Validated methods for isolation and qualification of mesenchymal stromal/stem cells from different sources

Abstract

Mesenchymal Stromal/Stem Cells (MSCs) have attracted considerable attention in the field of regenerative medicine. Their unique properties make them suitable for various therapeutic applications. This article reviews accepted methods and guidelines for the isolation and characterization of MSCs from various sources. Common sources include bone marrow, adipose tissue, perinatal and umbilical cord tissue, dental pulp, etc. Naturally, the techniques used to isolate MSCs can vary depending on the source from which they are derived. However, several methods have been widely accepted by the scientific community. These include enzymatic digestion, density gradient centrifugation, the use of Percoll, adherence-based techniques and selective culture conditions. To characterize MSCs, basic criteria established by the International Society for Cell and Tissue Transplantation and the International Federation for Adipose Tissue are routinely used. These criteria include the ability of MSCs to adhere to plastic surfaces under standard culture conditions, the expression of specific membrane markers and their differentiation potential. Various techniques are used to assess these characteristics, including mixed lymphocyte reactions, flow cytometry and immunophenotyping profiles. These assessments aim to confirm the purity of the MSCs and validate their mesenchymal properties. In summary, the isolation and characterization of MSCs requires careful consideration of the different available methods. Each source presents unique challenges and advantages. By following established guidelines, researchers can ensure successful isolation and characterization of MSCs. This knowledge will ultimately improve their use in regenerative medicine.

Introduction

Cell-based therapy is a fast-growing medical procedure that uses different cell products for a variety of indications. The first successful stem cell transplantation was performed in 1958 by George Mathe when bone marrow was transplanted to treat six nuclear researchers exposed to radioactive substances [1]. Such a seminal procedure rapidly led to a better understanding of the immune system and allowed the development of diverse autologous and allogeneic cell therapies. Among these, Mesenchymal Stromal Cells (MSCs) are undoubtedly the most used cells for a wide range of corrective or interventional therapies [2, 3]. More than 30 years ago, Dr. Caplan provocatively reinterpreted the term in “mesenchymal stem cells” , in response to their multipotency and highly proliferative capacity and such an abbreviation has become extremely popular as the stem cell field was growing and gaining attention leading to a large part of the scientific literature being classified under this stemness characteristic [4]. . The still popular concept of mesenchymal “stem” cells lays roots in classical experiments on the transplantation of bone marrow (the first and still most common site of extraction for adult MSC) to heterotopic anatomical sites, resulting in de novo generation of ectopic bone and marrow. The Russian scientist A.J. Friedenstein is generally credited as the father of the MSC. Friedenstein started his pioneering work with the critical observation that bone marrow may serve as a reservoir of precursor cells for osteogenic and hematopoietic tissues. Friedenstein and colleagues isolated adherent, fibroblast-like cells with a high replicative capacity in vitro [5]. But such important findings have seminal observations back to the 19th century [6] and to the work performed by Tavassoli and Crosby, a couple of scientists who first hypothesized inherent osteogenic potential associated with the bone marrow [7, 8]. Indeed, it is only thanks to the studies carried out not on intact fragments of bone marrow, but on single, isolated MSCs performed by Friedenstein and co-workers in the early 70s, that the breakthrough characteristic of MSCs was shown, when BM-derived cells at clonal density resulted in distinct colonies generated by single cells (the colony-forming unit fibroblastic, CFU-F). The clonal nature of such colonies, generated by cells indistinguishable from most hematopoietic cells except for their fibroblast-like appearance and rapid adherence to culture vessels, was later supported by genomic analyses, time-lapse photography, and Poisson distribution statistics [5, 9, 10]. Friedenstein himself, ten years after, renamed these cells as osteogenic stem cells or BM stromal stem cells [8], paving the road to Caplan for a more popular mesenchymal stem cell identification.

Several years after officially including MSCs into the adult stem cell category, Dr Caplan publicly admitted the importance of refining such multipotent cells as Medicinal Signaling cells, whose paracrine action rather than differentiation capacity leads to regeneration induction, a rejecting their stemness affiliation (Dr Caplan last lecture offered at the Cell Transplant and Regenerative Medicine Society in 2023 was unequivocally restraining scientists from calling MSC as stem cells) [11]. Unfortunately, Dr Caplan’s latest definition of MSCs as medicinal signalling cells was less than clarifying and setting the ultimate name [11]. Indeed, under the umbrella of medicinal secretive cells a wide amount of somatic and perinatal cells can be grouped and fulfill such paracrine criteria. Thus, the nomenclature changed officially and was consolidated by the International Society for Cell and Gene Therapy (ISCT) in 2019, dogmatically stating MSC as Mesenchymal Stromal Cells.

Contradictory and questionable results have been published and debated during the past decades, with final proof yet to come. Nonetheless, MSCs are considered one of the most promising cellular tools in regenerative and reparative medicine, and the most abundant source of cells for tissue engineering and cell therapy [11, 13, 14].

Fifteen years after delineating the release criteria to identify and release MSC products [12], operated by the Mesenchymal and Tissue Stem Cell Committee of the ISCT, these multipotent cells have consolidated their leading role in regenerative medicine. The exact mechanism(s) of action for somatic or perinatal MSCs is still largely unknown. However, it is well accepted that MSCs home in sites of injury or induce regenerative effects through secreted bioactive factors and trophic mediators. Indeed, leading experts have highlighted that a patient’s tissue-resident progenitor cells are the real fabricator for new tissue, supported and enhanced by MSC-secreted bioactive factors [13].

During recent years, MSCs have been found in vascular and avascular tissue, and different amount of MSCs have been extracted from any organ (i.e., liver, heart) and tissue (i.e., skin, tendon), including aborted fetuses and menstrual blood, but most reliable and abundant source are somatic tissues like BM aspirate, lipoaspirate and adipose tissue (AT), dental pulp (DP), but also perinatal tissues such as umbilical cord (UC), placenta and amniotic fluid (AF), [15–24]. However, several limitations and hurdles are documented in relation with primary MSC extraction and release. Concurrently, induced pluripotent stem cells (iPSCs) have been recently suggested as an alternative source for MSC, upon differentiation [25]. Interestingly, iPSC-derived MSCs (iMSCs) have been described with a rejuvenated phenotype, characterized by a greater proliferative capacity compared to standard tissue-derived MSCs. iMSC preclinical validations are still ongoing, with preliminary encouraging results supporting similar or even superior therapeutic activity in several preclinical settings [26]. Safety and tolerability for iPS-derived cells are obviously the primary concerns that phase I clinical trials are examining [26]. Conversely, tissue-derived MSC have been promoted to phase II and III trials during the years, leading to controversial results in the short- and long-term run.

MSCs act as a cebll reserve in mammalian tissues, supporting physiological cell turnover and initiating a regenerative response to acute injuries. These adult multipotent cells are non-specialized, self-renewing cells that can be induced to mature, both in vitro and in vivo, towards different somatic lineages. The past decade has seen an explosion of research directed toward a better understanding of the mechanisms of action for MSCs during rescue and repair of injured organs and tissues. Different sources of MSC have necessarily led to the optimization and validation of different methods of isolation, qualification, and expansion for such cell products. It is exactly this kind of diversity and multi-step standard operative procedures that experts and competent operators, here gathered, have compared, and summarized. These are crucial for generating MSC products from perinatal and somatic sources.

Perinatal sources

The placenta is a transient internal organ that plays various crucial roles in supporting the development of the fetus. Its functions include the promotion of gas exchange, nutrient and waste transport, maternal immunoglobulin transport, and secretion of hormones critical for fetal growth and development. These exchanges and transfers occur due to diffusion gradients between fetal and maternal blood, the latter of which bathes the chorionic villi in the intervillous space of the placenta. The human placenta is a temporary organ composed of cells and tissues with two distinct genetic backgrounds, one from the mother and one from the (semi)allogeneic fetus. Perinatal tissues include both the placenta and the umbilical cord connecting the fetus to the maternal support and the blood flowing in it. So, perinatal tissues comprise both fetal and maternal portions. The amniotic fluid, the umbilical cords, the amnion, and chorion membranes are of fetal origin, while the different regions of the decidua are maternal. The cord blood outflowing from the fetus (arteries) carries fetal-derived blood, while the umbilical vein is the sole venous vessel in our body vehiculation oxygenated blood received from the mother and “filtered” by the placenta. Both fetal and maternal layers contain different cells [27], including MSCs that can be isolated selectively from different parts upon surgical dissection and processed separately.

Umbilical cord stem cells (UCSCs)

The umbilical cord (UC) is a 3-vessel conduit connecting the fetus to the placenta, supplying nutrients and gasses until childbirth. The UC (also known as funiculus umbilicalis) has been considered a useless medical waste for centuries, together with the rest of the placenta. During the last decades, these tissues have re-gained attention and validated in several medical applications. Historically, the human UC is the most used perinatal material, both in terms of decellularized tissue and cells isolated from such a tubular structure at birth. Three different progenitor and multipotent cells can be isolated from full-term UC: epithelial cells paving the surface of the cord, in direct contact with the amniotic fluid and the fetus; endothelial cells paving all the three coiling vessels; stromal cells embedded in the thick stroma surrounding such vessels (commonly known as Wharton Jelly; WJ) or in direct contact with the vessels (also known as pericytes). Several applications have benefitted from the isolation of UC stromal cells, as a whole or from dissected compartments (subendothelial, perivascular, intervascular and sub amniotic regions) [28–32]. UC-derived MSCs have been largely translated into medical practice for several medical conditions. UCSCs or WJ-derived MSCs (WJ-MSC) have been largely described and characterized [33], sharing identity profiles in common with somatic MSCs [12] as well as multipotency and several other stemness characteristics that have led to their identification as stem cells for more than 20 years.

WJ-MSC isolation from umbilical cord

Two distinct approaches have been largely described for the isolation of human MSCs from umbilical tissue: the isolation and the enzymatic digestion method [28, 33, 34]. McElreavey and collaborators, in 1991, were the first to describe an efficient method to extract fibroblast-like cells from an intact UC. McElreavey’s method relies on cell capacity to migrate out of the tissue spontaneously. In such a procedure, the WJ tissue is exposed by mincing the cord into small pieces of tissue, seeded on an untreated culture vessel, surrounded by a culture medium, to support cell migration and attachment to the culture dish [35]. Between 2003 and 2007, several reports described revised approaches, aimed to generate a larger amount of WJ-MSCs within a few hours after collection. Most of the methods start from the same initial surgical approach, where the WJ is mechanically exposed. In a revised method, one surgical incision is performed longitudinally along the cord, exposing the internal stroma and vessels. Some authors reported the beneficial effects of having the WJ spread on mesh gauze or a suitable carrier membrane (e.g. nitrocellulose), with additional mincing aimed to generate small fragments of tissue. A few reports have supported such a cell manufacturing method, describing the proliferative, structural, and differentiation properties of WJ-MSC after explant [36, 37]. In 2007, Friedman and co-authors improved the explant method by mechanical removal of blood vessels, with the intent to prevent contamination of endothelial and blood cells in the final cell product [38]. Sarugaser and Davies were the first to describe the selective isolation of a cell subpopulation from the perivascular region of the cord [39]. The human umbilical cord perivascular vein (hUCPV) cells can be found in the surrounding proximity of both vessels, embedded into the WJ. Such cells have previously not been selectively released using the explant method. Indeed, Friedman’s approach allowed extraction of such a population by removing the vessels before culture. Perivascular cells in the UC have been described and characterized by identity and functional markers [32] commonly identified on mesenchymal phenotype, suggesting a specific role as human allogeneic cells source for cell therapies [39]. Significant differences can be found in explant protocols, affecting the quality of isolation. For example, we can find that different outcomes in WJ-MSC isolation rely on different sizes of explant fragments [40], the use of particular culture vessels (plates, petri dishes, or flasks), or the supplementation of specific additives/growth factors in the culture medium [41]. Another critical parameter affecting cellular yield relies on the migration/adhesion time granted to cells before the first medium change [40, 42, 43]. The second largely used method to efficiently isolate WJ-MSC is based on extracellular matrix digestion and cell release driven by specific enzymatic solutions. The main used enzymes are collagenase, mixture of collagenase and hyaluronidase, with or without trypsin, or a mix of collagenase, dispase, and hyaluronidase [32] (Table 1). Salehinejad et al., compared different enzymatic preparations in terms of efficacy to release of UCSCs, and showed that different formulations may significantly affect the purity and growth capacity of isolated MSC populations. A mixture of collagenase and trypsin was proven superior to any other enzymatic solutions, when applied to the isolation of human WJ-MSC [44]. For a long time, the enzymatic formulations used for primary cell isolation procedures comprised of minimally purified biochemical products. Enzymatic solutions derived from Clostridium histolyticum culture supernatants containing collagenase, neutral proteases, endotoxin, and other enzymes [45]. The commonly used commercial collagenase products can be subdivided into three grades: crude, enriched, or purified collagenases. These products differ by the degree of purification, a critical step resulting in specific activity of the collagenase enzymes. The main disadvantage of early 2000s protocols relied on poorly defined enzymatic solutions, characterized by a large lot-to-lot variability in enzymatic activity and endotoxin, affecting the broad range of incubation times, between a few minutes (15–30 min) to several hours (up to 16 hours) (Table 1 summarized different protocols using different enzymatic solutions, concentrations, and incubation time). Such variability raises the risk of degradation of the extracellular lamina, preventing MSCs from adhering to the surface of culture vessels after enzymatic digestion [28, 46]. Only a few studies have directly compared explant UC cultures to enzymatic isolation strategies [46, 47]. Since there is currently no standard protocol, researchers revised and improved the isolating protocols by introducing additional washing steps (to remove blood) or surgical removal of blood vessels [46, 48, 49].

Table 1.

Enzymatic digestion protocols used for UC and UCB-MSCs isolation

Enzymatic solution	Conditions	References
0.5 mg/mL Collagenase type I	18–24 h at 37  °C	[50]
1 mg/mL Collagenase type I	18–24 h at 37  °C 18–24 h at RT 1 h at 37  °C	[51] [52] [53]
0.2 mg/mL Collagenase type I  + 0.2% Trypsin	24 h at 37  °C  + 30 min at 37  °C	[54]
2 mg/mL Collagenase  + 2.5% trypsin	16 h at 37  °C  + 30 min at 37  °C	[55] [56]
1 μg/mL collagenase type B	4 h at 37  °C	[57]
1 mg/mL Collagenase type B  + 0.25 mg/mL Trypsin	3 h at 37  °C  + 15 min at 37  °C	[44]
1 mg/mL Collagenase, 300ug/mL Hyaluronidase  + 0.25 mg/mL Trypsin	2 h at 37  °C  + 15 min at 37  °C	[20, 58, 59]
300U/mL Collagenase, 1 mg/mL Hyaluronidase  + 0.1% Trypsin- EDTA	1 h at 37  °C  + 30 min at 37  °C	[59]
300U/mL Collagenase, 1 mg/mL Hyaluronidase  + 0.25% Trypsin–EDTA	45 min at 37  °C  + 15 min at 37  °C	[60]
0.8 mg/mL Collagenase type IV, 0.2 mg/mL Hyaluronidase, 1.38 mg/mL Dispase  + 0.25% Trypsin- EDTA	45 min (× 2) at 37  °C  + 20 min at 37  °C	[61]
2.5 mg/ml Trypsin, 0.2 mg/mL EDTA	30 min at 37  °C	[44]
150 USP U/mL Hylenex + 5 mg/mL Liberase	1.5–2 h at 37  °C	[56]

In summary, explant protocols are cost-effective and relatively easy to perform but result in low yield and unreliable outcome based on the MSCs ability to migrate out of the UC tissue and adhere to the surface of culture vessels [41]. Enzymatic digestion may increase cell recovery per gram of processed tissue, but it also affects cell adhesion and homogeneity, particularly in prolonged enzymatic exposure.

Stromal cell isolation from umbilical cord blood

Multipotent MSCs have also been detected in umbilical Cord Blood (UCB-MSC), leading to a subclass of MSC from a different origin but similar characteristics, such as anti-inflammatory, immune-modulatory, anti-oxidative stress, anti-apoptotic and more [62]. Historically the cord blood collected from perinatal tissues extracted after delivery has been a reliable source of hematopoietic stem cells (HSCs). The blood flowing into the umbilical cord has been the first perinatal product collected and infused in patients: in 1974 it was first reported as a source of HSCs and progenitor cells and infused [63, 64] and, in 1988, infused in a child with Fanconi anemia [64]. Hematopoietic stem cells (HSC), CD34-positive cells are routinely collected and banked for cell therapies. Usually, UCB is collected from full-term deliveries by venipuncture immediately after cord clamping and before placenta release. UCB can be recovered in high volumes and based on the thickness and length of the conduit, an approximate volume of 50-100 mL of blood can be harvested after delivery [65]. UCB is commonly stored in bags containing an anticoagulant and processed for HSC enrichment or eventually for MSC isolation. In both procedures, an initial purification of the mononuclear cells (MNCs), isolated by density gradient centrifugation, is performed (Figure 1). To generate UCB-MSC, all the MNCs are seeded and allowed to adhere overnight, to perform an initial selection of highly adherent cells as MSC [66–68].

Fig. 1.

Schematic representation of UCSCs isolation from UC and UCB

Nevertheless, the amount of MSC in the cord blood is quite low, and the established procedure to collect CD34 cells has not supported an additional selective manipulation for MSC. Similarly to other tissues, UCB can be collected, processed, and cryogenically preserved, granting CB-MSC off-the-shelf [69–71]. Moreover, thanks to the presence of the placental barrier, a standard processing in closed systems, CB-MSCs have a lower risk of bacterial and viral infections than MSCs isolated from somatic sources. Perinatal MSC have been described as excellent candidates for allogeneic transplants due to immune-modulatory properties and paracrine effects [72] and low/null risk for graft versus host disease (GvHD) [73]. Adherent cells are cultured until the outgrowth of fibroblastoid cell colonies appears. After reaching a sub-confluent stage, cells are harvested, re-seeded in culture and exposed to cell expansion in an MSC-selective medium. UCB-MSC, as any other MSC, is characterized according to standard criteria: presence of static cell markers (CD73, CD90, CD105), plastic adherence, and differentiation potential into mesoderm lineages such as osteoblasts, adipocytes and chondroblasts [12]. Such release criteria were established almost 20 years ago by the Mesenchymal and Tissue Stem Cell Committee, part of the International Society for Cell and Gene Therapy.

UC- and UCB-derived MSC culture conditions

WJ-MSC, once released from UC, are cultured with culture media supporting MSC proliferation. The first standard medium used was the canonical Dulbecco’s Modified Eagle Medium (DMEM), rapidly replaced with formulation with low glucose concentration (1.5 g/L or below), such as DMEM/Ham’s F12 nutrient medium (DMEM/F12), low glucose DMEM, or alpha Minimal Essential Medium (α-MEM) (Table 2). Standard cell culture media are commonly supplemented with high concentrations of fetal bovine serum (FBS or FCS) (10%). Moreover, for UCB-MSCs culture, the expansion medium consists of Iscove modified Dulbecco (IMDM) medium supplemented with 20% FBS [66]. Recently, Hassan et al. proposed an alternative culture method to replace FBS, which has an important limitation related to its animal origin, with cord blood serum (CBS) and human platelet lysate (hPL) [74, 75]. UCSCs are usually stored at 37 °C in a humid atmosphere with 5% CO₂ and fresh medium is replaced every 2 days and 80% confluency is achieved for optimal cell harvesting.

Table 2.

Culture media for UCSC and UCB-MSC propagation

Basal medium	Supplements	References
UCSC
α-MEM	10% FBS + 1% L-glutamine + 1% nonessential amino acid (NEAA), + 20 IU/mL basic fibroblast growth factor (bFGF) 20% FCS + 2 mM L-glutamine + penicillin/streptomycin 10% hPL + 2 mM L-glutamine + Penicillin/streptomycin 10% FBS + 2 mM L-glutamine + penicillin/streptomycin 10% human serum + gentamicin	[76–80]
DMEM/F12	10% FBS + penicillin/streptomycin/amphotericin B 10% FCS + penicillin/streptomycin 10% CBS + penicillin/streptomycin	[44, 53, 81]
DMEM	10% FBS + 2 mM L-glutamine + penicillin/streptomycin/gentamicin 10% FBS + penicillin/streptomycin 10% FCS + penicillin/streptomycin 20% FCS + penicillin/streptomycin/amphotericin B 15% FBS + Glutamax + MEM-NEA + MEM vitamins + penicillin/streptomycin	[82–87]
DMEM-HG	10% FBS + penicillin/streptomycin/amphotericin B 20% FBS + 16 ng/ mL bFGF + 1:200 insulin-transferrin-selenium (ITS) + penicillin/streptomycin/amphotericin B	[43, 88]
DMEM-LG	10% FBS + penicillin/streptomycin 10% FBS + 2 mM L-glutamine + penicillin/streptomycin/amphotericin B 10% FBS + 2 mM L-glutamine + NEA + penicillin/streptomycin/amphotericin B 10% FBS + 2 mM L-glutamine + 5 ng/mL bFGF + penicillin/streptomycin/amphotericin B 10% FBS + 10 ng/mL bFGF + penicillin/streptomycin	[41, 52, 89–94]
DMEM-KO	10% FBS + 2 mM L-glutamine + penicillin/streptomycin	[95]
RPMI	10% FBS + penicillin/streptomycin/amphotericin B/Ciprofloxacin
UCB-MSC
IMDM	20% FBS + 10 ng/mL bFGF + penicillin/streptomycin, and 2 mM L-glutamine 10% FBS or (5%,10%) platelet-rich plasma (PRP) + 10 ng/mL EGF + penicillin/streptomycin/amphotericin B	[38, 96]
DMEM	10% hPL + penicillin/streptomycin 10% cord blood serum (CBS) 10% FBS	[74, 75]
DMEM-LG	10% FBS + penicillin/streptomycin	[90]
Culture conditions	37  °C in a humid atmosphere with 5% CO2	[74]
Cryoconservation	10% DMSO in complete medium or FBS	[74]

MSC from amniotic membrane

As previously stated, MSC can be released and collected from many other parts of the human placenta, at the end of pregnancy (or earlier if needed). Mesenchymal stromal cells have been largely described as embedded in the amniotic membrane (AM) surrounded by rich amount of extracellular matrix. The amnion is a thin, avascular membrane composed of a compact layer of cuboidal and columnar epithelial cells, which are in contact with the amniotic fluid on the external side and attached to a basal lamina on the inner side. This lamina is connected to the amniotic mesoderm, a layer of extracellular components (collagens and fibronectin) into which is embedded a network of dispersed fibroblast-like mesenchymal cells.

Human AMSCs are isolated from human amnion that, like other placental tissues, would normally be discarded post-delivery. For sterility purposes, placentas are normally obtained from cesarean section; however, theoretically, all placentae could be useful for MSC isolation. Amnion-derived MSCs have been shown to express major cartilage components after chondrogenic induction, with deposition of collagen II after in vivo implantation into the abdominal muscle of mice [97]. Since there is no maternal contribution to the amnion membrane, amnion-derived MSCs are entirely fetal and characterized by low immunogenicity.

Isolation of AMSCs

Human amnion cells can be isolated from the amnion membrane either as a heterogenous suspension composed by epithelial and stromal cells, or through a multi-step procedure based on selective enzymatic solutions developed to release epithelial and mesenchymal cells effectively (Figure 2).

Fig. 2.

Schematic representation of isolation and culture of AMSCs. The amniotic membranes separated from chorion, fragmented, and immediately placed in culture or subjected to enzymatic digestion. After centrifugation the resulting pellet is placed in culture together with culture medium

In most protocols, trypsin is used for isolation of epithelial cells, followed by a second step with collagenase to release AMSCs [26, 97–105]. However, other digestive mixtures have been proposed and tested, including collagenase enriched with proteases (DNase) [101, 106, 107] or dispases [101, 103, 108, 109]. The removal of epithelial cells paving the inner layer of AM has been described as beneficial to generate a large yield in AMSC. A considerable number of epithelial elements may still be attached to the membrane before collagenase digestion, and this may result in the presence of epithelial cells in the AMSC preparations. Additional steps in washing and purification by density gradients have been shown to generate a homogeneous suspension, further characterized for MSC identity markers [12] and some surface proteins commonly associate with pluripotent stem cells largely described as characteristics in amnion-derived cells (such as SSEA-3 and SSEA-4, but lacking TRA-1-60 and TRA-1-81 [110, 111]. Alternatively, it has been reported that the “purity” of the preparation may be enhanced by exploiting the slow adhesion of epithelial cells to culture substrates, as opposed to the fast attachment of hAMSCs, it is good practice to change the culture medium 1 to 2 h after plating. This will remove many of the undesired cells that might be present in the preparation. The morphological differences between the two cell types will allow you to easily determine whether heterogeneity in the preparation.

Expansion of AMSCs is possible for at least 5 passages without morphological alterations [106, 108, 112]. Studies have shown that maintaining AMSCs in a controlled environment with specific growth factors and nutrients can support their proliferation without compromising their morphology. Some groups have even kept AMSCs in culture for 15 to 20 passages before reaching senescence [103, 113].

Cells from amniotic fluid

Another perinatal tissue frequently harvested during or at the end of pregnancy is the amniotic liquid. The Amniotic Fluid (AF) serves as a protective liquid for the developing embryo, providing mechanical support and the required nutrients during embryogenesis. The major component of AF is water and soluble mediators and nutrients initially released by amniotic epithelial cells and later released by the fetus. AF composition varies significantly throughout pregnancy. At the beginning of pregnancy, the amniotic osmolarity is like the fetal plasma. After keratinization of the fetal skin, amniotic osmolarity decreases relatively to maternal or fetal plasma, mainly due to the inflow of fetal urine [114].

Amniocentesis has been used for many decades as a routine procedure for fetal karyotyping and prenatal diagnosis, allowing the detection of a variety of genetic diseases. Interestingly, AF has been largely described as a rich source of cells exfoliated by the fetus or released by the amnion membrane, equipped with cellular properties for potential use in a plethora of clinical settings. The Amniotic Fluid Stem Cells (AFSC) represent a heterogeneous population derived from the three germ layers. These cells share an epithelial origin and are derived from either the developing embryo, or from the inner surface of the amniotic membrane, which are characterized as amniotic membrane stem cells. The AFSCs are mainly composed of three groups of adherent cells, categorized based on their morphological, growth and biochemical characteristics. Epithelioid (E-type) cells are cuboidal to columnar cells derived from the fetal skin and urine, amniotic fluid (AF-type) cells are originating from fetal membranes, and fibroblastic (F-type) cells are generated mainly from fibrous connective tissue. Both AF- and F- type cells share a fibroblastoid morphology and the dominant cell type appears to be the AF-type, co-expressing keratins and vimentin [115–117].

AFSC isolation

Several studies have documented that human AFSCs can be easily obtained from a small amount of second trimester liquid, collected during routine amniocentesis, a procedure with spontaneous abortion rate ranging from 0.06% to 0.5% [118, 119]. Up to date, several isolation protocols based on the seeding and cultivation of cells have been reported, leading to enriched stem cell populations [120]. The isolation of AFSC and the respective culture protocols were summarized in a recent review by Klemmt et al (Figure 3) [121] and are divided into i) a single-stage cultivation protocol, in which the primary culture was not disturbed for at least 7 days until the first colonies appeared; ii) a two-stage cultivation protocol, in which unattached amniotic cells are collected after 5 days and further expanded; iii) cell surface marker selection for CD117 (c-kit receptor); iv) mechanical isolation of mesenchymal progenitor cell colonies formed in the initial cultures; v) short-term cultures for the isolation of fibroblastoid colonies. The resulting cell products shared a multipotent mesenchymal phenotype, exhibited higher proliferation capacity, and broader differentiation potential compared to adult MSCs.

Fig. 3.

Schematic representation of isolation and culture of AFSCs. Example of the isolation and culture of AFSCs. The amniotic fluid is centrifuged, and the resulting pellet is placed in culture vessel together with culture media for approximately 10–15 days, when the first colonies appear

Some authors reported issues in maintaining such properties after long-term culture, prevented to some extent by exposing AFSC to selected growth factors. Specifically, some axiomatic strategies include the use of large amount of FBS (20% vs 10% commonly supplemented), or by modified version of basic culture medium, such as Iscove's modified Dulbecco's medium (IMDM), [122] or AmnioMAX C100 [123], or even mixture of media (i.e., 88% αMEM mixed with Chang B basal medium) [124]. Apart from these peculiar approaches, any other modifications or approaches are quite similar to any other MSC protocols (e.g., supplementation with TGFbeta or other mitogenic factors).

AMSC and AFSC culture conditions

Early attempts to expand AMSCs ex vivo used standard culture methods (10% serum supplemented basal tissue culture medium) and isolated MSCs exhibited typical spindle morphology [125, 126]. Subsequently, several efforts to establish long-term culture conditions for AMSCs were based on the optimization of serum-supplemented culture media, with or without additional growth factors, to study the growth kinetics and stem cell phenotype of these cells (Table 3).

Table 3.

Culture media for propagation of AMSCs and AFSCs

Basal medium	Supplements	References
AMSCs
DMEM DMEM/F12	10% FBS/FCS	[125–127, 135, 136]
DMEM	FBS 10% ITS, HS, bFGF, L-Ascorbic acid FBS 10% + FGF-2 FBS 10% + EGF 10 ng/mL + Na-bicarbonate	[130–132]
α-MEM	FBS/FCS 10% FBS 20%	[100, 134, 137]
α-MEM	PAA, 15% FBS, 1% Penicillin/Streptomycin, 2% Chang C and 18% Chang B	[138]
AFSCs
DMEM	20% FBS	[118, 120]
Iscove's modified Dulbecco's medium (IMDM)	20% FBS 5 ng/mL bFGF	[139]
α-MEM	10% FBS, 1% GlutaMAX, 10% Chang B basal medium, and 2% Chang C supplement	[124]
AmnioMAX C100	10% FBS	[123]

In these works, the complete adherence of MSCs without any additional supplements is reported [108, 127, 128]; however, some researchers recommended using 20% FBS instead of 10% FBS to support cell adherence [129]. In some studies, the addition of growth factors (such as FGF, EGF), or ascorbic acid, or Na-bicarbonate to serum-supplemented culture medium was found to support the expansion of cells [130–132].

Standard culture medium for amnion or amniotic fluid cells is like UCSC (low glucose media, as αMEM) supplemented with 10-20% fetal animal serum. Similarly to what described afore, customary media (αMEM mixed with Chang C and Chang B formulations) have shown some level of benefit [133]. In works like these, the researchers demonstrated the ability of placenta stromal cells to achieve higher proliferation rates and maintain stemness characteristics (including multipotency) for several passages (up to sixth passages) [100, 134]. Once again, the replacement of animal-derived supplements with human products (platelet lysates or sera) are currently under investigation and validation.

Somatic sources

Bone marrow MSC (BM-MSC)

The bone marrow (BM) is considered the only permanent hematopoietic organ in humans, contained in the marrow cavity of the bones. Such hematopoietic tissue is divided into yellow marrow, where adipocytes prevail, and red marrow, where the marrow microenvironment is located. Active BM is rich in replicating HSC and precursors of mature blood cells; the prevalence of maturing erythrocytes determines a dark red color from which the denomination of red marrow derives. In the stroma of the BM are disseminated several stromal cells (BM-MSC). BM-MSCs represent approximately 0.01% of mononuclear cells (MNCs) and provide structural and functional support to hematopoietic stem cells (HSCs) [9, 128, 140]. The multipotent precursors of the BM stroma were the first MSCs to be identified [9]. Bone marrow is usually aspirated from the iliac, femur, tibia, and sternum bones and processed within a few hours to extract BM-MSCs [141, 142]. These cells are the subject of interest for their ability to migrate into damaged tissues and their active action in supporting repair, regeneration, and modulation of the immune response. The identity qualification for MSC was initially set on BM-MSC relying on cell adhesion to plastic (when cultured in presence of serological proteins), the expression of specific surface markers (co-expression of CD73, CD90, and CD105) and the absence of hematopoietic markers (CD45, CD34, HLA-II), the ability to differentiate into osteoblasts, chondrocytes, adipocytes [12].

BM-MSCs isolation

The most common and safe site to collect BM tissue is the iliac crest. Through a bioptic needle, a sample volume between 20 and 50 mL is obtained [143, 144]. The patient is subjected to general or local anesthesia; imaging guidance using fluoroscopy and computed tomography support the safety, success, and yield of BM sample (Figure 4). Once the BM tissue is collected, the sample is immediately processed and subjected to Ficoll density gradient centrifugation [145] which separates the MNCs from the polymorphonuclear and non-nuclear cells. MSCs can be identified based on their ability to form adherent colonies in vitro by designating these cells as colony-forming units (CFU-F) [146] or by advanced isolation techniques such as flow cytometry (Figure 4). However, a interindividual variability in cell adhesion efficiency has been largely reported. Furthermore, MSCs represent only a rare fraction of the initial cell population, which contains other abundant cell types that may adhere to standard cell culture surfaces [147]. Therefore, cultures of BM-MSCs initiated with unselected cells are heterogeneous and undesirable interactions between MSCs and non-MSC may occur. Based on flow cytometry, more sensitive techniques are then used, which exploits the surface markers expressed by BM-MSCs [143, 147]. BM-MSCs are fibroblast-like cells with crucial features such as multipotent differentiation potential, mielo-supportive capacity, anti-inflammatory and immunomodulatory properties, and extensive use in cell therapy and tissue repair [148–151].

Fig. 4.

Schematic representation of BM-MSCs isolation

BM-MSCs culture conditions

Data reported in the literature support the frequency of stromal cells equal to 1 every 18,000 MNCs in a BM aspirate. Consequently, the recovery of BM-MSCs obtained from each donor may largely vary, and it is therefore essential to resort to in vitro cell expansion [143, 152]. Moreover, automated culture systems, such as bioreactors, have been validated to for ex vivo expansion of human BM-MSC. In example, the Quantum system commercialized by Terumo BCT [153, 154] has been described support such critical step in GMP conditions, based on a functionally closed, automated hollow fiber bioreactor system designed to grow adherent and suspension cells. Such an automated system could potentially reduce costs and time [155]. Similar technologies commercialized by different biotech companies have been proposed and validated, with still some controversial outcome in efficiency for adherent cells like MSC. However, a proper comparison between commercial support is beyond the scope of this study. Regarding culture conditions, after separation by Ficoll density gradient, the MNCs are cultured with same culture conditions previously described. Briefly, DMEM/F12 or α MEM culture media supplemented with animal or human serological products, with the exclusion of antibiotics and antimicotic not allowed in GMP-grade conditions for clinical uses (Table 4). To minimize the risk of xenogenic protein-related allergies in patients, animal-free and serum-free media can be used. FBS can be replaced with autologous serum, platelet lysate, platelet-rich plasma (PRP), and growth factors without serum or commercial xeno-free media [156]. Culture formulations have been developed and commercialized for the culture of BM-MSCs [147, 150]. Compared with traditional culture media, human MSCs that were expanded in serum- and xeno-free media showed better expansion and multipotentiality and are suitable for producing large-scale functionally competent MSCs for clinical applications [140, 155, 156]. The mean yield of MNCs for each aspirate is approximately 4x10⁷ cells. Such mononuclear cells are resuspended in the specific culture medium and placed in an incubator at 37 °C and 5% CO₂ for 24h. After 24 hours of culture, the suspended cells are transferred to a new flask. This step is essential because it allows for an initial purification of the cells from monocytes, which adhere to the plastic starting from the first 24–48 hours. Then, the flask is placed again in the incubator at 37 °C. Once confluence is reached (2–3 weeks), the cells are detached, and a cell count is performed. The cells can be re-seeded after immunophenotypic characterization.

Table 4.

Examples of culture media implemented in research-grade production of BM-MSCs

Basal medium	Supplements	References
α-MEM	10% platelet lysate, 2i.u/mL Na-heparin 1% Penicillin/Streptomycin, 10%, platelet lysate, 2i.u/mL Na-heparin 10% FBS + 100 U/mL penicillin, 0.1 mg/mL Streptomycin, and 100 μmol/L ascorbic acid-2 phosphate 10% FCS + 1% Penicillin/Streptomycin + L-Glutamine	[156–159]
DMEM or DMEM/F12	10% FBS + Penicillin/Streptomycin + L-Glutamine	[147],[160]
DMEM-LG	10% FCS + 1% Penicillin/Streptomycin	[159]
IMDM	9% FCS, 9% HS, 100 U/mL Penicillin, 100 μg/mL Streptomycin, and 12 μM L-Glutamine	[161]
Culture conditions Cryoconservation	37  °C in a humid atmosphere with 5% CO2 10% DMSO in complete medium or FBS	[162]

Dental pulp cells (DPSCs)

MSC extracted from dental pulp (DPSC) are characterized by properties similar to BM-MSCs, and they were firstly isolated by Gronthos and colleagues in 2000 [23]. Although the presence of such multipotent cells in dental pulp was previously reported by Yamamura in 1985 [163]. Since then, the presence of different types of MSC populations in teeth has been described, which, depending on the site of collection, have been referred to as DPSCs, human exfoliated deciduous tooth stem cells (SHEDs), ligament stem cells (PDLSCs), dental follicle stem cells (DFSCs), apical papilla stem cells (SCAPs), and gingival MSCs (GMSCs) [164–168]. DPSCs can be isolated from human pulp tissues both in deciduous and adult teeth (removed for orthodontic purposes, impacted teeth, and supernumerary teeth) and characterized based on surface marker expression (22, 168). DPSCs are multipotent cells equipped with all MSC properties [169–171]. Such cells are embedded in the central cavity of each tooth of healthy subjects’ pulp of human teeth, a soft and highly vascularized connective tissue consisting of four zones: the peripheral odontogenic zone, the intermediate cell-free zone; the cell-rich zone; the pulp core [172]. DPSCs originate from ectomesenchyme, derived from migratory neural crest stem cells during early development [173]. Such an origin has related to their neurogenic capabilities, reported superior to other MSCs, envisioning their use in neurodegenerative diseases [174].

DPSCs isolation

Given the importance of DSCs in regenerative and biomedical research, several methods for DPSC isolation and expansion have been published. Starting from the Gronthos' protocol, several researchers have revised dental pulp cell isolation, characterization, differentiation, and storage procedures providing a significant revision of both non-invasive techniques and ethical constraints [175–177]. The main factors that can influence DPSC properties, during the experimental procedures, are the selection of the tooth type (e.g., molar, third molar), the stage of tooth development, the characteristics of the donors (e.g., sex, age, lifestyle), tooth transport and short-term storage [22]. In this regard, Perry et al., reported as DPSCs remained viable up to five days after tooth extraction if preserved at 4 °C in phosphate-buffered saline (PBS). However, cell viability decreased dramatically after 24 h of storage [178]. The three main procedures described in the literature for the DPSC isolation are based on (i) tissue explants (OG); (ii) enzymatic digestion (ED); (iii) mechanical extraction (or combination of mechanical and ED procedures) [179–181] (Figure 5). The OG method is technically simpler rather than others, since consists of placing pulp fragments (pieces of 1-2 mm³) directly into the culture dish and let the stromal cells migrating out and adhere on culture vessel [179, 182, 183]. As previously mentioned, the enzymatic digestion of stromal extracellular tissue has been documented efficient in releasing single cell. Different enzymatic formulations, based on collagenases (type I, II, IV), dispase, trypsin, or accutase have been described [22, 184]. The enzymatic digestion remains the most used method to obtain DPSCs [185–188]. Some researchers compared OG and ED, reporting higher proliferation and differentiation rate in MSC released after enzymatic digestion [180, 185, 189]. ED remains the most used method to obtain DPSCs [185–188]. Based on the first Gronthos procedure (Figure 5), the pulp tissue is gently separated from the crown and root and then digested in an enzymatic solution of 3 mg/ml collagenase type I and 4 mg/ml dispase, the most used enzymatic cocktail [190], for 1 hour at 37 °C [22]. Several alternative digestion protocols for DP dissociation have been reported (Table 5).

Fig. 5.

Schematic representation of DPSCs isolation

Table 5.

Enzymatic digestion protocols used for DPSCs isolation

Enzymatic solution	Conditions	Teeth	References
3 mg/mL collagenase type I, 4 mg/mL dispase	1 h at 37  °C 30–60 min at 37℃	Third molars and molars	[22, 23, 176, 189, 191–193]
4 mg/mL collagenase type I, 4 mg/mL dispase type II	24 h at 4  °C	Deciduous teeth	[168, 194, 195]
4 mg/mL collagenase type I, 2 mg/mL dispase	1 h at 37  °C	Incisors	[196]
0.2 mg/mL collagenase type I, 2 mg/mL dispase	70 min 37  °C	Third molars	[197, 198]
1 mg/mL collagenase type I, 2.4 mg/mL dispase	1 h at 37  °C	Deciduous teeth, adult molars	[199]
3 mg/mL collagenase type I	1 h at 37  °C	Third molars	[200, 201]
2 mg/mL collagenase type I, dispase	30 min at 37  °C	Deciduous teeth, third molars	[202]
0.2% collagenase type I	1 h at 37  °C	Incisors, canines, molars, deciduous teeth	[203, 204]
0.2% collagenase type II	30 min at 37  °C	Third molars	[205]
1 mg/mL collagenase type I	30 min at 37  °C	Third molars	[184]
0.1% collagenase type IV	15–30 min at 37  °C	Third molars	[169, 205, 206]
0.2% trypsin	5 min at 37  °C	Third molars	[187]
1 mg/mL accutase solution	30 min at 37  °C	Third molars	[207]
Liberase	40 min at 37  °C	Third molars	[208]

DPSCs culture conditions

To improve primary culture establishment, a good cell attachment in the plastic dish is essential, which can be eventually enhanced through the pre-coating of the plastic surfaces with ECM proteins, peptide-modified surfaces, synthetic polymer cations or culture-treated surfaces [209]. Several ECM molecules, such as recombinant vitronectin, laminin-511 and laminin-521/E-cadherin, and synthetic polymers for example, polyethyleneimine and poly [2-(methacryloyloxy) ethyl dimethyl-(3-sulfopropyl) ammonium hydroxide] (PMED-SAH) have been reported to support the long-term culture of pluripotent stem cells [210–213]. Some authors, report that ECM components can enhance long-term DPSC culture, like in other stem cells [214, 215]. Kim et al., argue that ECM provides more than a substrate for attachment but also, plays a key role in signaling events essential to maintaining the stem cell niche [216]. Once enzymatically released DPSC can be resuspended and seeded in different Media based on commercial formulations such as αMEM, high- or low-glucose DMEM, DMEM/F12, nd DMEM-KO media [181] (Table 6). Standard cell culture media are commonly supplemented with high concentrations of FBS (10-20%) to stimulate growth and proliferation [217]. However, although the in vitro use of FBS is critical, as it provides many components required for cell growth, metabolism, and proliferation [217], some researchers have raised some concerns about the use of FBS for the DPSC isolation direct to human cell therapy due to the high risk of contamination, severe immune reactions, and therapeutic outcome controversies [218–220]. For these reasons, researchers are increasingly oriented towards the study of alternative compounds to FBS such as the use of culture media supplemented with a human-serum origin, umbilical CB serum and human hPL or, alternatively, chemically defined serum-free media [199, 218, 221–223]. DPSCs are usually stored at 37 °C in a humid atmosphere with 5% CO₂ and fresh medium is replaced every 2 days and 80% confluency is achieved for optimal cell harvesting [169, 206, 224]. During cell culture, different environmental conditions, such as temperature, humidity, pH, and oxygen levels, which play a crucial role in cell growth, must be considered. Recently, 21% O₂ has been shown to reduce cell proliferation and promote the activation of antioxidant defenses in DPSCs compared with 3% O₂ [225].

Table 6.

Culture media for DPSC propagation

Basal medium	Supplements	References
α-MEM	Platelet lysate (PL) 20% FBS + penicillin/streptomycin 10% FBS + 2 mM L-glutamine + 100 units/mL Penicillin and 100 µg/mL Streptomycin 10% FBS + 2 mM l-glutamine + 100 µM l-ascorbic acid-2-phosphate + 100 U/mL penicillin-G + 100 µg/mL streptomycin, and 0.25 µg/mL fungizone 10% FBS + 1% penicillin/streptomycin solution 20% FBS + antibiotics (100 IU/mL Penicillin and 100 µg/mL Streptomycin 15% FBS + 100 U/mL Penicillin/Streptomycin	[192, 208, 215, 226–230]
DMEM	10–20% FBS + penicillin/streptomycin	[203, 231]
DMEM/F12	10% FBS + penicillin/streptomycin	[193, 204]
DMEM-LG	10% FBS + penicillin/streptomycin 20% FBS + 2 mM L-glutamine + penicillin/streptomycin 10% FBS + 1% L-glutamine + 100 U/mL penicillin + 100 µg/mL streptomycin	[169, 201, 202, 232]
DMEM-KO	10–20% FBS + penicillin/streptomycin	[204, 233, 234]
Culture conditions Cryoconservation	37  °C in a humid atmosphere with 5% CO2 10% DMSO in complete medium or FBS	[169, 178, 206, 235–237]

Adipose-derived stem cells

Adipose tissue (AT) is derived from the mesoderm during embryonic development, and it is present in every mammalian species located throughout the body, and it is functionally classified into two categories: white and brown. Deposits of brown AT are responsible for energy production and are typical of fetal and neonatal life. Brown AT, in adult life, is converted into white AT with the primary function of energy reserve (through lipid storage), thermal insulation, and production of hormonal factors [238]. Furthermore, white AT can be classified into visceral adipose tissue (ATvis) and subcutaneous adipose tissue (ATsub) according to the body regions where it originates. ATvis is localized in organs such as the intestine, stomach, liver, and pancreas, while ATsub is more present in the abdomen, buttocks, and thighs [238]. The long-held belief about AT was that it was a relatively inert tissue in terms of biological activity, and it was believed that its central role was as energy storage. However, such old theory was changed thanks to the discovery of the large abundance of adult stem/stromal cells in this tissue. Adipose-derived Stromal Cells (ADSCs) are a subpopulation of MSCs and the actual progenitors of fat cells/adipocytes [239], largely used as source of MSCs [240] granted by large abundance (particularly in Western Countries), easily accessible, and renewable source of adult stem cells. ADSCs are a cell population of great interest in medical research because they have no ethical limitations, and it is possible to isolate them from an accessible source, AT, through minimally invasive procedures. They have been used in tissue engineering to repair damaged tissues and have been explored for their potential in wound healing and scar reduction for several diseases. Furthermore, the estimated percentage of ADSCs residing in AT deposits is approximately 1%, higher than BM [241, 242]. The most common source of ADSCs is the white ATsub, and these ADSCs have a greater proliferative capacity and adipogenic potential than those deriving from ATvis [243]. The biological features of these cells depend on the donor’s physiology and health status, isolation procedure, culture conditions and different protocols used [244]. Epicardial adipose tissue, the fatty deposit surrounding the heart and located beneath the pericardium, is also a source of ADSCs. The so-called epicardial ADSCs (eADSCs) shows a heightened potential to develop into heart muscle cells compared to other subtypes of ADSCs [245], such as those derived from ATvis or ATsub, and significant paracrine effects that promote angiogenesis and modulate the immune system. Therefore, the mobilization of resident eADSCs could be a potential therapeutic strategy to stimulate cardiac regeneration in patients with heart failure [246].

ADSCs isolation

Adipose tissue is collected through biopsy or liposuction surgery [247, 248] (Figure 6). The initial method for isolating fat cells from AT was pioneered by Rodbell and colleagues in the 1960s and were based on biopsy [248]. Briefly, rat fat pads were minced with a scalpel and washed extensively to remove contaminating hematopoietic cells. Subsequently, the tissue fragments were incubated with collagenase, followed by centrifugation to recover the fat cells. To date, the method of processing AT taken by biopsy for ADSC isolation is not massively changed; the tissue is chopped into small pieces using sterilized scissors or a scalpel to facilitate subsequent tissue digestion. If the sample volume of AT is of the size between 150-250 mg, it is recommended to increase the volume of collagenase solution (up to 400 μl for an equivalent of 150-250 mg) to improve the efficiency of tissue digestion[249, 250]. Among the techniques for harvesting AT by liposuction, there is the method described by Coleman, in which the AT sub sample is taken from areas where it is abundant, such as hips, buttocks, and abdomen. These areas are injected with saline and local anesthetic, such as lidocaine, and then a cannula is inserted, avoiding damage to the aspirate tissue. Next, the harvested AT, the harvested amount of which depends on many variables, can be subjected to a centrifugation process to remove the nonviable components and to obtain tiny parcels of fat [251]. In 2001, Zuk and colleagues were the first to demonstrate that fat tissue could be a source of autologous adult stem cells. They processed human AT obtained by liposuction to recover a fibroblast-like population of cells, first called processed lipoaspirate (PLA). Such PLA cells showed low levels of senescence, mesenchymal origin and ability to differentiate into adipogenic, chondrogenic, myogenic, and osteogenic cells in the presence of lineage-specific induction factors, this identifying them as ADSCs [17]. Since the countless studies and protocols have been performed to isolate ADSCs from AT, and many different protocols have been developed. Manual and automated methods can be differentiated, which can be further divided into enzymatic (Table 7) and non-enzymatic processes. All the protocols have in common an initial phase of extensive washing of the starting material (AT) to clean it from blood and oily components. Such a washing step is generally followed by an enzymatic phase to digest the ECM and release the cellular components. This step is generally performed at 37 °C using enzymes such as collagenase (type I or II), dispase, or trypsin. The enzymatic incubation may take a few hours and later neutralized with protein-enriched solution. The process continues with a mechanical phase aimed at isolating the stromal vascular fraction (SVF), a heterogeneous cell population composed of ADSCs, endothelial cells, erythrocytes, lymphocytes, monocytes/macrophages, fibroblasts, and preadipocytes. Digested tissue is first filtered through a nylon filter to remove the various cellular debris present, and SVF is collected by centrifugation. Subsequently, an erythrocyte lysis step is frequently included to avoid erythrocyte contamination and to decrease the number of cells with hematopoietic origin. Hypotonic sodium chloride and ammonium chloride are generally used as erythrocytes lysis reagents, since they have no negative effects on ADSC survival and differentiation [252]. Sodium chloride is preferred for clinical application, since it is convenient and cost-effective, it ensures the highest ADSC proliferative potential and is safe for patients, while ammonium chloride may be harmful to the immune system and represent a threat to patient safety [253]. There are numerous protocols through which it is possible to obtain SVF from AT, but these are not always standardized and reproducible in terms of yield. It is known that minimal variables in manual protocols lead to the isolation of different cells according to the procedure used [254]. These variants are represented by the type of reagents used, such as the enzymatic solution, and by the time or temperature of application. Other modifications may include mechanical comminution of the AT, centrifugation parameters, and the use of buffer solution for red cell lysis. For these reasons, in addition to conventional manual methods of tissue digestion, various closed automated systems have been produced and disseminated recently, which can isolate the SVF from AT, maintaining the sterile conditions and ensuring a high-quality process. Systems focus on the automated isolation of SVF from liposuction aspirates by collagenase-based digestion. The most important commercial systems using this method include AdiStem (AdiStem Pty Ltd, China) [255], Sepax (Biosafe Group SA, Switzerland) [256], Celution (Cytori Therapeutics Inc, Usa) [257]. All these systems have been tested and show real effectiveness. However, the systems can differ with respect to the yield of nucleated cells, the cell viability, and the percentage of stem cells among the SVF subpopulations. A colony-forming unit fibroblast assay, together with the subsequent differentiation assays, should be performed to better characterize the stem cell potential of the cell products obtained from the automated systems. Enzymatic digestion may have adverse effects on cell survival and the expression of surface antigens, this influencing the safety and efficacy of the cellular product obtained [43]. The use of enzymatic versus non-enzymatic methods is also generally accompanied by high costs. Conversely, non-enzymatic methods give more significant guarantees in terms of the safety of the cellular product, however they cannot ensure the yield in terms of SVF achieved by enzymatic methods. Specifically, the lower yield is due to the low ability of these methods to break down ECM compared to enzymatic methods. Therefore, a large proportion of cells belonging to the SVF remain trapped in larger and coarser portions of the tissue that are discarded during AT processing [258]. The non-enzymatic methods base their principle of action on the use of mechanical forces such as pressure, centrifugal force, shear force, radiant force, or sonication [259]. Manual processes achieve mechanical digestion of AT fragments by pipetting, shaking, filtration, and centrifugation. Raposio et al. described a protocol for mechanical isolation based on the use of a vibrating shaker followed by a centrifugation step and the final collection of cell pellets through an automated pipetting system. This method allowed to isolate a conspicuous number of ADSCs in approximately 15 minutes; moreover, it does not require collagenase, serum or other animal-derived reagents and can be performed within the operating room, with minimal manipulation [260]. Various ADSC automated isolation systems on the market also exploit these mechanical forces, such as Puregraft (Bimini Technologies LLC, USA), Lipogems (Lipogems International SPA, Italy), MyStem® (MyStem -LLC, USA) [255], which are already widely used in clinical applications. In the Lipogems device, for example, the lipoaspirate is microfragmented by using mild mechanical forces without the addition of collagenase or other enzymes/additives. Subsequent washings allow red blood cells and ADSCs to accumulate into the drain bag. The pellet obtained after centrifugation can be seeded to expand and characterize ADSCs [261]. To date, no system seems to achieve the ideal characteristics to meet the requirements established by the "International Federation for Adipose Therapeutics and Science" (IFATS) together with the ISCT. Therefore, one can only make "hypotheses" about the efficacy of one system compared to another or a set of different techniques that can complement each other. However, it should be considered that enzymatically isolated ADSCs are regarded as beyond the scope of “minimal manipulation” and therefore a drug. Thus, their production for clinical application requires compliance with “cell manufacturing” in accordance with the European Good Manufacturing Practice (eGMP) Guidelines. Instead, the clinical application of mechanically isolated ADSCs during the same operative session with minimal manipulation falls under the jurisdiction of the practice of medicine and is thus allowed.

Fig. 6.

Schematic representation of ADSCs isolation

Table 7.

Enzymatic digestion protocols used for ADSC isolation

Enzymatic solution	Conditions	References
0.075% (230 U/mg) Collagenase type I	30 min at 37  °C	[17, 19, 262, 263]
1 mg/mL Collagenase type I, 12.5–25-37.5 μg/L Trypsin 1 g/l Collagenase type I	30 min + 10 min at 37  °C 60 min at 37  °C under shecking/75RPM)	[249, 264, 265]
0.1% Collagenase type I	60 min at 37  °C	[266–268]
3 mg/mL Collagenase type I + 4 mg/mL dispase	60 min + 2–3 h at 37  °C	[269, 270]

ADSCs culture conditions

Under standard culture conditions, ADSCs are known to adhere to the plastic of culture flasks through the production of cytoplasmic protrusions conferring them typical fusiform morphology defined as fibroblastoid. This adhesion ability is also exploited in ADSC isolation processes to select these cells from cell populations deriving from the hematopoietic lineage that are unable to adhere to the plastic [250]. ADSCs are a relatively homogenous population that expands after 7-14 days of culture [271, 272], under controlled conditions at 37 °C and 5% CO₂. In the traditional culture of ADSCs, based on low-glucose media (DMEM or DMEM/F12), supplemented with animal sera or human-related substitutes, some authors have reported the need of additional growth factors to support a stable cell growth (Table 8) [272]. Lindroos et al. adopted serum-free and xeno-free media to culture ADSCs, and these media maintained the proliferative ability of the cells [273, 274]. The SVF resulting from enzymatic digestion is seeded on culture vessels and after 24 h the non-adherent cells and erythrocytes are removed. The resulting cell population mainly consists of mesenchymal origin cells, which are cultured in a non-inductive culture medium and at subconfluent level to prevent spontaneous differentiation [17, 275]. To obtain pure populations of ADSCs, it is also possible to add a filtration step using polyurethane and silkscreen/poly (lactide-co-glycolic acid) filter membranes in various pore sizes. Higuchi et al. described a membrane migration method in which SVF passed through a filter membrane, then membrane bearing attached ADSCs was cultured for 14–16 days to allow ADSCs to migrate out of the membrane [244]. Finally, the final products (adherent cells) are validated by flow cytometry to confirm the expression of specific markers.

Table 8.

Culture media for ADSCs propagation

Basal medium	Supplements	References
DMEM-LG	10% mesenchymal stem cell growth supplements (MSCGS); 10% FBS + 1% antibiotic/antimycotic	[19, 263, 268]
DMEM/F12	10% FBS + 1% antibiotic/antimycotic 3% FBS + 1% antibiotic/antimycotic + (0.1, 1.0 or 10 ng/mL) EGF and/or (0.1, 1.0 or 10 ng/mL) b-FGF	[276, 277]
Culture condition and cryoconservation	37  °C in a humid atmosphere with 5% CO2 10% DMSO in complete medium or FBS and/or Trehalose	(278, 279)

MSCs qualification

The International Society for Cellular Therapy (ISCT) has established criteria for the characterization of MSCs to ensure standardization and reproducibility in research and clinical applications [302]. Following the ISCT and IFATS, every MSCs isolated from any adult or fetal tissue is called to respond to a tripartite set of properties: (i) MSCs should adhere to plastic vessels when exposed to a culture medium supplemented with animal serum or human serum replacement. Indeed,MSCs can secrete and adapt to the extracellular environment, allowing them to adhere to newly secreted extracellular matrix (ECM) proteins; (ii) The presence of static cell markers on the plasma membrane has been recognized to validate the identity of MSCs, just as it has been for hematopoietic cells for decades. MSC should express CD73, CD90, and CD105 on their surface, and in several works has also been reported a constitutive presence of CD44; while they are negative for hematopoietic markers CD34, CD45, CD11b or CD14, CD 79 or CD 19. A low level of polymorphic Human Leukocyte Antigen (HLA) class 1 is present, while the class 2 and co-stimulatory molecules CD40 and CD80 are absent. Still debated is the constitutive presence of non-polymorphic HLA forms (e.g., HLA-G and HLA-E), both as membrane-bound or soluble mediators; (iii) MSCs can be reliably induced to mature into osteoblasts, adipocytes, and chondroblasts both in vitro and in vivo. In this regard, various assessments such as mixed lymphocyte reaction (MLR), flow-cytometric, and immunophenotypic profiling, etc. are performed to evaluate the characteristic of MSCs. These analyses aim to verify the purity of the cells and confirm their mesenchymal nature [264, 308, 309]. the identity qualification for MSCs has been set on cell adhesion to plastic even though some studies have shown the presence of a subpopulation of non-adherent MSCs equipped with the same multipotency properties and ability to migrate into damaged tissues as adherent cells [157, 280]. Another critical qualification test is based on FACS analyses, to certify the expression of specific surface markers (co-expression of CD73, CD90, and CD105) and the absence of hematopoietic markers (CD45, CD14, CD34, HLA-II) and endothelial proteins (CD31), the ability to differentiate into osteoblasts, chondrocytes, adipocytes since 2006 [12].

The cell surface markers of MSCs may vary among different sources [281]. For example, in addition to the other markers commonly used for MSCs, such as CD73, CD90, and CD105, there are other markers that are commonly used to identify BM-MSCs such as Stro-1, CD146, and CD271 that are expressed by BM-MSCs and have been defined as specific marker for BM-MSCs at early passages in culture. BM-MSCs are also positive for other surface markers such as SH2, SH3, CD10, CD29, CD44, CD49e, CD71, CD106, CD120a, CD124, CD133, CD166, HLA-ABC, MSCA -1 and SSEA-4, while they are negative for CD3, CD11b, CD14, CD19, CD31, CD34, CD45, CD79a. These markers help in further characterizing BM-MSCs. Moreover, the characteristics of BM-MSCs are closely linked to the ages and pathological conditions of the donors. [12, 143, 147, 185, 205, 282–286]. Mesenchymal stem cells sourced from the amniotic membrane (referred to as AMSCs) and amniotic fluid (known as AFSCs) display unique characteristics in terms of surface marker expression and their ability to suppress the immune response. AMSCs typically show the presence of MSC markers like CD44, CD73, CD90, CD105, and CD29, while lacking CD34 and CD45. Similarly, AFSCs also exhibit typical mesenchymal marker expression, including CD90, CD73, CD105, CD29, CD166, CD49e, CD58, and CD44, as confirmed through flow cytometry analysis [118]. Additionally, these cells expressed the HLA-ABC antigens, whereas the expression of the hematopoietic markers CD34 and CD45, the endothelial marker CD31 and the HLA-DR antigen was undetected. More importantly, the majority of cultured AFSCs expressed pluripotency markers such as, the octamer binding protein 3/4 (Oct-3/4), the homeobox transcription factor Nanog (Nanog) and the stage-specific embryonic antigen 4 (SSEA-4) [117, 287–289]. It was also reported that amniocyte cultures contain a small population of CD117 (a tyrosine kinase specific for stem cell factor present primarily in ESCs and primordial germ cells) positive cells, that can be clonally expanded in culture. The differentiation properties of CD117+ AFS were tested for the first time in vivo, proving in this way their stem cell identity. Experimental evidence suggested that AFSCs are derived from spindle shaped fibroblastoid cells [290]. To analyze the AFSCs subpopulations, two morphologically distinct populations of AFSCs of mesenchymal origin were identified, with different proliferation and differentiation properties, termed as spindle shaped (SS) and round shaped (RS). Both subpopulations were expressing mesenchymal stem cell markers at similar levels. However, it was identified that SS colonies expressed higher levels of CD90 and CD44 antigens compared to RS colonies [289]. Dental pulp stem cells (DPSCs) typically exhibit a blend of mesenchymal and embryonic stem cell markers. These include pluripotent stem cell markers like Oct4, Nanog, Sox2, SSEA, and c-Myc [291]. Besides the commonly found mesenchymal stem cell markers (CD73, CD90, CD105), research has shown the presence of additional markers in DPSCs such as STRO-1, CD10, CD13, CD106, CD117, CD29, CD44, CD146, CD166, and CD271 [292–294]. Moreover, other studies have also reported several cell surface antigens for prospective isolation of DPSCs, such as STRO-1, VCAM-1, SH2, SH3/SH4, CD271, GD2 and SSEA4 [282, 284, 295–297]. Apart from stem cell markers, DPSCs also express specific markers such as dentin sialophosphoprotein (DSPP), dentin matrix protein-1 (DMP-1), osterix (Osx), osteocalcin (OCN), osteopontin (OPN) alkaline phosphatase (ALP), and type I collagen [206, 298, 299]. Unlike other mesenchymal stem cells, ADSCs display varying levels of CD166 (ALCAM), CD271 (NGFR), CD248, CD146 (MCAM), CD13 (ANPEP), CD29 (ITGB1), CD36, CD44, CD200, and CD201 (PROCR). Some combinations of these markers are only detectable simultaneously through multichromatic flow cytometry. ADSCs also demonstrate high levels of CD10 (MME), CD26 (DPP4), CD49d (ITGA4), and CD49e (ITGA5). Negative markers associated with ADSCs include CD14/CD11b, CD19/CD79a, CD56 (NCAM1), CD3, and CD235a (GYPA). Attention should be given to the CD34 antigen, which was initially considered a negative marker for characterizing MSCs but has shown expression on ADSC surfaces during the early stages post-adipose tissue isolation. Its expression decreases with in vitro expansion. For precise identification and isolation of ADSCs, a combination of positive and negative markers is recommended [300–302] with a constant reduction following in vitro expansion [303, 304]. In conclusion, the combination of multiple positive and negative markers is recommended for increasing the specificity of ADSC identification and isolation. Regarding UCSCs, they do not display markers associated with endothelial and hematopoietic cells, such as CD31, CD45, CD34, CD117, CD14, CD11b, CD79a, CD19, and HLA-DR. Unlike some other types of MSCs, UCSCs do not exhibit CD133 expression. However, it is important to note that the characterization of these cells may vary depending on factors such as culture passage number, medium, and technique used. Furthermore, UCSCs have been found to express pluripotency markers including Octamer-binding transcription factor 4 (OCT-4), sex-determining region Y box 2 (SOX-2), Kruppel-like factor 4 (KLF4), and NANOG at both the gene and protein levels. This information is supported by studies conducted by various researchers such as [12, 34, 82, 90, 305, 306]. A third requirement relies on MSC maturation into mesoderm lineages, such as adipocytes, osteoblasts, and chondrocytes even thoughtheir commitment towards other mesoderm phenotypes as well as transdifferentiation into endoderm or ectoderm lineages has still to be proven. The researchers’ sparked enthusiasm for the generation of somatic cells starting from multipotent cells (as MSC) has historically not always been followed by solid proves and measurements. Mature and functional hepatocytes, cardiomyocytes, skeletal muscle, and neural precursors have gained quite a sparked enthusiasm in early 2000s, briefly later disproven and jeopardized by contradictory results. The existence of a subpopulation of BM-MSCs capable of differentiating not only in the mesenchymal lineage, but also in the ectodermal and endodermal germ layer lineages is still intensely debated [307, 308].. Actually, regulatory authorities, such as ISCT, also propose[309] the development of potency assays as release criteria for use them in advanced-phase clinical trials [309]. In particular, multifunctional assays that can characterize MSCs immunomodulatory abilities, through the combination of molecular genetics and secretome analysis, may serve as a platform for robust potency analysis . Unfortunately, a common mechanisms of action of MSCs have not yet been fully clarified. This has led to the frequent use of potency assays based on surrogate markers, an array of multiple complementary assays, or a combination of these [309, 310]. Despite the many unknowns, the T-cell inhibition assay remains the most popular method for assessing the potency of MSCs . However, since potency tests must be evaluated on a case-by-case basis depending on the source tissue, it is not possible to rule out any of the tests a priori. Tests based on the properties of the mesenchymal stromal cells (MSCs) or responding cells may also be appropriate, provided the relevance of the proposed test and readout parameters is adequately justified and corroborated by sufficient clinical and non-clinical data [313].

Beyond the identification criteria, the qualification of MSCs for clinical use requires a series of rigorous safety tests to ensure compliance with clinical-grade standards. The sterility of the final cell product is of critical importance. MSCs must therefore undergo thorough, multi-stage testing to rule out microbial contamination (bacteria and fungi), mycoplasma, and to quantify endotoxin levels, thereby ensuring patient safety [314]. Concurrently, the assessment of cell viability, typically measured by methods such as trypan blue exclusion or flow cytometry, is essential to guarantee that an adequate number of healthy and functional cells are administered [315]. Another critical aspect, especially for cellular products requiring significant in vitro expansion, is the evaluation of genetic stability. Long-term cultivation can increase the risk of chromosomal abnormalities and genetic instability, raising concerns about potential malignant transformation [315]. Therefore, genetic stability testing, including karyotype analysis to detect numerical and structural chromosomal aberrations, and more sensitive analyses like Copy Number Variation (CNV) assessment, are considered crucial to rule out cellular transformation and ensure the long-term safety of the product [316]. This comprehensive set of qualifications and safety tests is indispensable for translating MSC-based therapies from research to clinical practice [302, 309].

Strengths and weaknesses of methods and protocols to isolate and characterize MSC NICHE

A better understanding and characterization of the mesenchymal stem cell niche are of paramount importance for elucidating their biological behaviors and optimizing their therapeutic potential [3]. The methods and protocols now used for isolating and characterizing MSCs present both advantages and inherent limitations, particularly in terms of the spatial and temporal heterogeneity of the niche itself, which is difficult to reproduce in experimental systems, especially in vivo. [12, 317]. For example, DPSCs are found in the dental pulp, a tissue that is characterized by rich vascularization and innervation, and which is comprised of different and heterogeneous areas [171]. The main isolation procedures, including tissue explant culture (OG), enzymatic digestion (ED), and mechanical extraction [178–180], are designed to extract cells from their context, inevitably leading to the disruption of the native three-dimensional architecture, the loss of spatial cell-cell and cell-extracellular matrix (ECM) interactions, and the alteration of the vascular and neural microenvironment (ref). The bone marrow itself constitutes a complex microenvironment where BM-MSCs, representing approximately 0.01% of mononuclear cells, interact with hematopoietic stem cells (HSCs) and other stromal elements, providing structural and functional support [7, 127, 139]. Nevertheless, standard methodologies for BM-MSC isolation, such as bone marrow aspiration followed by density gradient centrifugation (e.g., Ficoll) and subsequent selection based on plastic adherence or flow cytometry [142, 144, 145], present significant weaknesses for niche studies.

Similarly, the adipose-derived stem cell (ADSC) niche within adipose tissue (AT) is a dynamic microenvironment that significantly influences the properties and regenerative potential of ADSCs. ADSCs are the precursors of adipocytes and primarily reside in a perivascular niche [238, 269]. However, the conventional method of isolating them—starting with a biopsy or lipoaspirate and involving digestion or mechanical processing to obtain the stromal vascular fraction (SVF)—destroys the three-dimensional architecture of TA [254–256]. The placenta, amnio, umbilical cord (UC), and umbilical cord blood (UCB) are sources and niches of stem cells. Compared with other sources, they are readily available, thus avoiding the need for invasive procedures or raising ethical issues. They also offer an advantage over adult tissues in that their harvesting avoids potential age-related problems. In addition, they demonstrate superior proliferation capacity, lifespan, and differentiation potential in comparison to BM-MSCs and AT-MSCs [318, 319].

Regarding the characterization of the niche, in situ techniques, such as immunohistochemistry (IHC) and immunofluorescence (IF) represent a strength, allowing the visualization of specific stem cells about their niche components, such as the vasculature, which is crucial to the concept of the perivascular niche [21, 146].

However, it is complicated by tissue heterogeneity and variations induced by the physiological or pathological state of the donor [242, 243]. Furthermore, many of the markers used to identify mesenchymal stromal cells (MSCs) have been defined on ex vivo cultured cells, which provides limited information on cellular behavior within the complexity of the native niche. Their expression can also vary significantly from the in vivo state, as discussed by the International Federation for Adipose Therapeutics and Science (IFATS) and the International Society for Cellular Therapy (ISCT) [299, 302]. Finally, the isolation and characterization of all cell subpopulations within the niche is a significant challenge, often complicated by a lack of specific markers and the cells' sensitivity to manipulation. Furthermore, the translation of results obtained from animal models to humans is frequently limited by species-specific differences in niche composition and regulation. Another critical issue is dependence on methodologies that can alter the physiological state of cells during isolation, requiring continuous optimization of protocols to preserve niche integrity and function [43, 320]. Therefore, a multidisciplinary approach combining various techniques while recognizing their limitations is essential for a comprehensive understanding of the MSC niche.

Strengths and weaknesses of methods and protocols of isolation of MSCs using microfluidic platform

Microfluidic systems provide a very small-scale environment for stem cells which is similar to the body one. As already noted by A. Aghlmandi, the main advantage of these systems is their small size, low sample, and reagent consumption [321]. Microfluidic platforms are also capable of performing several test steps with increased efficiency and high speed within short time intervals. The use of microfluidic platforms for isolating mesenchymal stem cells (MSCs) has gained significant attention in recent years, with several studies highlighting both its strengths and limitations [322]. One of the main strengths of microfluidic platforms is their ability to isolate MSC subpopulations with high precision and purity. Techniques like size-based sorting, as demonstrated by Tan et al. and Yang et al., allow for the enrichment of specific MSC populations that have superior regenerative potential, such as those with enhanced chondrogenic or osteogenic properties [323, 324]. This precision is specifically important for therapeutic applications, where the functionality of the isolated cells is just as crucial as their quantity. Additionally, label-free sorting, as described by Yin et al., ensures that the cells remain unaltered by the introduction of fluorescent markers, preserving their natural state and avoiding potential toxicity [321]. The scalability of these microfluidic devices also stands out, with Tan et al. showing that techniques like deterministic lateral displacement (DLD) can efficiently process larger volumes of bone marrow aspirates, making the method more feasible for clinical applications [323]. However, despite these advantages, there are several challenges to overcome. One key limitation is the technical complexity and high cost of microfluidic platforms, which can make them inaccessible for broader clinical use. The fabrication of microfluidic devices and the expertise required for their operation can add significant overhead to the process, limiting their accessibility in resource-limited settings. Another concern is the throughput of these systems. While scalable, some microfluidic platforms may still struggle to process large sample volumes quickly enough for high-demand applications [323]. Moreover, the potential for cell loss or damage during the sorting process remains a challenge. As Yin et al. highlight, the shear forces or long residence times in the device can compromise cell viability, which could undermine the therapeutic potential of isolated MSCs [325]. Finally, the reliance on biophysical properties like cell size for sorting can miss certain MSC subpopulations that are defined by other factors, such as surface markers or functional properties, which could limit the versatility of microfluidic devices. In conclusion, microfluidic platforms for MSC isolation offer significant advantages in terms of precision, purity, and scalability, which could greatly enhance their therapeutic potential. However, the technical barriers, cost, throughput limitations, and potential for cell damage are factors that still need to be faced and overcome before these platforms can be widely adopted in clinical practice.

Conclusion

To conclude, the research and application of mesenchymal stem cells (MSCs) continue to be a field of great interest in regenerative medicine. The emergence of innovative approaches such as on microfluidic platform or based on properties of specific stem niche, undoubtedly promises significant advantages in terms of cell purity and process automation. However, it is crucial to recognize that these methodologies are still in a development and validation phase. Their current limitations often include technical complexity, high cost and scalability that does not always can be adopted for a large-scale clinical application. In contrast, standardized isolation and characterization methods, adapted to MSCs from different sources, make them the preferred choice for most laboratories. In addition, their ability to handle larger volumes of tissue and to be easily replicated in different operational settings contributes significantly to their prevalence.. By following established protocols, researchers can increase the reliability and reproducibility of their findings in the field of regenerative medicine and cell therapy. .

Abbreviations

MSCs

Mesenchymal stromal/stem cells

UCSCs

Umbilical cord stem cells

WJ-MSCs

Wharton Jelly derived MSCs

UCB-MSCs

Umbilical cord blood-mesenchymal stromal cells

AFSCs

Amniotic fluid stem cells

AMSCs

Amniotic membrane stem cells

DPSCs

Dental pulp stem cells

BM-MSCs

Bone marrow-mesenchymal stromal cells

SHEDs

Human exfoliated deciduous teeth stem cells

PDLSCs

Periodontal ligament stem cells

DFSCs

Dental follicle stem cells

SCAPs

Apical papilla stem cells

GMSCs

Gingival mesenchymal stem cells

DSCs

Dental stem cells

ESCs

Embryonic stem cells

ADSCs

Adipose-derived stem cells

Authors contributions

Conceptualization, V.M. and S.D.M.; writing—original draft preparation, F. S., F.P., J.F., L.L., R.G., M.G.R., S. B., M.V., S. B., U. G., A. J. S., A. A., C. M., S.D. M.; writing—review and editing, C.S., F. M., S. C., E.P., R. G., V.M. and S.D.M.; supervision, V.M. and S.D.M. All authors have read and agreed to the published version of the manuscript.

Funding

Italian Ministry of Health, project code: GR-2021-12374686.

Data avalaibility

Not applicable.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.