The repair of bone fractures requires the generation of nerve cells throughout the injured area


In a December 2019 study, a team of Johns Hopkins Medicine researchers demonstrated in mice that repair of bone fractures requires the generation, growth and spread of nerve cells, or neurons, throughout the injured area.

This, they showed, partly relies on a protein known as nerve growth factor (NGF). Now, the researchers have dug deeper into this process to better understand how the nervous and immune systems work together with NGF to enable nerve regrowth during bone repair.

In a new study, published in the May 26, 2020, issue of the journal Cell Reports, the researchers found once again in mice that two proteins – tropomyosin receptor kinase-A (TrkA) and NGF – bind together to stimulate innervation (the supplying of nerves), and subsequently, new bone at an injured site.

What surprised them was that the NGF that mattered most in this process came from an unexpected source: macrophages, the white blood cells that alert the immune system to foreign invaders through inflammation, and then engulf and remove the attackers from the body.

“Previous research has shown that immune cells are clearly important in bone repair, but what we determined in our study is that macrophages and their inflammatory signals also kickstart nerve regrowth in injured bone,” says Aaron James, M.D., Ph.D., associate professor of pathology at the Johns Hopkins University School of Medicine and co-senior author of both studies.

In other words, James explains, the team’s experiments revealed “that NGF-TrkA signaling is how macrophages ‘talk’ to nerve fibers so that bone healing can begin.”

When bones are injured, there is a large release of the NGF neurotrophin (a protein that induces the survival, development and function of neurons). This activates sensory nerves to grow into the injured tissue.

These sensory nerves play multiple roles, including alerting the body through pain that the bone is broken and regulating the healing process.

To define the mechanism by which bone is repaired, the researchers removed the same small piece of skull from each of the mice in the study. By manipulating various steps of the NGF-TrkA signaling pathway in different mice, the team found that:

(1) the release of NGF coincides with the beginning of innervation,

(2) bone injury stimulates the increased production of NGF,

(3) inflammation at the injury site drives NGF production by macrophages (which are drawn by chemical signals released during inflammation),

(4) increased amounts of NGF elicit new nerve formation in the injured tissue,

(5) disrupting the production of NGF reduces innervation and impairs calvarial bone regeneration, and

(6) NGF produced by macrophages is the neurotrophin required for bone repair.

“We now understand that nerve growth and bone repair are linked processes,” James says. “Knowing this, we may be able to find ways to maximize our innate healing capacities.

Developing new methods to improve bone healing would greatly benefit many people, especially the elderly, where injuries such as hip fractures often lead to worse outcomes than heart attacks.”

The bones in our body are living tissues. They are composed of two types of tissues:

(1) The cortical (compact) bone as a hard outer layer, which is dense, strong, and tough; and

(2) The trabecular (cancellous) bone as a spongy inner layer[1].

Long bones, such as the tibia and femur, consist of articular cartilage, epiphyses, growth plate, metaphysis, diaphysis, periosteum, endosteum, and a marrow cavity[1]. Bones provide protection for vital organs and structural support for the body due to their tough and rigid structures resulting from a mineralized matrix[2].

Bones also act as a storage area for minerals (e.g., calcium) and provide a microenvironment for bone marrow (where blood cells are produced in long bones)[3].

During life, bones undergo organogenesis, modeling, and remodeling[4]. Bone modeling occurs when bone formation and bone resorption occur on separate surfaces, which means these two processes are not coupled during long bone increases in diameter and length[5].

Bone remodeling, the replacement of old bone by new bone, occurs primarily in the adult skeletal system to maintain bone mass[5]. This process involves the coupling of bone resorption and bone formation. Bone formation occurs by two distinct developmental processes.

Intramembranous ossification, which occurs by the direct differentiation of mesenchymal progenitors into osteoblasts, involves the replacement of connective tissue membrane with bone tissue[6].

Endochondral ossification involves the replacement of a hyaline cartilage model with bone tissue[7]. Bone repair or fracture healing proceeds through four phases: inflammation, intramembranous ossification, endochondral ossification, and bone remodeling[8].

Bone repair depends on the function of specific cell types, such as mesenchymal stem cells (MSCs) and osteoblasts[9,10]; the expression of soluble molecules (cytokines and growth factors)[11-13]; the scaffold (hydroxyapatite and extracellular matrix molecules)[14,15]; and various mechanical stimuli during the entire repair process[16,17].

Stem cells are defined as cells with the ability to self-renew and differentiate into different cell types[18]. According to their differentiation capacity, stem cells can be categorized as totipotent, pluripotent, multipotent, or unipotent[8].

Totipotent stem cells are capable of generating all of the cell types in animals, such as early blastomeres[19]. Pluripotent stem cells are capable of generating embryonic tissues from all three primary germ layers.

Induced pluripotent stem cells experimentally derive from adult somatic cells, and embryonic stem cells (ESCs) originate from the inner cell mass of the blastocyst[20-24].

Multipotent stem cells can differentiate into multiple specific cell types in a specific tissue or organ[25] and are located in specialized niches, where they can interact with the local microenvironment to maintain the stemness or differentiation potential.

The musculoskeletal system contains many multipotent stem cells. The most studied multipotent stem cells in the musculoskeletal system are the hematopoietic stem cells (HSCs)[26], which are the source of all types of blood cells, and bone marrow mesenchymal stem cells (BMMSCs), also known as bone marrow stromal cells (BMSCs)[27]. Unipotent stem cells can develop into only a single cell type[28,29].

The skeletal system contains multiple tissue types including bone, cartilage, blood vessels, nerves, and fat. Each tissue in the skeletal system is generated and maintained by the accurate management of specific stem cells.

Among the most well-known stem cells in the skeleton are the HSCs, defined as having the critical role of the long-term maintenance and production of all mature blood cell lineages during life[30,31].

The isolation of non-hematopoietic stem cells in the bone marrow relies on the ability of the cells to attach to plastic plates, which are thought to be ‘‘mesenchymal stem cells’’ or “skeletal stem cells.” These stem cells contain heterogeneous mixtures of cells with different potencies, such as bone, cartilage, adipo-tissue, endothelial cells, fibroblasts, and stroma.

At this time, the MSCs have two opposing descriptions. MSCs can be the self-renewing, postnatal, and multipotent stem cells for bone tissue, which are considered a specific type of bone marrow perivascular cell.

In contrast, MSCs can be ubiquitous in connective tissues and are defined by in vitro characteristics, such as adipose tissue[32,33], periosteum[34,35], the synovial joint[36-38], and muscle tissue[39,40]. In 2006, the International Society for Cellular Therapy proposed minimal criteria for defining the concept of human MSCs:

They must be plastic-adherent; highly express CD105, CD73, and CD90 while lacking expression of CD45, CD34, CD14 or CD11b, CD79a or CD19, and HLA-DR surface molecules; and be able to differentiate to osteoblasts, chondroblasts, and adipocytes in vitro[41].

This set of standards for the definition of human MSCs is consistent with laboratory-based scientific investigations and preclinical studies. However, the relationships between MSCs and SSCs are still not definitively known.

The SSC concept derives from experiments conducted by Friedenstein et al[42], who found that heterotopic transplants of bone marrow form reticular tissue and bone[42,43]. They confirmed the presence of colony-forming unit fibroblasts in the tissue culture plastic (TCP), adherent, non-hematopoietic cells in the bone marrow.

However, there remained considerable heterogeneity within the TCP-adherent cell population. The formation of the ectopic ossicle was ascribed to a specific cell population in the TCP-adherent cells.

Subsequently, the generation of an ossicle has been assigned to multipotent clonogenic progenitor cells, which give rise to cartilage, bone, and adipocytes[44]. These progenitor cells were first termed as osteogenic by Friedenstein et al[42] or as stromal stem cells by Owen et al[44]; they were then named MSCs by Caplan[45] and Pittenger et al[46]. Finally, they were considered SSCs by Bianco et al[47].

In past decades, several studies have attempted to identify cell surface markers that are expressed by SSCs, including the STRO-1 antigen, CD73, CD44, CD166, CD105, CD90, CD146, and CD271, or by negative selection for hematopoietic markers, such as CD45, CD34, CD14, CD79a, CD19, CD11b, and HLA-DR surface markers[48,49].

However, due to variation in certain markers, there is still a lack of consensus regarding the cell surface markers unique to SSCs. The absence of a set of specific surface markers may have contributed to the presence of confusing data in the literature related to the identification of SSCs.

Concerning the present controversy, the definition of SSCs states that the SSC population should have the capacity to produce four distinct lineages: bone, cartilage, adipo-tissue, and hematopoiesis-supportive stroma in vivo. Nevertheless, a list of specific surface markers, which could be extensively studied, would be widely accepted.

In 2013, Chan et al[50] reported a lineage-restricted and self-renewing skeletal progenitor that was isolated from the skeletal elements of fetal, neonatal, and adult mice and could form bone, cartilage, and bone marrow; it was named bone-cartilage-stromal progenitors (BCSPs). However, the main aim of the study was to focus on the regulation of the vascularization and hematopoiesis of HSCs by BCSPs, and they did not intensively study the role of BCSPs in bone regeneration or repair.

In 2015, two reports published in Cell helped to advance the SSC field and provide insight into the cell hierarchy[51,52]. A study by Worthley et al[51] used the secreted bone morphogenetic protein (BMP) agonist, Gremlin 1 (Grem1), to label skeletal progenitor cells. They found Grem1 positive cells beside the growth plate and determined that the trabecular bone could self-renew and generate diverse cells, such as osteoblasts, reticular marrow stromal cells, and chondrocytes but not adipocytes.

They later named them osteo-chondro-reticular (OCR) stem cells. In the femoral fracture callus, they found that Grem1+ OCR stem cells contributed to the expansion and differentiation into osteoblasts and chondrocytes. In another study, Chan et al[52] found clonal regions in the bone, especially at the growth plate, that encompassed bone, stromal tissue, and cartilage in mice. Subsequently, they showed that the CD45- Ter119- Tie2- AlphaV + Thy- 6C3- CD10- CD200+ cell population in the growth plate could self-renew in vitro and generate other subpopulations, such as pre-BCSP and BCSP.

These cell populations could specify their differentiation toward bone, cartilage, or stromal cells but not toward fat or muscle, which are regulated by soluble factors. They concluded that the CD45- Ter119- Tie2- AlphaV+ Thy- 6C3- CD105- CD200+ cell population represented SSCs in postnatal skeletal tissues.

Furthermore, they found that the SSC number increased in the callus of a femoral fracture more than in the uninjured femur with enhanced osteogenic capacity. In a similar study, Marecic et al[53] found that BCSP expansion preceded ossified callus formation in femoral fractures and that irradiation reduced the fracture-induced BCSP expansion.

The fracture-induced BCSPs (f-BCSPs) possessed greater plating efficiency, viability, alkaline phosphatase (ALP) activity, and Alizarin Red staining (ARS) than did the uninjured femur BCSPs (u-BCSPs). The f-BCSPs formed significantly larger bone specimens compared with u-BCSPs when transplanted under the renal capsules of immunodeficient mice. Although the hierarchy of stem cells and the differential capacity were studied in depth in these studies, little is known about the involvement of SSCs in bone development, modeling, and remodeling.

As mentioned above, SSCs are multipotent cells that differentiate into bone, cartilage, and stromal niches; however, they are unable to differentiate into other cell types, such as adipocytes, fibroblasts, muscle cells, or hematopoietic cells.

Chan et al[54] published another study in 2018, which focused on the human SSC. Using single cell RNA sequencing, fluorescence-activated cell sorting, and in vivo differentiation assays, they showed that the PDPN+ CD146- CD73+ CD164+ fetal growth plate cells produced the most colony-forming units in vitro and determined that they possessed self-renewal and multipotency, which were thought to be putative human SSCs.

Further hierarchical studies showed that this cell population was capable of the linear generation of osteogenic and chondrogenic subpopulations and was at the top of the differentiation tree. These studies established an ingenious human bone xenograft mouse model, transplanting human fetal phalangeal grafts with intact periosteum into immunodeficient mice; they found that fracture of the implanted bone induced the expansion of human SSCs near the fracture site. Furthermore, they found that human SSCs favored hematopoiesis and, conversely, that HSCs supported the human SSC lineage.

Another study published in 2018 by Mizuhashi et al[55] reported that SSCs were generated from PTHrP-positive chondrocytes in the resting zone of the growth plate in a mouse model. Mouse SSCs (41.6% ± 4.4%), pre-BCSP (31.7% ± 6.2%), and BCSP (53.4% ± 16.9%) were positive for PTHrP.

The analysis showed that PTHrP-positive chondrocytes, which are considered a unique SSC class in the resting zone, were multipotent and could longitudinally form columnar chondrocytes, which underwent hypertrophy, then became multiple types of cells, such as osteoblasts and marrow stromal cells, beneath the growth plate.

Additionally, these stem cells were able to send a signal to the transit-amplifying chondrocytes to maintain their proliferation so that they could maintain the integrity of the growth plate; transit-amplifying chondrocytes sent cues to determine the cell differentiation fates of PTHrP-positive chondrocytes in the resting zone.

The SSCs were derived from the growth plate in most of the abovementioned studies, which focused on their multipotency by transplanting stem cells under the renal capsules of immunodeficient mice involved in endochondral ossification.

Duchamp found that periosteal cells (PCs) and BMSCs were derived from the same embryonic Prx1-mesenchymal lineage and that postnatal PCs had an enhanced clonogenicity, growth, and differentiation capacity compared to BMSCs[56]. Although they did not identify the SSCs in the periosteum, they concluded that the presence of SSCs in the periosteum was associated with greater regenerative potency.

Another study, from Weill Cornell Medical School, identified SSCs, periosteal stem cells (PSCs), which were present in the periosteum of the long bones and calvarium of mice[57]. The PSCs displayed self-renewal and multipotent capacities and possessed different transcriptional signatures compared to the other SSCs.

As previously mentioned, other SSCs form bones through endochondral ossification, whereas PSCs form bones via a direct intramembranous pathway in the long bone or cranial bone. The differentiation capacity of PSCs for bone formation would therefore be enhanced in response to a fracture.

In 1991, Caplan[45] introduced the term “mesenchymal stem cells” to define the putative stem cells of skeletal tissues (bone and cartilage). The concept of MSCs extended to include bone marrow[58,59], adipose tissue[33,60], the periosteum[61], the synovial lining[62], muscle tissue[63], the umbilical cord[64], and different types of dental tissues[65]. Among them, BMMSCs were one of the well-studied sources.

It is currently thought that BMMSCs show an essential role in supporting bone healing through the secretion of nutritional and immunomodulatory factors rather than via a direct effect on the formation of the bone callus. BMMSCs secrete growth factors and cytokines to influence bone regeneration via paracrine and autocrine systems; this process includes vascular endothelial cell growth factors, platelet-derived growth factors, BMPs, fibroblast growth factors, insulin-like growth factor, and epidermal growth factor[65,66].

Inflammation is essential for any wound healing including bone repair. The first phase of fracture repair is the inflammation phase. Besides the trophic role, BMMSCs are critical regulators of the local inflammation micro-environment during bone repair. Macrophages are a key cell population that contributes to the inflammatory environment, whereas BMMSCs show an immunomodulatory effect on macrophages[67,68].

These inflammation factors include prostaglandin-E2[69], monocyte chemoattractant proteins (MCP-1 and MCP-3)[70], tumor necrosis factor-α[71], transforming growth factor-β[72], and numerous interleukins (IL-1, IL-3, IL-4, IL-6, and IL-10)[73,74].

Zuk et al[75] first described the isolation of adipose tissue-derived MSCs (ADSCs) from adipose tissue and characterized their phenotype and multipotency. Although ADSCs do not have superior osteogenic potential compared to BMMSCs in vitro[76-79], ADSCs are easier to acquire than BMMSCs.

ADSCs have been reported to exhibit high angiogenesis with either the ability to differentiate into endothelial cells or to secrete angiogenic factors, which favor osteogenesis and bone healing[80]. Moreover, ADSCs have a favorable effect on bone regeneration in vivo[81] and are widely used in clinical trials.

The periosteum is a tough layer of dense connective tissue that surrounds the bone surface, which contains different bone cells that enable bone to grow in thickness, which favors fracture repair and nourishes bone tissues[82].

The innermost layer contains stem cells that contribute to bone homeostasis and fracture healing, which respond to bone injury within 48 h through rapid proliferation. The stem cells from the periosteum have enhanced clonogenicity, growth, and differentiation capabilities[56,57]. Studies using reporter mice have identified Prx1 as a periosteal marker[83,84].

Studies in adult animals have shown that Prx1 is expressed in the periosteum and contributes to the formation of fracture callus[85]. Although only a limited number of studies have focused on the identification of MSCs in the periosteum, it is generally accepted that the periosteum plays an essential role in bone modeling and remodeling and is an important trophic pool for fracture healing.

Synovial tissue-derived mesenchymal stem cells (SMSCs) are obtained by a minimally invasive procedure and have been used for cartilage repair[86-89]. They are effective in regenerating critically sized bone defects when combined with polyether ketone[90], although few studies of SMSCs have focused on bone regeneration.

Muscle-derived MSCs also had high osteogenic potential in a mouse model[91] but need to be further characterized. Umbilical cord MSCs (UCMSCs) show a favorable osteogenic potential, similar to that of BMMSCs, and are able to contribute to bone and vessel regeneration[92]. UCMSCs also show great potential for bone regeneration in the presence of secretion factors[93-95], biomaterials[96-98], exosomes[99], and gene modification therapy[100,101].

Dental tissue-derived MSCs have been well-characterized and have shown features originally ascribed to BMMSCs. At least six different dental tissue-derived mesenchymal stem cell types have been isolated and have been described by Bartold et al[65]. Briefly, dental pulp stem cells and periodontal ligament stem cells exhibit considerable bone regenerative capabilities, whereas human apical papilla stem cells, dental follicle stem cells, exfoliated deciduous teeth stem cells, and gingival mesenchymal stem cells require further study[65].

Although hematopoietic cells are developmentally derived from the mesoderm in a manner similar to osteoblasts, they have no direct role in fracture healing or heterotopic ossification[102]. Other circulating cells, such as CD34+ cells from endothelial progenitor cells (EPCs), exhibit accelerated bone healing[103,104].

The EPCs, induced into the peripheral circulation by trauma, contribute to neovascularization and are involved in fracture healing[105,106]. CD31+ cells from peripheral blood facilitate bone endogenous regeneration by supporting immunomodulation and vascularization[107].

The circulating osteogenic progenitor cells, a type I collagen+/CD45+ subpopulation of mononuclear adherent cells in bone marrow, serve as osteogenic precursors for heterotopic ossification[108]. AMD3100, an antagonist of the chemokine receptor 4 that rapidly mobilizes stem cell populations into the peripheral blood, exerts significant beneficial effects, involving improved neovascularization and osteogenesis, on bone healing[109-111].

Using surgically conjoined transgenic mice which constitutively express green fluorescent protein (GFP) in no erythroid tissue and syngeneic wild-type mice models, circulating osteogenic connective tissue progenitors (GFP+ cells) from transgenic mice are mobilized to fracture sites in wild-type mice and contribute to osteogenic differentiation in the early stage of fracture healing[112].

Additionally, exposure to young cells, by heterochronic parabiosis, rejuvenates bone repair in aged animals[113]. Taken together, these results demonstrate that circulating progenitor cells play an important role in bone regeneration.

More information: Carolyn A. Meyers et al. A Neurotrophic Mechanism Directs Sensory Nerve Transit in Cranial Bone, Cell Reports (2020). DOI: 10.1016/j.celrep.2020.107696


1. le Noble F, le Noble J. Bone biology: Vessels of rejuvenation. Nature. 2014;507:313–314.

2. Buck DW 2nd, Dumanian GABone biology and physiology: Part I. The fundamentals. Plast Reconstr Surg. 2012;129:1314–1320.

3. Russell RG, Espina B, Hulley P. Bone biology and the pathogenesis of osteoporosis. Curr Opin Rheumatol. 2006;18 Suppl 1:S3–S10.

4. Raut N, Wicks SM, Lawal TO, Mahady GB. Epigenetic regulation of bone remodeling by natural compounds. Pharmacol Res. 2019;147:104350.

5. Sugiyama T, Oda H. Osteoporosis Therapy: Bone Modeling during Growth and Aging. Front Endocrinol (Lausanne) 2017;8:46.

6. Takarada T, Nakazato R, Tsuchikane A, Fujikawa K, Iezaki T, Yoneda Y, Hinoi E. Genetic analysis of Runx2 function during intramembranous ossification. Development. 2016;143:211–218.

7. Zhang Z, Leung WN, Li G, Lai YM, Chan CW. Osthole Promotes Endochondral Ossification and Accelerates Fracture Healing in Mice. Calcif Tissue Int. 2016;99:649–660.

8. Haffner-Luntzer M, Weber B, Lam C, Fischer V, Lackner I, Ignatius A, Kalbitz M, Marcucio RS, Miclau T. A novel mouse model to study fracture healing of the proximal femur. J Orthop Res. 2020:Online ahead of print.

9. Oryan A, Kamali A, Moshiri A, Baghaban Eslaminejad M. Role of Mesenchymal Stem Cells in Bone Regenerative Medicine: What Is the Evidence? Cells Tissues Organs. 2017;204:59–83.

10. Chu C, Wei S, Wang Y, Wang Y, Man Y, Qu Y. Extracellular vesicle and mesenchymal stem cells in bone regeneration: recent progress and perspectives. J Biomed Mater Res A. 2019;107:243–250.

11. Hu K, Olsen BR. The roles of vascular endothelial growth factor in bone repair and regeneration. Bone. 2016;91:30–38.

12. Martino MM, Briquez PS, Maruyama K, Hubbell JA. Extracellular matrix-inspired growth factor delivery systems for bone regeneration. Adv Drug Deliv Rev. 2015;94:41–52.

13. Samorezov JE, Alsberg E. Spatial regulation of controlled bioactive factor delivery for bone tissue engineering. Adv Drug Deliv Rev. 2015;84:45–67.

14. Chen X, Fan H, Deng X, Wu L, Yi T, Gu L, Zhou C, Fan Y, Zhang X. Scaffold Structural Microenvironmental Cues to Guide Tissue Regeneration in Bone Tissue Applications. Nanomaterials (Basel) 2018;8

15. Holt BD, Wright ZM, Arnold AM, Sydlik SA. Graphene oxide as a scaffold for bone regeneration. Wiley Interdiscip Rev Nanomed Nanobiotechnol. 2017;9

16. Glatt V, Evans CH, Tetsworth K. A Concert between Biology and Biomechanics: The Influence of the Mechanical Environment on Bone Healing. Front Physiol. 2016;7:678.

17. Betts DC, Müller R. Mechanical regulation of bone regeneration: theories, models, and experiments. Front Endocrinol (Lausanne) 2014;5:211.

18. Guan JL, Simon AK, Prescott M, Menendez JA, Liu F, Wang F, Wang C, Wolvetang E, Vazquez-Martin A, Zhang J. Autophagy in stem cells. Autophagy. 2013;9:830–849.

19. Baker CL, Pera MF. Capturing Totipotent Stem Cells. Cell Stem Cell. 2018;22:25–34.

20. Hayashi K, Saitou M. Generation of eggs from mouse embryonic stem cells and induced pluripotent stem cells. Nat Protoc. 2013;8:1513–1524.

21. Ran D, Shia WJ, Lo MC, Fan JB, Knorr DA, Ferrell PI, Ye Z, Yan M, Cheng L, Kaufman DS, Zhang DE. RUNX1a enhances hematopoietic lineage commitment from human embryonic stem cells and inducible pluripotent stem cells. Blood. 2013;121:2882–2890.

22. Phelan DG, Anderson DJ, Howden SE, Wong RC, Hickey PF, Pope K, Wilson GR, Pébay A, Davis AM, Petrou S, Elefanty AG, Stanley EG, James PA, Macciocca I, Bahlo M, Cheung MM, Amor DJ, Elliott DA, Lockhart PJ. ALPK3-deficient cardiomyocytes generated from patient-derived induced pluripotent stem cells and mutant human embryonic stem cells display abnormal calcium handling and establish that ALPK3 deficiency underlies familial cardiomyopathy. Eur Heart J. 2016;37:2586–2590.

23. Zhang M, Ngo J, Pirozzi F, Sun YP, Wynshaw-Boris A. Highly efficient methods to obtain homogeneous dorsal neural progenitor cells from human and mouse embryonic stem cells and induced pluripotent stem cells. Stem Cell Res Ther. 2018;9:67.

24. De Los Angeles A, Ferrari F, Xi R, Fujiwara Y, Benvenisty N, Deng H, Hochedlinger K, Jaenisch R, Lee S, Leitch HG, Lensch MW, Lujan E, Pei D, Rossant J, Wernig M, Park PJ, Daley GQ. Hallmarks of pluripotency. Nature. 2015;525:469–478.

25. Mirzaei H, Sahebkar A, Sichani LS, Moridikia A, Nazari S, Sadri Nahand J, Salehi H, Stenvang J, Masoudifar A, Mirzaei HR, Jaafari MR. Therapeutic application of multipotent stem cells. J Cell Physiol. 2018;233:2815–2823.

26. Monteiro R, Pinheiro P, Joseph N, Peterkin T, Koth J, Repapi E, Bonkhofer F, Kirmizitas A, Patient R. Transforming Growth Factor β Drives Hemogenic Endothelium Programming and the Transition to Hematopoietic Stem Cells. Dev Cell. 2016;38:358–370.

27. Gao X, Usas A, Tang Y, Lu A, Tan J, Schneppendahl J, Kozemchak AM, Wang B, Cummins JH, Tuan RS, Huard J. A comparison of bone regeneration with human mesenchymal stem cells and muscle-derived stem cells and the critical role of BMP. Biomaterials. 2014;35:6859–6870.

28. Lilja AM, Rodilla V, Huyghe M, Hannezo E, Landragin C, Renaud O, Leroy O, Rulands S, Simons BD, Fre S. Clonal analysis of Notch1-expressing cells reveals the existence of unipotent stem cells that retain long-term plasticity in the embryonic mammary gland. Nat Cell Biol. 2018;20:677–687.

29. Ko K, Araúzo-Bravo MJ, Kim J, Stehling M, Schöler HR. Conversion of adult mouse unipotent germline stem cells into pluripotent stem cells. Nat Protoc. 2010;5:921–928.

30. Dzierzak E, Bigas A. Blood Development: Hematopoietic Stem Cell Dependence and Independence. Cell Stem Cell. 2018;22:639–651.

31. Baum CM, Weissman IL, Tsukamoto AS, Buckle AM, Peault B. Isolation of a candidate human hematopoietic stem-cell population. Proc Natl Acad Sci USA. 1992;89:2804–2808.

32. Bacakova L, Zarubova J, Travnickova M, Musilkova J, Pajorova J, Slepicka P, Kasalkova NS, Svorcik V, Kolska Z, Motarjemi H, Molitor M. Stem cells: their source, potency and use in regenerative therapies with focus on adipose-derived stem cells – a review. Biotechnol Adv. 2018;36:1111–1126.

33. Rodriguez AM, Elabd C, Amri EZ, Ailhaud G, Dani C. The human adipose tissue is a source of multipotent stem cells. Biochimie. 2005;87:125–128.

34. Collette NM, Yee CS, Hum NR, Murugesh DK, Christiansen BA, Xie L, Economides AN, Manilay JO, Robling AG, Loots GG. Sostdc1 deficiency accelerates fracture healing by promoting the expansion of periosteal mesenchymal stem cells. Bone. 2016;88:20–30.

35. Kudva AK, Luyten FP, Patterson J. In Vitro Screening of Molecularly Engineered Polyethylene Glycol Hydrogels for Cartilage Tissue Engineering using Periosteum-Derived and ATDC5 Cells. Int J Mol Sci. 2018:19.

36. Yasui Y, Hart DA, Sugita N, Chijimatsu R, Koizumi K, Ando W, Moriguchi Y, Shimomura K, Myoui A, Yoshikawa H, Nakamura N. Time-Dependent Recovery of Human Synovial Membrane Mesenchymal Stem Cell Function After High-Dose Steroid Therapy: Case Report and Laboratory Study. Am J Sports Med. 2018;46:695–701.

37. Roelofs AJ, Zupan J, Riemen AHK, Kania K, Ansboro S, White N, Clark SM, De Bari C. Joint morphogenetic cells in the adult mammalian synovium. Nat Commun. 2017;8:15040.

38. Neybecker P, Henrionnet C, Pape E, Mainard D, Galois L, Loeuille D, Gillet P, Pinzano A. In vitro and in vivo potentialities for cartilage repair from human advanced knee osteoarthritis synovial fluid-derived mesenchymal stem cells. Stem Cell Res Ther. 2018;9:329.

39. Owston H, Giannoudis PV, Jones E. Do skeletal muscle MSCs in humans contribute to bone repair? A systematic review. Injury. 2016;47 Suppl 6:S3–S15.

40. Fellows CR, Matta C, Zakany R, Khan IM, Mobasheri A. Adipose, Bone Marrow and Synovial Joint-Derived Mesenchymal Stem Cells for Cartilage Repair. Front Genet. 2016;7:213.

41. Dominici M, Le Blanc K, Mueller I, Slaper-Cortenbach I, Marini F, Krause D, Deans R, Keating A, Prockop Dj, Horwitz E. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy. 2006;8:315–317.

42. Friedenstein AJ, Piatetzky-Shapiro II, Petrakova KV. Osteogenesis in transplants of bone marrow cells. J Embryol Exp Morphol. 1966;16:381–390.

43. Tavassoli M, Crosby WH. Transplantation of marrow to extramedullary sites. Science. 1968;161:54–56.

44. Owen M, Friedenstein AJ. Stromal stem cells: marrow-derived osteogenic precursors. Ciba Found Symp. 1988;136:42–60.

45. Caplan AI. Mesenchymal stem cells. J Orthop Res. 1991;9:641–650.

46. Pittenger MF, Mackay AM, Beck SC, Jaiswal RK, Douglas R, Mosca JD, Moorman MA, Simonetti DW, Craig S, Marshak DR. Multilineage potential of adult human mesenchymal stem cells. Science. 1999;284:143–147.

47. Bianco P, Kuznetsov SA, Riminucci M, Gehron Robey P. Postnatal skeletal stem cells. Methods Enzymol. 2006;419:117–148.

48. Tare RS, Babister JC, Kanczler J, Oreffo RO. Skeletal stem cells: phenotype, biology and environmental niches informing tissue regeneration. Mol Cell Endocrinol. 2008;288:11–21.

49. Dawson JI, Kanczler J, Tare R, Kassem M, Oreffo RO. Concise review: bridging the gap: bone regeneration using skeletal stem cell-based strategies – where are we now? Stem Cells. 2014;32:35–44.

50. Chan CK, Lindau P, Jiang W, Chen JY, Zhang LF, Chen CC, Seita J, Sahoo D, Kim JB, Lee A, Park S, Nag D, Gong Y, Kulkarni S, Luppen CA, Theologis AA, Wan DC, DeBoer A, Seo EY, Vincent-Tompkins JD, Loh K, Walmsley GG, Kraft DL, Wu JC, Longaker MT, Weissman IL. Clonal precursor of bone, cartilage, and hematopoietic niche stromal cells. Proc Natl Acad Sci USA. 2013;110:12643–12648.

51. Worthley DL, Churchill M, Compton JT, Tailor Y, Rao M, Si Y, Levin D, Schwartz MG, Uygur A, Hayakawa Y, Gross S, Renz BW, Setlik W, Martinez AN, Chen X, Nizami S, Lee HG, Kang HP, Caldwell JM, Asfaha S, Westphalen CB, Graham T, Jin G, Nagar K, Wang H, Kheirbek MA, Kolhe A, Carpenter J, Glaire M, Nair A, Renders S, Manieri N, Muthupalani S, Fox JG, Reichert M, Giraud AS, Schwabe RF, Pradere JP, Walton K, Prakash A, Gumucio D, Rustgi AK, Stappenbeck TS, Friedman RA, Gershon MD, Sims P, Grikscheit T, Lee FY, Karsenty G, Mukherjee S, Wang TC. Gremlin 1 identifies a skeletal stem cell with bone, cartilage, and reticular stromal potential. Cell. 2015;160:269–284.

52. Chan CK, Seo EY, Chen JY, Lo D, McArdle A, Sinha R, Tevlin R, Seita J, Vincent-Tompkins J, Wearda T, Lu WJ, Senarath-Yapa K, Chung MT, Marecic O, Tran M, Yan KS, Upton R, Walmsley GG, Lee AS, Sahoo D, Kuo CJ, Weissman IL, Longaker MT. Identification and specification of the mouse skeletal stem cell. Cell. 2015;160:285–298.

53. Marecic O, Tevlin R, McArdle A, Seo EY, Wearda T, Duldulao C, Walmsley GG, Nguyen A, Weissman IL, Chan CK, Longaker MT. Identification and characterization of an injury-induced skeletal progenitor. Proc Natl Acad Sci USA. 2015;112:9920–9925.

54. Chan CKF, Gulati GS, Sinha R, Tompkins JV, Lopez M, Carter AC, Ransom RC, Reinisch A, Wearda T, Murphy M, Brewer RE, Koepke LS, Marecic O, Manjunath A, Seo EY, Leavitt T, Lu WJ, Nguyen A, Conley SD, Salhotra A, Ambrosi TH, Borrelli MR, Siebel T, Chan K, Schallmoser K, Seita J, Sahoo D, Goodnough H, Bishop J, Gardner M, Majeti R, Wan DC, Goodman S, Weissman IL, Chang HY, Longaker MT. Identification of the Human Skeletal Stem Cell. Cell. 2018;175:43–56.e21.

55. Mizuhashi K, Ono W, Matsushita Y, Sakagami N, Takahashi A, Saunders TL, Nagasawa T, Kronenberg HM, Ono N. Resting zone of the growth plate houses a unique class of skeletal stem cells. Nature. 2018;563:254–258.

56. Duchamp de Lageneste O, Julien A, Abou-Khalil R, Frangi G, Carvalho C, Cagnard N, Cordier C, Conway SJ, Colnot C. Periosteum contains skeletal stem cells with high bone regenerative potential controlled by Periostin. Nat Commun. 2018;9:773.

57. Debnath S, Yallowitz AR, McCormick J, Lalani S, Zhang T, Xu R, Li N, Liu Y, Yang YS, Eiseman M, Shim JH, Hameed M, Healey JH, Bostrom MP, Landau DA, Greenblatt MB. Discovery of a periosteal stem cell mediating intramembranous bone formation. Nature. 2018;562:133–139.

58. Zhong W, Zhu Z, Xu X, Zhang H, Xiong H, Li Q, Wei Y. Human bone marrow-derived mesenchymal stem cells promote the growth and drug-resistance of diffuse large B-cell lymphoma by secreting IL-6 and elevating IL-17A levels. J Exp Clin Cancer Res. 2019;38:73.

59. Su T, Xiao Y, Xiao Y, Guo Q, Li C, Huang Y, Deng Q, Wen J, Zhou F, Luo XH. Bone Marrow Mesenchymal Stem Cells-Derived Exosomal MiR-29b-3p Regulates Aging-Associated Insulin Resistance. ACS Nano. 2019;13:2450–2462.

60. Liu X, Xiang Q, Xu F, Huang J, Yu N, Zhang Q, Long X, Zhou Z. Single-cell RNA-seq of cultured human adipose-derived mesenchymal stem cells. Sci Data. 2019;6:190031.

61. De Bari C, Dell’Accio F, Vanlauwe J, Eyckmans J, Khan IM, Archer CW, Jones EA, McGonagle D, Mitsiadis TA, Pitzalis C, Luyten FP. Mesenchymal multipotency of adult human periosteal cells demonstrated by single-cell lineage analysis. Arthritis Rheum. 2006;54:1209–1221.

62. Murata Y, Uchida S, Utsunomiya H, Hatakeyama A, Nakashima H, Chang A, Sekiya I, Sakai A. Synovial Mesenchymal Stem Cells Derived From the Cotyloid Fossa Synovium Have Higher Self-renewal and Differentiation Potential Than Those From the Paralabral Synovium in the Hip Joint. Am J Sports Med. 2018;46:2942–2953.

63. Klimczak A, Kozlowska U, Kurpisz M. Muscle Stem/Progenitor Cells and Mesenchymal Stem Cells of Bone Marrow Origin for Skeletal Muscle Regeneration in Muscular Dystrophies. Arch Immunol Ther Exp (Warsz) 2018;66:341–354.

64. Liu KX, Zhu YX, Yan YM, Zeng Y, Jiao YB, Qin FY, Liu JW, Zhang YY, Cheng YX. Discovery of Populusone, a Skeletal Stimulator of Umbilical Cord Mesenchymal Stem Cells from Populus euphratica Exudates. Org Lett. 2019;21:1837–1840.

65. Bartold M, Gronthos S, Haynes D, Ivanovski S. Mesenchymal stem cells and biologic factors leading to bone formation. J Clin Periodontol. 2019;46 Suppl 21:12–32.

66. Caplan AI, Dennis JE. Mesenchymal stem cells as trophic mediators. J Cell Biochem. 2006;98:1076–1084.

67. Pajarinen J, Lin T, Gibon E, Kohno Y, Maruyama M, Nathan K, Lu L, Yao Z, Goodman SB. Mesenchymal stem cell-macrophage crosstalk and bone healing. Biomaterials. 2019;196:80–89.

68. Chen B, Ni Y, Liu J, Zhang Y, Yan F. Bone Marrow-Derived Mesenchymal Stem Cells Exert Diverse Effects on Different Macrophage Subsets. Stem Cells Int. 2018;2018:8348121.

69. Feigenson M, Eliseev RA, Jonason JH, Mills BN, O’Keefe RJ. PGE2 Receptor Subtype 1 (EP1) Regulates Mesenchymal Stromal Cell Osteogenic Differentiation by Modulating Cellular Energy Metabolism. J Cell Biochem. 2017;118:4383–4393.

70. Suzuki K, Chosa N, Sawada S, Takizawa N, Yaegashi T, Ishisaki A. Enhancement of Anti-Inflammatory and Osteogenic Abilities of Mesenchymal Stem Cells via Cell-to-Cell Adhesion to Periodontal Ligament-Derived Fibroblasts. Stem Cells Int. 2017;2017:3296498.

71. Du D, Zhou Z, Zhu L, Hu X, Lu J, Shi C, Chen F, Chen A. TNF-α suppresses osteogenic differentiation of MSCs by accelerating P2Y2 receptor in estrogen-deficiency induced osteoporosis. Bone. 2018;117:161–170.

72. Crane JL, Cao X. Bone marrow mesenchymal stem cells and TGF-β signaling in bone remodeling. J Clin Invest. 2014;124:466–472.

73. Walters G, Pountos I, Giannoudis PV. The cytokines and micro-environment of fracture haematoma: Current evidence. J Tissue Eng Regen Med. 2018;12:e1662–e1677.

74. Lin T, Pajarinen J, Kohno Y, Maruyama M, Romero-Lopez M, Huang JF, Nathan K, Khan TN, Yao Z, Goodman SB. Transplanted interleukin-4–secreting mesenchymal stromal cells show extended survival and increased bone mineral density in the murine femur. Cytotherapy. 2018;20:1028–1036.

75. Zuk PA, Zhu M, Mizuno H, Huang J, Futrell JW, Katz AJ, Benhaim P, Lorenz HP, Hedrick MH. Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng. 2001;7:211–228.

76. De Ugarte DA, Morizono K, Elbarbary A, Alfonso Z, Zuk PA, Zhu M, Dragoo JL, Ashjian P, Thomas B, Benhaim P, Chen I, Fraser J, Hedrick MH. Comparison of multi-lineage cells from human adipose tissue and bone marrow. Cells Tissues Organs. 2003;174:101–109.

77. Liu TM, Martina M, Hutmacher DW, Hui JH, Lee EH, Lim B. Identification of common pathways mediating differentiation of bone marrow- and adipose tissue-derived human mesenchymal stem cells into three mesenchymal lineages. Stem Cells. 2007;25:750–760.

78. Shafiee A, Seyedjafari E, Soleimani M, Ahmadbeigi N, Dinarvand P, Ghaemi N. A comparison between osteogenic differentiation of human unrestricted somatic stem cells and mesenchymal stem cells from bone marrow and adipose tissue. Biotechnol Lett. 2011;33:1257–1264.

79. Park SH, Sim WY, Min BH, Yang SS, Khademhosseini A, Kaplan DL. Chip-based comparison of the osteogenesis of human bone marrow- and adipose tissue-derived mesenchymal stem cells under mechanical stimulation. PLoS One. 2012;7:e46689.

80. Kim Y, Kim H, Cho H, Bae Y, Suh K, Jung J. Direct comparison of human mesenchymal stem cells derived from adipose tissues and bone marrow in mediating neovascularization in response to vascular ischemia. Cell Physiol Biochem. 2007;20:867–876.

81. Liao HT, Chen CT. Osteogenic potential: Comparison between bone marrow and adipose-derived mesenchymal stem cells. World J Stem Cells. 2014;6:288–295.

82. Evans SF, Parent JB, Lasko CE, Zhen X, Knothe UR, Lemaire T, Knothe Tate ML. Periosteum, bone’s “smart” bounding membrane, exhibits direction-dependent permeability. J Bone Miner Res. 2013;28:608–617.

83. Kawanami A, Matsushita T, Chan YY, Murakami S. Mice expressing GFP and CreER in osteochondro progenitor cells in the periosteum. Biochem Biophys Res Commun. 2009;386:477–482.

84. Ouyang Z, Chen Z, Ishikawa M, Yue X, Kawanami A, Leahy P, Greenfield EM, Murakami S. Prx1 and 3.2kb Col1a1 promoters target distinct bone cell populations in transgenic mice. Bone. 2014;58:136–145.

85. Murao H, Yamamoto K, Matsuda S, Akiyama H. Periosteal cells are a major source of soft callus in bone fracture. J Bone Miner Metab. 2013;31:390–398.

86. Lee JC, Min HJ, Park HJ, Lee S, Seong SC, Lee MC. Synovial membrane-derived mesenchymal stem cells supported by platelet-rich plasma can repair osteochondral defects in a rabbit model. Arthroscopy. 2013;29:1034–1046.

87. Noël D, Gazit D, Bouquet C, Apparailly F, Bony C, Plence P, Millet V, Turgeman G, Perricaudet M, Sany J, Jorgensen C. Short-term BMP-2 expression is sufficient for in vivo osteochondral differentiation of mesenchymal stem cells. Stem Cells. 2004;22:74–85.

88. Chen K, Man C, Zhang B, Hu J, Zhu SS. Effect of in vitro chondrogenic differentiation of autologous mesenchymal stem cells on cartilage and subchondral cancellous bone repair in osteoarthritis of temporomandibular joint. Int J Oral Maxillofac Surg. 2013;42:240–248.

89. Shimomura K, Moriguchi Y, Nansai R, Fujie H, Ando W, Horibe S, Hart DA, Gobbi A, Yoshikawa H, Nakamura N. Comparison of 2 Different Formulations of Artificial Bone for a Hybrid Implant With a Tissue-Engineered Construct Derived From Synovial Mesenchymal Stem Cells: A Study Using a Rabbit Osteochondral Defect Model. Am J Sports Med. 2017;45:666–675.

90. Lin Y, Umebayashi M, Abdallah MN, Dong G, Roskies MG, Zhao YF, Murshed M, Zhang Z, Tran SD. Combination of polyetherketoneketone scaffold and human mesenchymal stem cells from temporomandibular joint synovial fluid enhances bone regeneration. Sci Rep. 2019;9:472.

91. Liu X, Kumagai G, Wada K, Tanaka T, Asari T, Oishi K, Fujita T, Mizukami H, Furukawa KI, Ishibashi Y. High Osteogenic Potential of Adipose- and Muscle-derived Mesenchymal Stem Cells in Spinal-Ossification Model Mice. Spine (Phila Pa 1976) 2017;42:E1342–E1349.

92. Wang Q, Zhao G, Xing Z, Zhan J, Ma J. Comparative evaluation of the osteogenic capacity of human mesenchymal stem cells from bone marrow and umbilical cord tissue. Exp Ther Med. 2019;17:764–772.

93. Wang KX, Xu LL, Rui YF, Huang S, Lin SE, Xiong JH, Li YH, Lee WY, Li G. The effects of secretion factors from umbilical cord derived mesenchymal stem cells on osteogenic differentiation of mesenchymal stem cells. PLoS One. 2015;10:e0120593.

94. Deng M, Luo K, Hou T, Luo F, Xie Z, Zhang Z, Yang A, Yu B, Yi S, Tan J, Dong S, Xu J. IGFBP3 deposited in the human umbilical cord mesenchymal stem cell-secreted extracellular matrix promotes bone formation. J Cell Physiol. 2018;233:5792–5804.

95. Todeschi MR, El Backly R, Capelli C, Daga A, Patrone E, Introna M, Cancedda R, Mastrogiacomo M. Transplanted Umbilical Cord Mesenchymal Stem Cells Modify the In Vivo Microenvironment Enhancing Angiogenesis and Leading to Bone Regeneration. Stem Cells Dev. 2015;24:1570–1581.

96. Wang P, Liu X, Zhao L, Weir MD, Sun J, Chen W, Man Y, Xu HH. Bone tissue engineering via human induced pluripotent, umbilical cord and bone marrow mesenchymal stem cells in rat cranium. Acta Biomater. 2015;18:236–248.

97. Day AGE, Francis WR, Fu K, Pieper IL, Guy O, Xia Z. Osteogenic Potential of Human Umbilical Cord Mesenchymal Stem Cells on Coralline Hydroxyapatite/Calcium Carbonate Microparticles. Stem Cells Int. 2018;2018:4258613.

98. Zhao L, Weir MD, Xu HH. An injectable calcium phosphate-alginate hydrogel-umbilical cord mesenchymal stem cell paste for bone tissue engineering. Biomaterials. 2010;31:6502–6510.

99. Zhang Y, Hao Z, Wang P, Xia Y, Wu J, Xia D, Fang S, Xu S. Exosomes from human umbilical cord mesenchymal stem cells enhance fracture healing through HIF-1α-mediated promotion of angiogenesis in a rat model of stabilized fracture. Cell Prolif. 2019;52:e12570.

100. Bougioukli S, Saitta B, Sugiyama O, Tang AH, Elphingstone J, Evseenko D, Lieberman JR. Lentiviral Gene Therapy for Bone Repair Using Human Umbilical Cord Blood-Derived Mesenchymal Stem Cells. Hum Gene Ther. 2019;30:906–917.

101. Ciavarella S, Dammacco F, De Matteo M, Loverro G, Silvestris F. Umbilical cord mesenchymal stem cells: role of regulatory genes in their differentiation to osteoblasts. Stem Cells Dev. 2009;18:1211–1220.

102. Otsuru S, Overholt KM, Olson TS, Hofmann TJ, Guess AJ, Velazquez VM, Kaito T, Dominici M, Horwitz EM. Hematopoietic derived cells do not contribute to osteogenesis as osteoblasts. Bone. 2017;94:1–9.

103. Kawakami Y, Matsumoto T, Mifune Y, Fukui T, Patel KG, Walker GN, Kurosaka M, Kuroda R. Therapeutic Potential of Endothelial Progenitor Cells in the Field of Orthopaedics. Curr Stem Cell Res Ther. 2017;12:3–13.

104. Kuroda R, Matsumoto T, Kawakami Y, Fukui T, Mifune Y, Kurosaka M. Clinical impact of circulating CD34-positive cells on bone regeneration and healing. Tissue Eng Part B Rev. 2014;20:190–199.

105. Ma XL, Sun XL, Wan CY, Ma JX, Tian P. Significance of circulating endothelial progenitor cells in patients with fracture healing process. J Orthop Res. 2012;30:1860–1866.

106. Matsumoto T, Kuroda R, Mifune Y, Kawamoto A, Shoji T, Miwa M, Asahara T, Kurosaka M. Circulating endothelial/skeletal progenitor cells for bone regeneration and healing. Bone. 2008;43:434–439.

107. Sass FA, Schmidt-Bleek K, Ellinghaus A, Filter S, Rose A, Preininger B, Reinke S, Geissler S, Volk HD, Duda GN, Dienelt A. CD31+ Cells From Peripheral Blood Facilitate Bone Regeneration in Biologically Impaired Conditions Through Combined Effects on Immunomodulation and Angiogenesis. J Bone Miner Res. 2017;32:902–912.

108. Suda RK, Billings PC, Egan KP, Kim JH, McCarrick-Walmsley R, Glaser DL, Porter DL, Shore EM, Pignolo RJ. Circulating osteogenic precursor cells in heterotopic bone formation. Stem Cells. 2009;27:2209–2219.

109. Wang XX, Allen RJ, Jr, Tutela JP, Sailon A, Allori AC, Davidson EH, Paek GK, Saadeh PB, McCarthy JG, Warren SM. Progenitor cell mobilization enhances bone healing by means of improved neovascularization and osteogenesis. Plast Reconstr Surg. 2011;128:395–405.

110. Toupadakis CA, Granick JL, Sagy M, Wong A, Ghassemi E, Chung DJ, Borjesson DL, Yellowley CE. Mobilization of endogenous stem cell populations enhances fracture healing in a murine femoral fracture model. Cytotherapy. 2013;15:1136–1147.

111. Meeson R, Sanghani-Keri A, Coathup M, Blunn G. VEGF with AMD3100 endogenously mobilizes mesenchymal stem cells and improves fracture healing. J Orthop Res. 2019;37:1294–1302.

112. Kumagai K, Vasanji A, Drazba JA, Butler RS, Muschler GF. Circulating cells with osteogenic potential are physiologically mobilized into the fracture healing site in the parabiotic mice model. J Orthop Res. 2008;26:165–175.


Please enter your comment!
Please enter your name here

Questo sito usa Akismet per ridurre lo spam. Scopri come i tuoi dati vengono elaborati.