970; Friedenstein, et al. 1978). Since this time, it has been recognized that MSC populations can be isolated from a variety of tissues including adipose, muscle, tendon, peripheral blood, umbilical cord, oral soft tissue, alveolar bone tissue, and tooth tissues (Akintoye, et al. 2006; Crisan, et al. 2008; Gronthos, et al. 2000; Mason, et al. 2014; Matsubara, et al. 2005; Seo, et al. 2004; Zhang, et al. 2012). Presently, there are three criteria used to define adult MSCs, regardless of their source of origin: 1) adherence to plastic; 2) positive expression of CD73, CD90, and CD105, and negative expression of CD45, CD34, CD14 or CD11b, CD79α or CD19 and HLA-DR surface molecules; and 3) the ability to differentiate into osteoblasts, adipocytes and chondroblasts in vitro (Dominici, et al. 2006).
Despite being similar in phenotype and immunotype, the trophic and regenerative properties of different MSC populations have been shown to be highly variable in different systems (Caplan 2015; Chen, et al. 2004; Hendrich, et al. 2009; Kaigler, et al. 2015; Lloyd, et al. 2017; Nakajima, et al. 2018; Quarto, et al. 2001; Rodrigo, et al. 2013). Because there have been limited studies comparing the regenerative properties of MSCs from different tissues, their differences are ill-defined and poorly understood. A deeper understanding of these underlying differences has significant therapeutic implications and holds broad-based impact.
We previously demonstrated a standardized method for isolation and expansion of MSCs from alveolar bone tissue (aBMSCs) (Mason, et al. 2014) and the phenotypic properties of aBMSCs were comparable to those of MSCs derived from bone marrow of the iliac crest (BMSCs) (Cao, et al. 2020). The objectives of the present study were to define the molecular, trophic, and regenerative properties of human BMSCs and aBMSCs and compare them directly to one another. The hypothesis underlying the study was that the molecular identities of MSC populations from different bone tissue sources are variable and further, that this variability provides prospective insight toward the regenerative properties of these cells.
2. Materials and Methods
2.1. Cells
BMSCs (n = 4) and aBMSCs (n = 4) were isolated from 6 human subjects’ samples and cultured as previously described (Cao, et al. 2020; Mason, et al. 2014) in accordance with our Institutional Review Board approved protocol (HUM00064770). Two of the human subjects donated both iliac crest marrow aspirate for isolation of BMSCs and alveolar bone tissues for aBMSCs, whereas 2 additional iliac marrow aspirate samples and 2 additional alveolar bone sample were obtained from the other 4 different human subjects. Both BMSCs and aBMSCs were maintained in the same fashion using the same growth media, i.e., the nucleosides-containing Minimum Essential Media α (MEM α; Thermo Fisher Scientific, Waltham, MA) supplemented with 15% fetal bovine serum (FBS; MilliporeSigma, Burlington, MA), 0.1 mM L-ascorbic acid-2-phosphate (MilliporeSigma), and 25 μg/mL gentamicin (Thermo Fisher Scientific). Primary human umbilical vein endothelial cells (HUVECs) were obtained from American Type Culture Collection (ATCC, Manassas, VA), and cultured in Endothelial Growth Medium-2 (EGM-2; Lonza, Walkersville, MD).
2.2. Immunotyping−Flow cytometry
Flow cytometry to determine the expression levels of the cell surface markers CD90, CD73, CD105, and Stro-1 was performed. BMSCs and aBMSCs were harvested from T150 flasks, washed and aliquoted equally into tubes. Cells were first incubated with a blocking solution containing an anti-CD16/CD32 antibody (Biolegend, San Diego, CA) at 4°C for 10 minutes followed by washing. Cells were then incubated with the specific antibodies conjugated with fluorochromes (Biolegend) at 4°C for 30 minutes. After washing, these cells were analyzed on a MoFlo flow cytometer (Beckman Coulter, Brea, CA).
2.3. Angiogenesis protein array
BMSCs and aBMSCs were washed with serum-free MEM α, followed by the addition of 5 ml serum-free MEM α before the cells were incubated at 37°C for 24 hours. After incubation the cells and conditioned media were harvested. Relative levels of 55 angiogenesis related proteins were detected in this conditioned media using the Human Angiogenesis Antibody Array (R&D Systems, Minneapolis, MN; Cat. #ARY007). The DNA concentration was measured from the harvested cells. Array membranes were analyzed by quantifying the mean spot pixel densities using the freeware Image J software (NIH http://rsb.info.nih.gov/ij). Samples were standardized using DNA content.
2.4. 3-dimensional microcarrier angiogenesis assay
The angiogenic potential of BMSCs and aBMSCs was assessed using a microcarrier-based angiogenesis assay as described previously (Ghajar, et al. 2008). Primary endothelial cells (HUVECs) cultured in EGM-2 were trypsinized at 90% confluence, and counted. Four million HUVECs were combined with 10,000 Cytodex 3 microcarrier beads (MilliporeSigma) in 5 ml EGM-2 and pipetted into an upright T-25 culture flask. The suspension was cultured for four hours in a 37°C, 5% CO2 incubator and agitated gently every 30 minutes. Following cell adhesion, the suspension was removed and an additional 5ml EGM-2 was added. The suspension was then pipetted into a new T-25 and cultured overnight in a normal cell culture position.
Fibrin gels were prepared the following day. Bovine fibrinogen (MilliporeSigma) was dissolved in serum-free EGM-2 and sterile filtered. Gels were created by combining 2.5mg/ml fibrinogen solution containing approximately 100 HUVEC-coated microcarriers with bovine thrombin (MilliporeSigma), and pipetting the resulting solution into a 24-well plate. The beads were allowed to settle for 5 minutes, and then to finish gelation, the solution was incubated for 25 minutes at 37°C. During incubation, BMSCs and aBMSCs were trypsinized and counted. Following gelation, 25,000 cells of the appropriate type were plated on top of each gel and co-cultures were incubated in a 37°C, 5% CO2 incubator with media changes every other day. The total network lengths of the capillary-like networks were quantified in the 3D co-culture model of angiogenesis as previously described (Ghajar, et al. 2010). Briefly, capillary networks formed by HUVECs in the presence of either BMSCs or aBMSCs were imaged at 7 and 14 days using phase contrast microscopy (Olympus IX81, Olympus America, Center Valley, PA). With each condition, a total of 10 beads were imaged at low magnification (4×) and tracked longitudinally across both time points. Total vessel network lengths per bead were measured using the Angiogenesis Tube Formation module within MetaMorph Premier Software (Molecular Devices, Sunnyvale, CA).
2.5. In vivo angiogenesis model−subcutaneous mouse implants
The blood vessel forming capacity of BMSC and aBMSC populations was evaluated qualitatively in a subcutaneous mouse model, as previously described (Kaigler, et al. 2005; Nor, et al. 2001) in accordance with our Institutional Animal Care and Use Committee (IACUC) approved protocol (PRO00008804). Briefly, gelatin sponges (Gelfoam®, Pfizer, New York, NY) containing either BMSCs or aBMSCs were implanted into subcutaneous pockets on the dorsal region of 7- to 9-wk-old male severe combined immunodeficient (SCID) mice (cb17/SCID) (Taconic Laboratories, Germantown, NY) (n=5). Animals were first anesthetized with an intraperitoneal injection of 1 ml of ketamine (100 mg/ml) and 0.15 ml of xylazine (20 mg/ml) and a small (about 1.5-cm) longitudinal incision was then made into the skin. Scissors were inserted under the skin to perform blunt dissection of the subcutaneous fascial tissue and to create subcutaneous pockets into which the implants could be placed. Following placement of the implants, the incisions were closed with interrupted sutures and 4 weeks post-operatively, animals were sacrificed and implants were analyzed for blood vessel formation, as previously described (Yeasmin, et al. 2014). Briefly, blood vessels were quantified in each histologic section based on the presence of endothelial cells lining well-defined capillary lumens containing red blood cells. The number of microvessels was counted blindly in 10 random fields per implant at × 100 magnification. Vessel numbers were then reported as an average of the number vessels per high powered field.
2.6. In vivo bone regeneration model−rat calvarial defect
The ability of BMSCs and aBMSCs to regenerate bone was evaluated in vivo in a rat calvarial defect model as we have previously described (Kaigler, et. al. 2006) in accordance with our IACUC approved protocol (PRO00009333). Briefly, a mid-longitudinal incision was made on the dorsal surface of the cranium of RNU Nude rats (Charles River Laboratories, Mattawan, MI) prior to exposing the calvaria (n=5). A 5 mm diameter trephine bur was then used to create two circular osseous defects in the rat cranium on both sides of the sagittal suture. The full thickness of the cranial bone was removed, and collagen scaffolds with BMSCs or aBMSCs were placed in the defects. The areas were sutured with 4–0 Silk sutures and the sites allowed to heal for 8 weeks. Following 8 weeks, calvaria samples and 3-dimensional micro-CT images of the non-decalcified calvariae were captured and 3-dimensionally reconstructed. Bone mineral density (BMD) and bone volume fraction (BVF) were calculated and following this imaging, samples were decalcified and prepared histologically for standard hematoxylin and eosin (H&E) staining after which bone histomorphometry was performed as previously described.
2.7. Transcriptome analysis−RNA sequencing
Total RNA from BMSCs and aBMSCs were isolated using the RNeasy Plus kit (Qiagen, Hilden, Germany; Cat. #74134). The University of Michigan Sequencing Core prepared the cDNA library and performed HiSeq 50 Cycle Single Read sequencing on an Illumina High-Seq Platform.
Data analysis was provided by the UM Bioinformatics Core. FastQC (version 0.10.1) was utilized for pre-alignment analysis to determine the quality of the reads (low quality scores, over-represented sequences, inappropriate GC content, etc.) and for post-alignment to ensure that only high quality data would be input to expression quantitation and differential expression analysis. The Tuxedo Suite Tools were used for alignments of reads (TopHat version 2.0.9, Bowtie version 2.1.0). Cufflinks/CuffDiff version 2.1.1 was used for the expression, quantitation, normalization and differential expression analysis. Diagnostic plots were generated using the CummeRbund R package. NCBI Entrez GeneIDs was used for gene and isoform annotation and text descriptions. Differentially expressed genes were further annotated using Gene Ontology (GO). DAVID version 6.7 was used for enrichment analysis of the set of differentially expressed genes to identify significantly enriched functional categories.
3. Results
3.1. Immunophenotypes of BMSCs and aBMSCs
Consistent with established standards for characterization of MSCs (Dominici, et al. 2006), we have previously reported that aBMSCs consistently express high levels of CD73, CD90, CD105 while being negative for CD11b, CD19, CD34, and CD45 (Mason, et al. 2014). As such, we isolated aBMSCs from two individuals and from the same two individuals, isolated BMSCs. Following expansion of BMSCs and aBMSCs, flow cytometry was performed to establish their immunophenotype. At 15 population doublings (PDs), high levels (> 95%) of CD73+, CD90+, and CD105+ cells were expressed in BMSCs and aBMSCs from both individuals (Fig. 1a and b). Additionally, despite being derived from two different tissue sources, there was no difference in the expression levels of these markers between MSCs from these two tissues (Fig. 1c and d). Immunophenotypes of BMSC and aBMSC populations from four additional patients were consistent with those of these two individuals and again, showed no difference between the two cell types (Supplementary Fig. 1).
Figure 1.
Cell surface marker expression of aBMSCs and BMSCs. (a and b) Mesenchymal stem cell marker expression of CD73, CD90, and CD105 were compared between different BMSC populations (a) and between different aBMSC populations (b), and populations from both subjects yielded high levels of expression of these markers. (c and d) MSC surface marker expression was then compared between BMSCs and aBMSCs from each of these subjects, i.e., Patient 1 (c) and Patient 2 (d), with no difference in expression between the two cell populations.
3.2. Trophic properties of BMSCs and aBMSCs
In addition to demonstrating multipotency, MSCs have been evaluated extensively in preclinical and clinical studies for their capacity to induce angiogenesis and stimulate blood vessel formation, namely through their production of trophic factors. We next evaluated the trophic factors produced by BMSCs and aBMSCs using an angiogenesis proteome profiler array to assess secreted molecules from both BMSCs and aBMSCs of the same individual. Of the 55 angiogenesis related proteins screened, the array showed that BMSCs and aBMSCs produced varying levels of 20 of these analytes (Fig. 2a and b). In both BMSCs and aBMSCs, the angiogenic proteins endothelin, insulin-like growth factor binding protein 3 (IGFB-3), pentraxin 3, plasminogen activator inhibitor-1/serpin E1 (SE1), pigment epithelium-derived factor/serpin F1 (SF1), thrombospondin-1, tissue inhibitor of matrix metalloproteinase 1 (TIMP-1), and vascular endothelial growth factor (VEGF) were most highly produced. In comparing angiogenic protein production between BMSCs and aBMSCs, quantitative analyses showed that BMSCs secreted higher concentrations of angiogenin, C-X-C motif chemokine ligand 16 (CXCL16), angiopoietin-2, endocrine gland-derived VEGF (EG-VEGF), insulin-like growth factor binding protein 2 and 3 (IGFBP-2 and IGFBP-3), matrix metalloproteinase 9 (MMP-9), and placental growth factor (PlGF) (Fig. 2c).
Figure 2.
Angiogenic factors produced by BMSCs vs. aBMSCs. (a and b) Representative results of anigogenic proteome profiler arrays probed with serum-free conditioned medium (CM) from BMSCs (a) and aBMSCs (b); boxes indicate individual factors in duplicate. (c) Quantitative analysis of angiogenic factors and chemokines in CM from BMSCs and aBMSCs (normalized to DNA content). * denotes > two-fold difference. R, reference spots; AN, angiogenin; AP1, angiopoietin-1; AP2, angiopoietin-2; AR, artemin; CX, CXCL16; DP, DPPIV (dipeptidyl peptidase 4); EV, EG-VEGF; EN, endothelin; I1, IGFBP-1 (insulin-like growth factor binding protein 1); I2, IGFBP-2; I3, IGFBP-3; M9, MMP-9; P3, pentraxin 3; Pe, persephin; Pl, PlGF; SE1, serpin E1; SF1, serpin F1; T1, TIMP-1; Th1, thrombospondin-1; Up, uPA (urokinase-type plasminogen activator); V, VEGF.
To evaluate the functional effect of the trophic properties of BMSCs and aBMSCs, a 3D angiogenesis system was used whereby endothelial cells (ECs) differentiate in an extracellular matrix into capillary tubes under the influence of exogenous angiogenic factors. BMSCs and aBMSCs were co-cultured with ECs (without direct contact) and over the course of two weeks, and EC differentiation into tubular vessels was measured. After one week, vascular network formation appeared to be similar under conditions for which ECs were cultured in the presence of either BMSC or aBMSCs (Fig. 3a), while no tubular structures were identified in conditions with ECs only. Quantitative analysis of the angiogenic response was determined by measuring the total network length of the capillary-like networks. Following two weeks of culture, ECs grown in the presence of aBMSCs exhibited a more robust angiogenic response than those grown in the presence of BMSCs (Fig. 3b). Quantitative analysis of the total network length confirmed that there was a significant (p = 0.03) two-fold increase in the vessel network length of ECs cultured in the presence of aBMSCs relative to those in the presence of BMSCs.
Figure 3.
BMSC and aBMSC induction of endothelial cell (EC) vessel formation. (a-f) Photomicrographs (10× magnification) of EC-coated microbeads at 7 (a-c) and 14 days of culture (d-f) within fibrin gels containing BMSCs (a and d), aBMSCs (b and e), or no MSCs (c and f). Images show differentiating ECs forming vessels elongating from microbeads (yellow arrows indicate vessels). Scale bar depicts 200 µm. (g) Quantification of the total vessel network length formed by ECs when cultured in the presence of BMSCs, aBMSCs, or no other interstitial cell type shows that by 14 days, aBMSCs induce formation of a more robust vascular network. * denotes p < 0.05.
The capacity of BMSCs and aBMSCs to form capillary networks in vivo was also evaluated following subcutaneous transplantation in SCID mice. High and lower power magnification shows that four weeks following transplantation, relative to transplantation of BMSCs, aBMSCs induced the formation of a greater number of vessels, and these vessels appeared much larger in diameter (Fig. 4a-f). Quantitative analysis of total vessels formed revealed significantly (p < 0.05) more vessels in the condition in which aBMSCs were transplanted (Fig. 4g).
Figure 4.
In vivo induction of de novo blood vessel formation by BMSCs and aBMSCs. (a-f) Low (40×; a-c) and high (100×; d-f) magnification photomicrographs of representative sections of tissue sections of scaffolds without cells (a and d) and those containing BMSCs (b and e) showed relatively low densities of blood vessels at 4 weeks compared to those formed in scaffolds containing aBMSCs (c and f). (g) Quantitative analysis of vessel formation showed that scaffolds containing aBMSCs had significantly greater number of blood vessels formed compared to conditions with no cells or BMSCs. * p < 0.05.
3.3, In vivo bone regeneration following transplantation of BMSCs and aBMSC
Because BMSCs and aBMSCs were harvested from different bone tissues within the same individual, we wanted to determine if there were differences in their ability to regenerate bone in an orthotopic bone regeneration model. Using a Type I collagen based scaffold material (Colla-tape, Zimmer Biomet; Warsaw, IN) as a cell carrier, BMSCs and aBMSCs were transplanted in bone defects created in the calvaria of nude rats. After eight weeks, the bone regenerative capacities of BMSCs and aBMSCs were evaluated. Following harvest of calvaria, micro-CT was used to generate 3D reconstructions of the regenerated bone within the defect. These images revealed that the greatest coverage of the defects with mineralized tissue was achieved in the condition where aBMSCs were transplanted (Fig. 5b). Quantitative micro-CT analyses of the regenerated defects confirmed this qualitative assessment and showed that the bone volume fraction (BVF) of the area treated with aBMSCs was significantly (p < 0.05) higher (BVF = 0.21) than that of the defect areas treated with BMSCs (0.14) (Fig. 5c). In comparing the bone mineral density (BMD) of this regenerated tissue, the BMD of tissue regenerated by aBMSCs (1,213.1 mg/cc) was almost twice (p < 0.01) that of the regenerated tissue derived from BMSCs (681.8 mg/cc) (Fig. 5d). Histological evaluation of these tissues (H&E) showed that these regenerated tissues had the morphology indicative of early woven bone formation and areas of more mature, dense lamellar bone (Fig. 5e-g).
Figure 5.
Eight week BMSC- and aBMSC-induced bone regeneration of calvarial defects. (a and b) 3-D reconstructed micro-CT images of calvarial defects which received the following treatments at the time of defect creation: no treatment (empty; a left), collagen scaffold only (a right), scaffold loaded with BMSCs (b left), and scaffold loaded with aBMSCs (b right). (c) Bone volume fraction (BVF) of defects treated with BMSCs and aBMSCs is significantly greater than BVF of non-treated defects or defects only treated with the scaffold; however, BVF of defects treated with aBMSCs is significantly greater than defects treated with BMSCs. (d) Bone mineral density (BMD) of defects treated with BMSCs and aBMSCs is significantly greater than BMD of non-treated defects or defects treated with the scaffold only. (e-g) Representative H&E histological images of calvaria showing defects treated with BMSCs (e left and f) and aBMSCs (e right and g) and the tissue morphology of regenerated bone (green arrows show new bone; red arrows show native bone; black arrows show fibrous tissue).
3.4. Molecular identities of BMSCs and aBMSCs
Next generation sequencing was used to determine the molecular identities of BMSCs and aBMSCs. Whole transcriptome analysis was performed on BMSCs and aBMSCs from two subjects and differential gene expression determined between these MSCs from different tissues. In contrast to these cell populations having the same cell surface marker expression, there were 564 differentially expressed (DE) genes between them (Fig. 6a). Among these genes, 237 were more highly expressed in BMSCs while 327 were more highly expressed in aBMSCs (See Supplementary Table 1 and Supplementary Table 2 for the complete list of DE genes). Strikingly, when comparing the gene expression of BMSCs between the different subjects, it was observed that there were only 16 DE genes (Fig. 6b). Similarly, in comparing aBMSCs from the two subjects, of the > 23,000 genes probed there were only a very small number (18) which were DE (Fig. 6b). The transcriptomes of aBMSC samples from two additional subjects were determined as were the transcriptomes of two additional BMSC samples; however, unlike the first two samples, these BMSC and aBMSC samples were not derived from the same subjects. Transcriptome analyses of these additional samples revealed that between BMSCs and aBMSCs, there were 589 DE genes (Fig. 6c). Taken together, the key novel finding is that MSCs from two different osseous tissue sources within the same subject exhibit much greater difference in gene expression relative to MSCs from the same tissue source taken from different subjects.
Figure 6.
Differential gene expression. (a-c) Numbers of differentially expressed genes between BMSCs and aBMSCs (a and c) or between patients (b). (a) While there are many differentially expressed genes between BMSCs and aBMSCs from the same subject (n = 2), there is strikingly less (b) differential gene expression between aBMSC from different subjects and between BMSC from different subjects. (c) RNA sequencing of additional samples (n = 4 in total for either BMSCs or aBMSCs) confirms that gene expression of MSC populations from the same tissue type are strikingly similar from one individual to another.
Further analysis of the BMSC and aBMSC transcriptomes was performed in order to establish a classification system according to the biological process or role in which the gene plays a predominant role. The following groups were used: immunity, immune response, inflammatory response and chemotaxis (Table 1), angiogenesis (Table 2), and osteogenesis/osteoblast differentiation (Table 3). Of these categories, angiogenesis and osteogenesis are of particular interest in therapeutic applications of bone regeneration (Table 2 and Table 3). Of the 38 DE angiogenic genes, 21 were more highly expressed in aBMSCs. Of the 23 DE osteogenic genes, 9 were more highly expressed in aBMSCs and 14 more highly expressed in BMSCs. Significant to note is that one of the more potent bone morphogens, BMP-4, is more highly expressed (8-fold) in aBMSCs. qPCR was performed to confirm these findings (Supplementary Fig. 2).
Table 1.
Differential expression of genes related to immunity, immune response, inflammatory response and chemotaxis between BMSCs and aBMSCs.
| Gene | Description | Fold | |
|---|---|---|---|
| Genes expressed more in BMSCs | C1QA | complement component 1, q subcomponent, A chain | * |
| C1QB | complement component 1, q subcomponent, B chain | * | |
| C1QC | complement component 1, q subcomponent, C chain | * | |
| S100A8 | S100 calcium binding protein A8 | * | |
| S100A9 | S100 calcium binding protein A9 | * | |
| CHI3L1 | chitinase 3 like 1 | 236.216 | |
| ADORA1 | adenosine A1 receptor | 34.184 | |
| GGT5 | gamma-glutamyltransferase 5 | 27.068 | |
| BST2 | bone marrow stromal cell antigen 1 | 26.885 | |
| HLA-DRB1 | major histocompatibility complex, class II, DR beta 1 | 22.517 | |
| HLA-DRA | major histocompatibility complex, class II, DR alpha | 19.978 | |
| NLRP3 | NLR family, pyrin domain containing 3 | 13.841 | |
| HLA-DRB5 | major histocompatibility complex, class II, DR beta 5 | 12.644 | |
| SBSPON | somatomedin B and thrombospondin type 1 domain containing | 12.484 | |
| SPP1 | secreted phosphoprotein 1 | 12.378 | |
| FPR1 | formyl peptide receptor 1 | 11.190 | |
| CXCL6 | chemokine (C-X-C motif) ligand 6 | 10.317 | |
| NGFR | nerve growth factor receptor | 9.256 | |
| CFI | complement factor I | 8.042 | |
| CD74 | CD74 molecule, major histocompatibility complex, class II invariant chain | 7.654 | |
| CSF1R | colony stimulating factor 1 receptor | 7.459 | |
| TINAGL1 | tubulointerstitial nephritis antigen-like 1 | 6.651 | |
| ITGB2 | integrin subunit beta 2 | 5.921 | |
| TNFRSF21 | tumor necrosis factor receptor superfamily member 21 | 5.889 | |
| SUSD4 | sushi domain containing 4 | 5.538 | |
| S1PR1 | sphingosine-1-phosphate receptor 1 | 5.510 | |
| LSP1 | lymphocyte-specific protein 1 | 5.396 | |
| C3AR1 | complement component 3a receptor 1 | 5.307 | |
| SPHK1 | sphingosine kinase 1 | 5.110 | |
| SEMA7A | semaphorin 7A, GPI membrane anchor (John Milton Hagen blood group) | 4.763 | |
| CCL26 | chemokine (C-C motif) ligand 26 | 4.175 | |
| CIITA | class II, major histocompatibility complex, transactivator | 4.131 | |
| ADARB1 | adenosine deaminase, RNA-specific, B1 | 3.773 | |
| CHST1 | carbohydrate (keratan sulfate Gal-6) sulfotransferase 1 | 3.583 | |
| EPHA2 | EPH receptor A2 | 3.479 | |
| TSPAN2 | tetraspanin 2 | 3.474 | |
| Genes expressed more in aBMSCs | MASP1 | mannan-binding lectin serine peptidase 1 (C4/C2 activating component of Ra-reactive factor) | 86.109 |
| SCN9A | sodium channel, voltage gated, type IX alpha subunit | 26.301 | |
| PTGS1 | prostaglandin-endoperoxide synthase 1 (prostaglandin G/H synthase and cyclooxygenase) | 15.693 | |
| MBP | myelin basic protein | 15.092 | |
| VNN1 | vanin 1 | 14.316 | |
| SECTM1 | secreted and transmembrane 1 | 13.400 | |
| SPON2 | spondin 2, extracellular matrix protein | 13.382 | |
| CXCL14 | chemokine (C-X-C motif) ligand 14 | 9.860 | |
| OAS1 | 2’−5’-oligoadenylate synthetase 1 | 8.443 | |
| F2R | coagulation factor II (thrombin) receptor | 7.797 | |
| CD55 | CD55 molecule, decay accelerating factor for complement (Cromer blood group) | 6.661 | |
| TEC | tec protein tyrosine kinase | 6.188 | |
| IFIT2 | interferon induced protein with tetratricopeptide repeats 2 | 6.093 | |
| P2RX7 | purinergic receptor P2X, ligand gated ion channel, 7 | 6.048 | |
| CCRL2 | chemokine (C-C motif) receptor-like 2 | 5.153 | |
| IL12A | interleukin 12A | 4.928 | |
| NLRP10 | NLR family, pyrin domain containing 10 | 4.802 | |
| AMOT | angiomotin | 4.729 | |
| PAG1 | phosphoprotein membrane anchor with glycosphingolipid microdomains 1 | 4.329 | |
| IL7R | interleukin 7 receptor | 4.032 | |
| TNFSF15 | tumor necrosis factor superfamily member 15 | 3.923 | |
| IFIT3 | interferon induced protein with tetratricopeptide repeats 3 | 3.782 | |
| APOL3 | apolipoprotein L3 | 3.716 | |
| IFIH1 | interferon induced, with helicase C domain 1 | 3.509 | |
| CFD | complement factor D (adipsin) | 3.484 | |
| IFIT1 | interferon induced protein with tetratricopeptide repeats 1 | 3.467 | |
| PRKCE | protein kinase C, epsilon | 3.437 | |
| CFB | complement factor B | 3.405 | |
| PTGER4 | prostaglandin E receptor 4 | 3.346 | |
| C1S | complement component 1, s subcomponent | 3.273 | |
| ERAP2 | endoplasmic reticulum aminopeptidase 2 | 3.119 | |
| ROBO1 | roundabout guidance receptor 1 | 3.041 | |
| C1R | complement component 1, r subcomponent | 3.033 | |
| TGFBR3 | transforming growth factor beta receptor III | 3.032 |
No detectable expression in aBMSCs
Table 2.
Differential expression of angiogenic genes between BMSCs and aBMSCs.
| Gene | Description | Fold | |
|---|---|---|---|
| Genes expressed more in BMSCs | HOXA3 | homeobox A3 | * |
| HOXB3 | homeobox B3 | 74.496 | |
| PGF | placental growth factor | 66.280 | |
| MEOX2 | mesenchyme homeobox 2 | 25.901 | |
| MCAM | melanoma cell adhesion molecule | 11.024 | |
| S1PR1 | sphingosine-1-phosphate receptor 1 | 5.510 | |
| FGFR2 | fibroblast growth factor receptor 2 | 4.992 | |
| NRXN3 | neurexin 3 | 4.940 | |
| NRP2 | neuropilin 2 | 4.442 | |
| EGF | epidermal growth factor | 4.367 | |
| MEIS1 | Meis homeobox 1 | 4.260 | |
| TEK | TEK tyrosine kinase, endothelial | 4.004 | |
| COL4A1 | collagen, type IV, alpha 1 | 3.987 | |
| NDNF | neuron-derived neurotrophic factor | 3.830 | |
| EPHA2 | EPH receptor A2 | 3.479 | |
| Genes expressed more in aBMSCs | EREG | epiregulin | 26.044 |
| TBX4 | T-box 4 | 6.917 | |
| LAMA5 | laminin subunit alpha 5 | 6.500 | |
| NOV | nephroblastoma overexpressed | 5.455 | |
| RAMP1 | receptor (G protein-coupled) activity modifying protein 1 | 5.053 | |
| EPGN | epithelial mitogen | 4.821 | |
| ESM1 | endothelial cell-specific molecule 1 | 3.833 | |
| COL18A1 | collagen, type XVIII, alpha 1 | 3.827 | |
| CCBE1 | collagen and calcium binding EGF domains 1 | 3.416 |
No detectable expression in aBMSCs
Table 3.
Differential expression of osteogenic genes between BMSCs and aBMSCs.
| Gene | Description | Fold | |
|---|---|---|---|
| Genes expressed more in BMSCs | SPP1 | secreted phosphoprotein 1 | 12.378 |
| IBSP | integrin-binding sialoprotein | 6.212 | |
| MGP | matrix Gla protein | 5.868 | |
| SEMA7A | semaphorin 7A, GPI membrane anchor (John Milton Hagen blood group) | 4.763 | |
| DLX5 | distal-less homeobox 5 | 4.534 | |
| PENK | proenkephalin | 3.772 | |
| ITGA11 | integrin subunit alpha 11 | 3.580 | |
| EPHA2 | EPH receptor A2 | 3.479 | |
| TNC | tenascin C | 3.047 | |
| Genes expressed more in aBMSCs | MSX2 | msh homeobox 2 | 9.273 |
| SFRP1 | secreted frizzled-related protein 1 | 9.217 | |
| GPNMB | glycoprotein (transmembrane) nmb | 9.030 | |
| BMP4 | bone morphogenetic protein 4 | 8.136 | |
| IGFBP5 | insulin like growth factor binding protein 5 | 5.040 |
4. Discussion
A recent report by Liu et. al identified aBMSCs as having greater osteogenic potential than BMSCs which are findings consistent with the results of our study (Liu, et. al 2020). However, our study is the first to evaluate the molecular, angiogenic, and osteogenic regenerative properties of MSC populations from osseous tissues of the same patients and as such, provides novel insights toward how the molecular identities of MSCs can be used to gain a deeper understanding of their regenerative properties. MSCs were isolated from two anatomically different osseous tissues from multiple subjects and their gene expression compared. There were 564 genes which were differentially expressed between BMSCs and aBMSCs of two different subjects; however, when comparing differential expression of genes between the same tissues, yet from these different subjects, it was very striking to note that among the over 23,000 genes, only 34 genes were DE between BMSCs and aBMSCs, i.e., 16 expressed at higher levels in BMSCs and 18 at higher levels in aBMSCs. These differences in gene expression did not manifest in differences of immunophenotype or multipotency; however, the trophic and regenerative properties of BMSCs and aBMSCs were different in model systems to evaluate their respective trophic and regenerative capacities.
To date, MSCs have been primarily characterized very broadly by their cell surface marker expression and multipotency in culture; however, our group and others have noted distinct differences in the properties of different MSC populations when placed in different clinical contexts (Behnia, et al. 2012; Bhansali, et al. 2014; Heldman, et al. 2014; Kaigler, et al. 2013). As such, it is clear that despite similarities in immunophenotype, the functional properties of different MSCs are unique and our understanding of these differences has remained very limited. qPCR and protein arrays have been used to compare different MSC populations and these data have provided initial clues about differences in MSC properties (Shi, et al. 2001; Sousa, et al. 2014). For the first time, our study used next generation sequencing to rigorously characterize MSCs based upon their molecular profile. In doing so, it was evident that the tissue source from which MSCs are derived has significant influence on the properties of these cells. Relative to their molecular profiles, it was striking to note that MSCs from the same site between different patients were more similar than MSCs from different sites within the same patient. Thus, contrary to what one may expect due to inherent variability between individuals, there was very little variability between individuals when MSCs were harvested from the same tissue sites. This has significant implications in cell therapies, particularly those involving allogeneic stem cells. Further studies evaluating MSCs from other tissues need to be performed to expand upon these findings.
Identification of specific molecular profiles is an important finding but extending these data to therapeutic applications is ultimately the goal. In the context of bone regeneration, while data from proteomic and molecular analyses suggest that some MSC populations are more apt to yield better bone regenerative outcomes than others, these studies have not linked data from the molecular properties of the cells to in vivo outcomes (Miranda, et al. 2012; Roson-Burgo, et al. 2014; Wagner, et al. 2007). In a randomized controlled clinical trial, our group demonstrated that clinical outcomes of mixed stem and progenitor cell transplantation in craniofacial bone defects is correlated with the percentage of CD90+ cells within these cell populations (Kaigler, et al. 2015). Even still, this correlation was limited to cell surface marker expression of only one of the numerous MSC markers. In our study, it was evident that aBMSC transplantation in calvarial defects resulted in greater bone formation in vivo than BMSCs and through next-generation sequencing, we were able to gain insight about why this difference may exist. Bone morphogenetic protein 4 (BMP-4), one of the most potent bone morphogens, was 8-fold more highly expressed in aBMSCs than in BMSC and this could in part underlie the regenerative outcome of these cells. There are clearly a multitude of other factors that could be implicated including the angiogenic capacity of these cells and their capacity to generate blood vessels where again, when incorporated into angiogenesis model systems, a more robust angiogenic response was observed from aBMSCs relative to BMSC. The use of next generation sequencing enables us to more rigorously characterize these cell populations and determine additional differences between them. Understanding these differences would enable us to extrapolate this knowledge to the clinical context where we could potentially screen different MSC populations based on their molecular profiles to determine their suitability for regenerating bone.
Though this work focused on MSC bone regenerative properties and angiogenesis, the impact for this work extends to other therapeutic properties of MSC, namely their immunomodulatory properties. The more recent identification of MSCs having the capacity to modulate immune responses has led to another important potential application of these cells. MSCs derived from different tissue sources have been evaluated to reduce inflammation and attenuate the immune response in acute and chronic conditions such as graft vs. host disease, lupus, degenerative joint diseases, and arthritis (Duijvestein, et al. 2010; Kikuiri, et al.; Liang, et al. 2010; Su, et al. 2011; Wong, et al. 2013; Zhang, et al. 2010). The outcomes seen in these preliminary reports have been very promising, yet no knowledge exists regarding how the molecular profile of these cells impact these outcomes. Greater depth of understanding is needed regarding the molecular characterization of these different cell populations. We previously reported that BMSCs and aBMSCs showed similar in vitro immunosuppressive effects on T cells and THP-1 monocytic cells, and both skewed THP-1 macrophages into a highly phagocytic phenotype (Cao, et al. 2020). However, the RNA sequencing data in this study show that there are numerous DE genes involved in immunity, immune responses, and inflammation between BMSCs and aBMSCs (Table 1). As such, further investigation is needed regarding how these differences manifest in an in vivo environment. It is worth noting that the lists include many genes directly involved in the complement system, chemotaxis, and modulation of immune cells. Our previously report showed that aBMSCs have an immunoevasive capacity similar to BMSCs, with neither eliciting T cell activation in vitro (Cao, et al. 2020). Nonetheless, the present study revealed that aBMSCs express higher levels of CD55, also known as decay-accelerating factor (DAF), which prevents the formation of the membrane attack complex in complement system. In an in vivo and clinical context, this may facilitate transplanted aBMSC survival (Li and Lin 2012) and could also contribute to the enhanced bone regenerative response observed from aBMSC transplantation relative to BMSCs. Additional studies are required to expand upon these findings.
The findings of our present study not only provide insight regarding how the molecular identities of BMSCs and aBMSCs relate to their properties, they provide evidence for us to consider revisiting how we have classified MSC populations (Dominici, et al. 2006). More rigorous characterization is needed to classify these cells based on their molecular identities and their functional properties vs. their cell surface marker expression and phenotype. In doing so, in a therapeutic context, we can determine which populations of MSCs may best be utilized for the therapeutic application of interest. While our findings are of interest, they require us to extend upon them and further studies are underway to address these questions which could have tremendous clinical impact.
Supplementary Material
Supplementary Fig. 1 BMSCs and aBMSCs immunophenotype.
Supplementary Fig. 2 BMP-4 gene expression levels in BMSCs and aBMSCs.
Supplementary Table 1 Genes expressed at higher levels in BMSCs compared to aBMSCs.
Supplementary Table 2 Genes expressed at higher levels in aBMSCs compared to BMSCs.
Acknowledgments
The authors would like to acknowledge David Adams and staff at the University of Michigan BRCF Flow Cytometry Core, Robert Lyons and staff at the University of Michigan Sequencing Core and Rich McEachin and staff at the University of Michigan Bioinformatics Core.
Funding
This study was funded by R01-DE028657 and R56-DE21410 from the NIH/NIDCR (awarded to D.K.) and R01-HL085339 from the NIH/NHLBI (awarded to A.J.P.).
Footnotes
Declarations
Conflict of interest
All authors declare that there is no conflict of interest concerning this work.
Ethical approval
All human tissues were collected from patients after obtaining informed consent following University of Michigan Institutional Review Board approval (IRB #HUM00034368) and all animal studies were conducted in accordance with University Institutional Animal Care and Use Committee approvals (PRO00009333, PRO00008804).
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Supplementary Materials
Supplementary Fig. 1 BMSCs and aBMSCs immunophenotype.
Supplementary Fig. 2 BMP-4 gene expression levels in BMSCs and aBMSCs.
Supplementary Table 1 Genes expressed at higher levels in BMSCs compared to aBMSCs.
Supplementary Table 2 Genes expressed at higher levels in aBMSCs compared to BMSCs.