Stem Cells International
Volume 2021 (2021), Article ID 6632052, 22 pages
https://doi.org/10.1155/2021/6632052

Review Article

Stromal Vascular Fraction and Amniotic Epithelial Cells: Preclinical and Clinical Relevance in Musculoskeletal Regenerative Medicine

Francesca Veronesi,¹ Melania Maglio,¹ Deyanira Contartese,¹ Lucia Martini,¹ Aurelio Muttini,² and Milena Fini¹

¹IRCCS Istituto Ortopedico Rizzoli, Complex Structure of Surgical Sciences and Technologies, Via di Barbiano 1/10, 40136 Bologna, Italy
²Faculty of Bioscience and Agro-Food and Environmental Technology, University of Teramo, Italy

Correspondence should be addressed to Melania Maglio; melania.maglio@ior.it

Received 5 November 2020; Revised 31 December 2020; Accepted 6 February 2021; Published 22 February 2021

Academic Editor: Elena Jones

Copyright © 2021 Francesca Veronesi et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract.

Musculoskeletal regenerative medicine is mainly based on the use of cell therapy to heal damaged tissues such as bone, cartilage, and tendons. Throughout the years, different cell types have been employed for the treatment of musculoskeletal diseases, in particular, mesenchymal stem cells (MSCs) derived from bone marrow (BMSCs) and adipose tissue (ADSCs). Though the results of these literature studies have been encouraging, there are some limitations, especially on long-term results. Recently, some interest has shifted towards new cell types such as the stromal vascular fraction (SVF) and amniotic endothelial cells (AECs). The aim of the present literature review is to evaluate preclinical and clinical studies that used SVF and AECs for musculoskeletal tissue regeneration. Forty-eight preclinical and clinical studies, performed in the last 10 years, were identified. Both SVF and AECs, injected or implanted with or without scaffolds, were shown to be valid alternatives, and in some ways superior, to ADSCs and BMSCs, being able to differentiate towards osteogenic, chondrogenic, and tenogenic lineages, and to promote cell and tissue regenerative potential. The use of SVF and AECs could represent a new regenerative treatment in several musculoskeletal pathologies, solving the problem of cell expansion in vitro.

1. Introduction

In several musculoskeletal tissue diseases that affect the cartilage, tendons, and bone, there is the need for new regenerative treatments instead of traditional conservative or surgical therapeutic approaches that mainly give palliative care or short-term curative effects [1–3]. In this regard, mesenchymal stem cells (MSCs) have been employed as substitutes and as a promising therapeutic strategy to restore tissue biology, having success in several pathologies, even if the ideal source of stem cells is still debated. In the ambit of cellular therapies, MSCs play a leading role because they possess paracrine activity, through which they exert anti-inflammatory, antiapoptotic, antifibrotic, proangiogenic, and mitogenic activities on the microenvironment, adjacent tissues, and cells [4]. Different tissues have been identified as a source of MSCs, and among them, MSCs from bone marrow (BMSCs) have been mostly used for the regeneration of musculoskeletal tissues, achieving satisfactory results [5]. However, MSCs from adipose tissue (ADSCs) show advantages over BMSCs: ADSCs are reported to have higher genetic stability and higher proliferation, differentiation, and immunoregulatory abilities, and they also show lower senescence than BMSCs [6, 7]. The clinical use of MSCs could be complicated due to donor site morbidity, ageing or disease of the donor, and the necessity of a previous in vitro expansion to obtain a large cell number able to produce a clinical effect (it is estimated that nearly cells should produce a clinically appreciable effect) with associated risks of cell transformation or infection, replaced by a one-step procedure with bone marrow [8].

With particular reference to therapies aimed at skeletal muscle regeneration, other cellular sources have also been tested, including mesoangioblasts, derived from blood vessels, or fibro/adipogenic progenitors (FAPs), multipotent mesenchymal cells derived from skeletal muscle. Mesoangioblasts are multipotent mesodermal progenitor cells that can be isolated by fetal muscle biopsy [9]. FAPs are involved in a dynamic crosstalk with the other cellular populations of the muscle stem cell niche, in particular, immediately after injury occurrence [10]. However, isolation of FAPs requires the application of a long and complex protocol, including muscle dissection or digestion, and subsequent characterization with antibody staining and cell sorting [11].

In an effort to find a smarter cell substitute for MSCs, the stromal vascular fraction (SVF) has been characterized to be employed in preclinical and clinical scenarios [12]. SVF includes not only ADSCs but also a heterogeneous group of cells, such as progenitor cells, endothelial cells, fibroblasts, monocytes, macrophages, immune cells, muscle cells, pericytes, CD34+ cells, growth factors (GFs), a few adipocytes, and stromal components [13]. Similar to MSCs, SVF is proangiogenic and immunomodulatory, and its cellular components are able to differentiate and proliferate, all of the features that make it suitable for tissue regeneration [14]. The advantage of using SVF with respect to expanded ADSCs is immediately clear since SVF, obtained with collagenase digestion and centrifugation of adipose tissue, can be easily harvested from a patient through lipoaspiration. Moreover, it is autologous, requires minimal manipulation, and contains ADSCs at a percentage ranging from 0.06 to 4 CFU-f. Therefore, SVF could be injected directly into a damaged tissue reducing inflammation and promoting regeneration, with consequent reduction in health costs and hours of hospitalization [15, 16]. Indeed, SVF allows the so-called one-step surgical procedure, through which it is possible to harvest and implant SVF in the same surgical session, not requiring in vitro expansion. This procedure consists of minimal cell manipulation and low risks linked to culture, without specific regulatory requirements for clinical translation, thus accelerating surgery. The process that goes from surgical adipose tissue harvest, SVF production, and its seeding onto a scaffold or onto hydrogels or its direct injection, lasts at most 4 hours [17, 18].

Another innovative cell source was found in the human placenta, a waste material with cells characterized by high plasticity [19]. The amniotic membrane is obtained from the placenta without an invasive procedure and could be employed as an autologous or allogenic graft due to its immunomodulation properties [20, 21]. The amniotic endothelial cells (AECs) are considered a valid alternative to MSCs because they differentiate into three lineages (osteogenic, adipogenic, and chondrogenic), express mesenchymal and embryonic stem cell markers, show a nontumorigenic phenotype, and have a high yield in terms of in vitro expansion. In addition, AECs replace embryonic stem cells that show a clear impact on ethical matters [22].

Currently, while BMSCs are widely analysed in musculoskeletal pathologies, little is known on the use of SVF and AECs as cell therapies for the regeneration of musculoskeletal diseases especially in comparison with other common cell types and sources [23–26]. These cells were mostly characterized in vitro and sporadically compared with other cell types, despite exhibiting noninferior characteristics [27]. However, considering the advantages of the fact that the source of these cells is waste material, which does not include any kind of sampling, it would be interesting to compare them with SVF, whose use in regenerative medicine is very promising, though require a more demanding harvesting. The aim of this review is to collect preclinical and clinical studies, performed in the last 10 years, which used SVF or AECs in bone, cartilage, and tendon tissue regeneration.

2. Materials and Methods

The review has been performed according to the Preferred Reporting Items for Systematic Reviews and Meta-Analyses (PRISMA) statement. The studies included in the present review were identified through the http://www.pubmed.com/ and http://www.webofknowledge.com/ databases. In the first database, the keywords used were the following: “(amniotic epithelial stem cells OR stromal vascular fraction) AND (tendon OR cartilage OR bone OR ligament).” The limits were use of English language and publication date from 2010/01/01 to 2019/06/31. In the second database, the keywords were the same, but the limits were use of English language, publication date from 2010 to 2019, and document type was article.

A total of 284 articles were found using the http://www.pubmed.com/ database, and from among them, 242 articles were excluded because they were reviews; or they were focused on cell isolation techniques; or they were not related to musculoskeletal tissues but were related to the heart, corneal epithelium, liver, skin, or bladder; or they were concerned with culture-expanded ADSCs, BMSCs, embryonic stem cells, or MSCs derived from amniotic fluid. Therefore, forty-two studies were accepted.

A total of 452 articles were found using http://www.webofknowledge.com/, and of these, 449 were excluded because they were not inherent, or they were reviews, or they overlapped with the previous search (38 studies). Therefore, three studies were accepted.

In addition, after reading the reference lists of the accepted studies, another six articles were included. Therefore, a final total of 51 studies were taken into consideration (Figure 1).

Figure 1: Schematic representation of the search strategy.

3. Results

Most of the studies (35/51 studies) dealt with SVF [7, 14, 25–57], and 16/51 studies dealt with AECs [58–73] (Figure 2).

(a)

(b)

(c)

Figure 2: Amount of in vitro, in vivo, and clinical studies. (a) Pie chart of the percentages of the literature studies on SVF (blue: 73%) and AECs (red: 27%). In addition, the other two pie charts for each type of cell present the percentages of literature studies on bone (blue), cartilage (red), and tendon (green) regeneration. (b) Bar graph on the number of clinical, in vivo, and in vitro studies found in the literature, for the regeneration of bone (blue), cartilage (red), and tendon (green) by AECs. (c) Bar graph on the number of clinical, in vivo, and in vitro studies found in the literature, for the regeneration of bone (blue), cartilage (red), and tendon (green) by SVF.

The 35 studies on SVF were conducted to treat bone (no of studies: 16) [17, 28–42], cartilage (no of studies: 13) [7, 43–54], and tendon (no of studies: 6) [55–60] defects (Figure 2). They were conducted in vitro [17, 28–30, 43] and in vivo [7, 29, 31–39, 42–45, 55–58], and some of them were clinical studies [40–43, 46–54, 59, 60].

Regarding AECs, seven studies were performed on bone [61–67], 2 on cartilage [68, 69], and 7 on tendon [70–76] defects. Among these 13 studies, some were carried out in vitro [61–64, 68–70] and others in vivo [65–67, 71–76] (Figure 2).

In addition, 11 studies compared two or more cell types in the same study [7, 17, 28, 29, 35, 43, 44, 56, 61, 67, 68]. More precisely, SVF were compared with the following: (1) BMSCs [28, 56], in vitro [28] and in vivo in rabbit tendon regeneration [56]; (2) ADSCs [7, 17, 35, 44], in vitro [17] and in vivo in mouse bone defect [35], goat osteochondral defects [44], and sheep knee osteoarthritis (OA) [7]; and (3) monocyte cell line (THP1) and ADSCs [29] or chondrocytes and ADSCs [43] in rat bone defects [29] or mouse subcutaneous pouches [43].

AECs were directly compared with the following: (1) ADSCs in vitro [61]; (2) BMSCs and amniotic fluid MSCs (AFMSCs) in mouse subcutaneous pouches [67]; and (3) chondrocytes, BMSCs, and amniotic MSCs (AMSCs) in vitro [68].

3.1. SVF and Bone

3.1.1. In Vitro Studies

Four in vitro studies were performed with human SVF (hSVF) obtained from subcutaneous tissue [29] or nonspecified sites [17, 28, 30] of donors (Table 1).

Table 1: In vitro, in vivo, and clinical studies on SVF in bone regeneration.

Materials
Treatment groups
Evaluations
Results
Ref

In vitro:
hSVF from 4 donors (42-62 yrs) ()
Purchased hBMSCs ()
Group 1: BMSCs
Group 2: BMSCs+SVF
ALP activity
RT-PCR (RUNX2, COLL I, ALP, and OSX)
12, 24, and 48 hrs
Group 2: ↑ gene expression than group 1
24 and 48 hrs
Group 2: ↑ ALP activity than group 1
[28]

In vitro:
hSVF () or hADSCs () from subcutis of 3 donors (38-52 yrs)
THP1 ()
In vivo:
46 nude rats with bilateral bone defects () in femoral condyles
hSVF () or hADSCs () from abdomen
THP1 ()
In vitro:
Group 1: THP1
Group 2: SVF
Group 3: THP1+SVF
Group 4: ADSCs
Group 5: THP1+ADSCs
In vivo:
Group 1: no treatment
Group 2: THP1
Group 3: SVF
Group 4: THP1+SVF
Group 5: ADSCs
Group 6): THP1+ADSCs
In vitro:
ALP activity
Calcium content
In vivo:
Histology
Histomorphometry
In vitro:
2 wks
Group 3: ↑ ALP activity than groups 2, 4, and 5.
1 mo
Group 3: ↑ calcium content than groups 2 and 5
In vivo:
4 and 10 wks
Groups 3, 4, 5, and 6: ↑ BA/TA than groups 1 and 2.
10 wks
Groups 3 and 4: ↑ BA/TA than groups 5 and 6
[29]

In vitro:
hSVF () from 8 donors ()
PLCL scaffold
COLL I/COLL III scaffold
Group 1: PLCL+SVF in NM
Group 2: COLL I/COLL III+SVF in NM
Group 3: PLCL+SVF in CM
Group 4: PLCL+SVF in OM
Group 5: COLL I/COLL III+SVF in CM
Group 6: COLL I/COLL III+SVF in OM
RT-PCR (ACAN, SOX6, SOX9, ALP, ONC, COLL I, COLL III, and COLL X)
4 days
Group 3: ↑ ACAN gene expression than group 5)
2 wks
Groups 3 and 5: ↑ SOX6, SOX9 gene expression during time
Groups 4 and 6: ↑ ALP, ONC, and COLL I gene expression during time
3 wks
Group 5: ↑ COLL II gene expression than group 3
Groups 3 and 5: ↓ COLL X gene expression during time
[30]

In vitro:
hSVF or hADSCs () from 7 donors
Xenohybrid bone graft scaffold ()
Group 1: SVF in plastic in NM
Group 2: ADSCs in plastic in NM
Group 3: scaffold+SVF in NM
Group 4: scaffold+ADSC in NM
Group 5: SVF in plastic in OM
Group 6: ADSCs in plastic in OM
Group 7: scaffold+SVF in OM
Group 8: scaffold+ADSC in OM
Micro-CT
IHC (OCN)
ALP activity
Mineralization
2 mo
Group 7: ↑ bone trabeculae than groups 4, 8, and 3
Groups 3 and 4: ↓ OCN than groups 7 and 8
Group 1: ↑ ALP activity than group 2
Groups 1 and 2: ↓ mineralization than groups 5 and 6
[17]

In vivo:
28 nude rats with subcutaneous pouch
hSVF () from 5 donors ()
HA scaffold (1 cm diameter, 1 cm height)
Group 1: scaffold
Group 2: scaffold+SVF previously cultured for 5 days
Micro-CT
Histomorphometry
1 wk
Group 2: ↑ vessel number, M2 macrophages than group 1
2 mo
Group 2: ↑ BV/TV than group 1
[31]

In vivo:
CD1nu/nu athymic mice with subcutaneous pouches
hSVF from abdomen or breast of donors ()
βTCP scaffold (8 mm diameter, 4 mm height)
HA scaffold (8 mm diameter, 4 mm height)
Group 1: βTCP+SVF ( CFU-f/cm³)
Group 2: βTCP+SVF ( CFU-f/cm³)
Group 3: βTCP+SVF ( CFU-f/cm³)
Group 4: HA+SVF ( CFU-f/cm³)
Group 5: HA+SVF ( CFU-f/cm³)
Group 6: HA+SVF ( CFU-f/cm³)
Histology
2 mo
Groups 1, 4, 3, and 6: ↑ dense matrix than groups 2 and 5
Groups 3 and 6: ↑ dense matrix similar to osteoid than groups 1 and 4
[32]

In vivo:
Nude mice with subcutaneous pouches
Nude rats with bilateral defects in parietal bone (4 mm)
Devitalized hypertrophic cartilage pellet from 5 donors ()
hSVF from 12 donors ()
Group 1: pellet in pouches
Group 2: pellet+SVF (/ml) in pouches
Group 3: pellet+SVF (/ml) in pouches
Group 4: pellet+SVF (/ml) in pouches
Group 5: pellet in bone defects
Group 6: pellet+SVF (/ml) in bone defects
Micro-CT
Histology
3 mo
Group 3: ↑ mineralized volume, BA than group 1
Groups 2 and 3: ↑ vessel length density than group 1
1 mo
Group 6: ↑ BA than group 5
[33]

In vivo:
24 syngenic Balb/c mice (8 wks) with subcutaneous pouches
Autologous SVF () from epididymis
Bio-Oss® scaffold (deproteinized sterilized bovine bone) (10 mg)
Group 1: scaffold
Group 2: scaffold+SVF+PRP
Histology
IHC (OPN)
Histomorphometry
RT-PCR (OSX, OCN, COLL I, and ALP)
2 wks
Group 2: ↑ OPN protein than group 1
2 and 4 wks
Group 2: ↑ OSX, OCN gene expression than group 1
1, 2, and 4 wks
Group 2: ↑ COLL I gene expression than group 1
2 mo
Group 2: ↑ ALP gene expression, osteoid-like tissue, OPN protein, vascularization than group 1
[34]

In vivo:
14 nude immunocompromised mice (8 wks) with bilateral critically sized calvarial defects
Nude mice with muscular pouches
hSVF or hADSCs () from flank, scapula, abdomen, or inner thigh of donors
PCL-DCB scaffold (4 mm diameter, 0.64 mm length)
Group 1: scaffold in bone defect
Group 2: scaffold+SVF in bone defect
Group 3: scaffold+ADSCs in bone defect
Group 4: scaffold in pouches
Group 5: scaffold+SVF in pouches
Group 6: scaffold+ADSCs in pouches
Micro-CT
Histology
3 mo
Groups 2 and 3: ↑ BV than group 1
Group 2: ↑ BV during time
10 days
Group 5: ↑ VA than groups 4 and 6
6 wks
Groups 5 and 6: ↑ VA than group 4
[35]

In vivo:
50 SD rats (8 wks) with critically sized calvarial defects
Autologous SVF () from inguinal region
DBM scaffold
PLA scaffold
Group 1: no treatment
Group 2: DBM
Group 3: DBM+SVF
Group 4: DBM+PLA
Group 5: DBM+PLA+SVF
Gross evaluations
Histology
IHC (OCN)
2 mo
Groups 3 and 5: ↑ defect filling, BA, OCN than groups 1, 2, and 4
Group 2: ↑ defect filling than groups 1 and 4
Group 4: ↑ defect filling than group 1
[36]

In vivo:
15 NZW rabbits (14-16 wks) with unilateral segmental bone defect in the middiaphysis of left ulna (20 mm)
Allogenic SVF () from suprascapular region
PLGA scaffold ()
Group 1: scaffold
Group 2: scaffold+SVF
Group 3: scaffold+osteogenically differentiated SVF
Radiography
Micro-CT
Histology
2 mo
Group 3: defect bridging, trabecular bone with smooth margins
Group 3: ↑ BVF than groups 1 and 2
Group 3: ↑ Tb.Th than group 1
Groups 2 and 3: scaffold absorbed
Group 2: ↑ BVF than group 1
[37]

In vivo:
28 C57BL/6J mice (8-12 wks old) with bilateral ONJ-like lesions
Allogenic SVF () from inguinal region
Group 1: saline
Group 2: SVF
Micro-CT
Histology
Histomorphometry
2 and 4 wks
Group 2: ↑ BA, osteocytes, bone filling, Tb.Th, BMD, collagen fibers, blood vessels, VA, F4/80⁺ macrophages; ↓ necrotic BA, empty lacunae, Tb.Sp, PMN infiltration than group 1
[38]

In vivo:
1 thoroughbred gelding in training (5 yrs) with lameness of the right carpal bone
Autologous SVF () from region above dorsal gluteal muscles
Group 1: SVF
Clinical evaluation
Lameness evaluation
4 mo
Group 1: return to racing
20 mo
Group 1: no injuries, no lameness, high performance level
[39]

Clinical trial:
10 pz (46-69 yrs old) partially edentulous requiring bilateral or unilateral dental implants
Autologous SVF (10⁷ nucleated cells)
βTCP scaffold
BCP (HA/βTCP 60%/40%) scaffold
Group 1: βTCP
Group 2: βTCP+SVF
Group 3: BCP
Group 4: BCP+SVF
Histology
Histomorphometry
6 mo
All groups: =bone formation, blood vessels
[40]

Clinical trial:
10 pz (46-69 yrs) edentulous in the posterior maxilla
Autologous SVF from abdomen
βTCP scaffold
BCP (HA/βTCP 60%/40%) scaffold
Group 1: βTCP
Group 2: βTCP+SVF ()
Group 3: BCP
Group 4: BCP + SVF ()
Micro-CT
Histomorphometry
6 mo
Groups 2 and 4: ↑ BV/TV, OV/TV than groups 1 and 3
≥36 mo
No adverse effects
[41]

Clinical trial:
8 pz (62-84 yrs) with displaced low-energy fractures of the proximal humerus
Autologous SVF from abdomen
Fibrin hydrogel+porous silicate HA microgranules scaffold
In vivo:
Adult nude rats with a critically sized segmental defect in femora (6 mm)
Fibrin hydrogel+porous silicate HA microgranules scaffold
hSVF () from abdomen
Clinical trial:
Group 1: scaffold+SVF pelleted
In vivo:
Group 1: scaffold
Group 2: scaffold+SVF
Clinical trial:
Safety
Histology
Micro-CT
In vivo:
Micro-CT
Histology
Clinical trial:
12 mo
Group 1: no adverse reactions; ↓ pain during time; ↑ formed bone ossicles during time
In vivo:
2 mo
Group 2: ↑ mineralized volume, BV/TV, newly formed bone, frank bone, early stage of maturation than group 1
[42]

In the first study, ALP activity and gene expression of Runt-related transcription factor 2 (RUNX2), collagen I (COLL I), alkaline phosphatase (ALP), and Osterix (OSX) of human BMSCs (hBMSCs) significantly increased when cocultured with hSVF in comparison to hBMSCs cultured alone [28].

In another study, ALP activity and calcium content of THP1 increased more when cocultured with hSVF than with hADSCs, after two and four weeks of culture. Additionally, in this study, hSVF or hADSCs alone or combined with THP1 cells were implanted in bone defects in femoral condyles of 46 nude rats. Ten weeks after implanting, it was observed that hSVF increased significantly more bone area (BA) than hADSCs [29].

Two types of scaffolds, namely, poly(L-lactide-co-caprolactone) (PLCL) and COLL I/COLL III, were seeded with hSVF and cultured in normal (NM), chondrogenic (CM), or osteogenic (OM) media to observe hSVF differentiation. The expression of sex-determining region Y box 6 (SOX6) and SOX9 significantly increased when the PLCL construct was cultured in CM. Gene expression of ALP, osteonectin, and COLL I increased when it was cultured in OM. Aggrecan gene expression increased when hSVF was cultured on PLCL, while COLL II gene expression increased with the COLL I/COLL III scaffold [30].

Finally, in another study, hSVF was cultured on a xenohybrid bone graft and compared with hADSCs. hSVF induced the formation of more bone trabeculae than hADSCs after 2 months of culture [14].

3.1.2. In Vivo Studies

Four in vivo studies were conducted in subcutaneous or muscular pouches of nude rats [31], athymic mice [32], nude mice [33], and syngenic mice [34]. SVF was obtained from the abdomen or breast [32], epididymis [34], or nonspecified sites [31, 33] of human donors (Table 1).

In one study, a hydroxyapatite (HA) scaffold was implanted alone or combined with hSVF. The addition of hSVF increased vessel number and M2 macrophages after 1 week and bone volume (BV) after 2 months [31].

In another study, beta-tricalcium phosphate (βTCP) or HA scaffolds were seeded with hSVF at 3 different concentrations ( CFU-f/cm³,  CFU-f/cm³, or  CFU-f/cm³). After 2 months from implantation, both scaffolds, seeded with hSVF at concentrations of  CFU-f/cm³ or  CFU-f/cm³, increased the formation of dense matrix [32]. An increase in mineralized volume, BA, osteoid matrix formation, and vessel length was observed 3 months after the implantation of a devitalized hypertrophic cartilage pellet with hSVF at a concentration of /ml or /ml in comparison to a concentration of /ml. Moreover, BA was significantly higher in nude rats with bilateral parietal bone defects treated with a scaffold and hSVF than with a scaffold alone after 1 month [33].

Najman et al. implanted a deproteinized sterilized bovine bone scaffold alone or combined with autologous SVF and platelet-rich plasma (PRP). Evaluations were performed after 1, 2, 4, and 8 weeks when it was observed that SVF and PRP significantly increased osteopontin (OPN) protein levels; gene expression of OSX, COLL I, ALP, and osteocalcin (OCN); osteoid-like tissue formation; and vascularization compared to a scaffold alone [34].

The other 5 in vivo studies were performed in critically sized calvaria defects in nude mice [35] or in rats [35], in segmental bone defects in the middiaphysis of the left ulna of rabbits [37], in osteonecrosis jaw- (ONJ-) like defects of mice [38], and in the right carpal bone of 1 horse [37]. SVF was obtained from the flank, scapula, abdomen, inner thigh [35], inguinal region [36], and gluteal muscle [39], and used as autologous [36, 39], allogenic [37, 38], or xenogenic grafts [32] (Table 1).

Both hSVF and hADSCs, seeded onto a polycaprolactone-decellularized bovine bone extracellular matrix (PCL-DCB) scaffold, increased BV more than the scaffold alone; in addition, hSVF increased BV during time, up to 3 months, in critically sized defects. In the same study, the PCL-DCB scaffold, seeded with hSVF and hADSCs, was also implanted in muscular pouches of nude mice, showing a high vascular volume in the presence of hSVF up to 6 weeks later [35]. Demineralized bone matrix (DBM) or DBM combined with polylactic acid (PLA), seeded or not with SVF, were always used in critically sized defects, and after 2 months, it was observed that when used alone, both scaffolds increased defect filling, but when combined with SVF, they also increased BA and OCN protein levels [36].

The poly(lactide-co-glycolide) (PLGA) scaffold was seeded with SVF, osteogenically induced or not, and implanted into rabbit ulna defects. Two months after implanting, SVF that was not osteogenically induced increased bone volume fraction (BVF), while SVF that was osteogenically induced also increased defect bridging, trabecular bone, and trabecular thickness (Tb.Th) [37].

In ONJ-like lesions, after 2 and 4 weeks, SVF increased living BA, osteocytes, bone filling, Tb.Th, bone mineral density (BMD), collagen fibers, blood vessels, and F4/80+ macrophages, while it reduced necrotic BA, empty lacunae, trabecular separation (Tb.Sp), and polymorphonuclear (PMN) infiltration in comparison with saline solution [38].

Finally, SVF injection in the carpal bone allowed a horse to return to racing after 4 months with no injuries and lameness and even with a higher performance level after 20 months [39].

3.1.3. Clinical Studies

Three clinical studies were performed in 10 edentulous patients [40, 41] and in 8 patients with displaced low-energy fractures of the proximal humerus [42]. SVF was autologous and was obtained from the abdomen [41, 42] or nonspecified sites [40] (Table 1).

In two studies, biphasic calcium phosphate (BCP) or βTCP scaffolds, with or without SVF, were implanted in edentulous patients [40, 43]. After 6 months, the first study showed similar bone formation and blood vessels in all treatments [40], while the second study showed higher BV and osteoid volume (OV) when SVF was added to scaffolds in comparison to the same scaffolds alone, with no adverse effects after 3 years [41].

Finally, through the setup of a clinical trial, it was shown that the SVF pellet, mixed with fibrin hydrogel and silicate HA microgranules, reduced pain and increased bone ossicles 1 year after implantation in low-energy fractures of the humerus. In the same study, a previous in vivo study was performed in a critically sized segmental defect of the femora in nude mice, in which porous silicate HA microgranules and hSVF increased mineralized volume, BV, and early stage of maturation, 2 months after implantation [42].

3.2. SVF and Cartilage

3.2.1. In Vitro Studies

One in vitro study was performed with hSVF or hADSCs harvested from the abdomen, and with human chondrocytes (Table 2). After 1 month, the addition of hSVF to chondrocyte cultures induced a higher amount of glycosaminoglycans (GAG) and increased cell proliferation than hADSCs cocultured with chondrocytes. In the same study, allogenic chondrocytes combined with allogenic SVF increased GAG and COLL II 2 months after they had been implanted in subcutaneous pouches of nude mice [43].

Table 2: In vitro, in vivo, and clinical studies on SVF in cartilage regeneration.

Experimental model
Treatment groups
Evaluations
Results
Ref.

In vitro:
Human chondrocytes () from healthy donors
hSVF or hADSCs () from abdomen
In vivo:
10 BALB/C nude mice (6 wks) with subcutaneous pouches
Allogenic SVF or ADSCs
Allogenic chondrocytes ()
In vitro:
Group 1: SVF pellet
Group 2: ADSC pellet
Group 3: chondrocyte pellet
Group 4: SVF+chondrocyte pellet (4 : 1)
Group 5: ADSCs+chondrocyte pellet (4 : 1)
In vivo:
Group 1: no treatment
Group 2: SVF+chondrocytes (4 : 1) in 2% alginate
Group 3: ADSCs+chondrocytes (4 : 1) in 2% alginate
Group 4: chondrocytes
In vitro:
Histology
GAG quantification
Cell proliferation
In vivo:
Histology
GAG quantification
IHC (COLL II)
In vitro:
1 mo
Group 4: ↑ GAG, chondrocytes % than group 5
In vivo:
2 mo
Group 2: ↑ GAG, COLL II protein than groups 1, 3, and 4
[43]

In vivo:
8 skeletally mature Dutch milk goats () with 2 osteochondral defects in the right troclea femoris and 2 defects in the right medial femoral condyles
Autologous SVF or ADSCs ()
COLL I/COLL III scaffold ()
Group 1: scaffold
Group 2: scaffold+SVF
Group 3: scaffold+ADSCs
Gross evaluations
Histology
Micro-CT
1 mo
Group 1: =gross appearance score, histological score than groups 2 and 3.
4 mo
Group 2: ↑ cartilage and SB regeneration than groups 1 and 3
Group 2: ↑ hyaline cartilage; ↓ fibrocartilage, cartilaginous tissue in SB than group 3
[44]

In vivo:
30 small tail Han sheep (6 mo) with OA in the right knee
Autologous SVF () or ADSCs from cervicothoracic region
HA scaffold (600–1500 kDa) (2.5 ml)
Group 1: saline (5 ml)
Group 2: scaffold
Group 3: scaffold+ADSCs ()
Group 4: scaffold+ADSCs ()
Group 5: scaffold+SVF
IHC (COLL X, SDF1)
3 mo
Group 5: ↓ COLL X protein than groups 3 and 4.
Group 5: ↑ SDF1 protein than groups 1 and 2
[7]

In vivo:
9 NOD/SCID mice with injured cartilage
hSVF () from abdomen
Group 1: PBS
Group 2: SVF+PRP
Movement recording
Histology
Histomorphometry
45 days
Group 2: ↓ time required that mice could move, cartilage lesion area; ↑ formed neocartilage than group 1
[45]

Clinical trial:
13 pz (65-82 yrs) with bilateral severe OA
Allogenic SVF () from abdomen
Group 1: SVF
Clinical evaluation
1 and 6 mo
Group 1: ↓ JKOM score, WOMAC score, VAS score than presurgery.
No differences between 1 and 6 mo
[46]

Clinical trial:
18 pz () with OA
Autologous SVF (35 ml) from abdomen
Group 1: SVF
Clinical evaluation
Radiography
3, 6, and 18 mo
Group 1: ↓ VAS score, WOMAC % than presurgery. No infection, thromboembolism, adverse reaction
18 mo
Group 1: =outerbridge score than presurgery
[47]

Clinical trial:
10 pz (≥50 yrs) with idiopathic knee OA
Allogenic SVF (5 ml) from abdomen
Group 1: SVF+PRP (3 ml)
Clinical evaluation
SF analysis
3, 6, 12, 18, and 24 mo
Group 1: ↓ WOMAC total, WOMAC stiffness, WOMAC pain, WOMAC physical function; ↑ six-minute walk distance than presurgery
24 mo
Group 1: no atypical cells in SF, restore SF properties, synovial metabolism; ↓ cartilage pathology than presurgery
[48]

Clinical trial:
4 pz (23-74 yrs) with OA of both knees
Autologous SVF (, , , and ) from periumbilical region
Group 1: SVF+PRP (3 ml) (4 injections every month)
Clinical evaluation
12 mo
Group 1: ↑ KOOS score pain, symptoms, ADL, sport/rec function, knee-related QOL. Pz regained normal functional activity
[49]

Clinical trial:
30 pz (>18 yrs) with OA
Autologous SVF from abdomen
Group 1: arthroscopic microfracture
Group 2: arthrosopic microfracture+SVF+PRP (5 ml)
Clinical evaluation
BM edema
12 mo
Group 2: ↓ BM edema than group 1
Group 1: ↓ WOMAC score during time
18 mo
Group 2: ↓ WOMAC score during time
Group 2: ↓ WOMAC score; ↑ Lysholm score than group 1
[50]

Clinical trial:
33 pz (>38 yrs) with OA
Autologous SVF (6 ml with cells) from abdomen
Group 1: arthroscopic microfracture
Group 2: arthroscopic microfracture+SVF
Clinical evaluation
BM edema
12 mo
Groups 1 and 2: ↓ VAS score, WOMAC score during time
24 mo
Group 2: ↓ VAS score, WOMAC score, Outerbridge score than group 1
Groups 1 and 2: ↑ Lysholm score during time
Group 2: ↓ BM edema during time
Group 1: ↑ BM edema during time
[51]

Clinical trial:
16 pz () with bilateral OA of grade II or III
Hyaluronic acid (4 ml)
Autologous SVF (4 ml) from abdomen
Group 1: arthroscopic debridement+SVF
Group 2: arthroscopic debridement+HA
Clinical evaluation
Radiography
1, 3, 6, and 12 mo
Group 1: ↓ VAS, WOMAC pain, WOMAC stiffness; ↑ ROM during time
Group 2: ↑ VAS, WOMAC pain, WOMAC stiffness; ↓ ROM than group 1
1 mo
Group 2: ↑ ROM during time
3, 6, and 12 mo
Group 2: ↓ ROM during time
1 and 3 mo
Group 2: ↓ VAS during time
6 and 12 mo
Group 2: ↑ VAS during time
Group 1: ↑ MOCART score during time
Group 1: ↑ MOCART score, complete tissue filling than group 2
Group 2: ↓ MOCART score during time
[52]

Clinical trial:
26 pz () with full-thickness chondral defect of the talar dome
Autologous SVF ( MSCs) from buttock
Group 1: arthroscopic marrow stimulation
Group 2: arthroscopic marrow stimulation+SVF
Clinical evaluation
Radiography
16-25 mo
Groups 1 and 2: ↓ VAS score; ↑ AOFAS score during time
Group 2: ↓ VAS score; ↑ AOFAS score, Tegner score, MOCART score than group 2
[53]

Clinical trial:
1 pz (36 yrs) with traumatic osteochondral lesion of the right medial femoral condyle 8 mo after injury
Autologous SVF () from abdomen
Group 1: microfracture+SVF
Clinical evaluation
BM edema
Radiography
24 mo
Group 1: ↑ IKDC, EQ-VAS than presurgery
12 and 24 mo
Group 1: recovery of CT; ↓ BM edema than presurgery
[54]

3.2.2. In Vivo Studies

Three in vivo studies were conducted in goats with osteochondral defects in the trochlea femoris and the medial femoral condyle [44], in sheep with knee OA [7], and in NOD/SCID mice with cartilage injury [45]. SVF was derived from nonspecified sites [44], the cardiothoracic region [7], or the abdomen [45] and was autologous [7, 44] or xenogenic [45] (Table 2).

In the first study, SVF and ADSCs were seeded onto COLL I/COLL III scaffolds and implanted into osteochondral defects. After 4 months, scaffolds with SVF induced higher hyaline cartilage and subchondral bone (SB) regeneration than a scaffold alone or a scaffold combined with ADSCs [44].

In the second study, in the presence of OA, SVF, combined with hyaluronic acid, reduced COLL X and increased stromal cell-derived factor 1 (SDF1) protein production in comparison to ADSCs or hyaluronic acid alone [7].

In the third study, hSVF mixed with PRP significantly reduced the recovery time (in terms of animal movement) and cartilage lesion area more than phosphate-buffered saline (PBS), 45 days from injection into cartilage injuries [45].

3.2.3. Clinical Studies

Eight clinical studies were performed in patients with OA, full-thickness chondral defect of the talar dome [50], and traumatic osteochondral defect of the right femoral condyle [54].

Allogenic or autologous SVF was obtained from the abdomen [42–44, 46–48, 50], periumbilical region [45], and buttocks [49] (Table 2).

In patients with OA, the treatments consisted of SVF alone [46, 47]; SVF and PRP [48, 49]; SVF, PRP, and arthroscopic microfracture [50, 51]; or SVF and arthroscopic debridement [52].

The injection of SVF alone reduced changes in Japanese Knee Osteoarthritis Measure (JKOM), Western Ontario and McMaster Universities Osteoarthritis Index (WOMAC), and visual analog scale (VAS) scores after 1, 3, 6, and 18 months [46, 47] with no infection, thromboembolism, or adverse reactions [47]. The combination of SVF and PRP reduced WOMAC scores and increased the six-minute distance parameter after 3 months, 6 months, 1 year, and 2 years [48] and ameliorated Knee injury and Osteoarthritis Outcome Score (KOOS), symptoms, and functional activity at 1 year [49]. Arthroscopic microfracture was accompanied with SVF injection with or without PRP. With the addition of PRP, bone marrow edema and WOMAC score was reduced and Lysholm score was increased after 12 and 18 months [50], while without PRP, bone marrow edema, VAS, WOMAC, and Outerbridge scores were reduced after 12 and 24 months [51]. In one study, in which OA was treated with arthroscopic debridement, the addition of SVF reduced VAS and WOMAC scores and increased the range of motion (ROM) more than hyaluronic acid at 1, 3, 6, and 12 months, while it increased the Magnetic Resonance Observation of Cartilage Repair (MOCART) score and complete tissue filling more than hyaluronic acid at 6 and 12 months [52].

Regarding the full-thickness chondral defect, SVF, associated with arthroscopic marrow stimulation, reduced the VAS score and increased the American Orthopedic Foot and Ankle Score (AOFAS) and the Tegner and MOCART scores more than the technique alone after 25 months [53].

In a traumatic osteochondral lesion, microfracture associated with SVF and fibrin sealant increased the International Knee Documentation Committee (IKDC) score, the EuroQol-visual analog scale (EQ-VAS) score, and the recovery of the cartilage thickness and reduced bone edema after 1 and 2 years [54].

3.3. SVF and Tendon

3.3.1. In Vivo Studies

Four in vivo studies were performed in rabbits [55–58] with transection of the midsubstance of the deep digital flexor (DDF) tendon [55], transection of the central one third of the flexor tendon [56], or with the supraspinatus tendon severed from the great trochanter [57, 58]. SVF was harvested from the inguinal region and was allogenic [55, 56] or autologous [57, 58] (Table 3).

Table 3: In vitro, in vivo, and clinical studies on SVF in tendon regeneration.

Experimental model
Treatment groups
Evaluations
Results
Ref

In vivo:
20 adult NZW rabbits (2.5-3 kg) with complete transaction of DDFT midsubstance
Allogenic SVF () from inguinal region
Group 1: PBS (0.2 ml)
Group 2: SVF
Histology
Histomorphometry
IHC (COLL I, COLL III)
Biomechanics
2 mo
Group 2: ↑ fibrillar linearity, fibrillar continuity, COLL I protein, ultimate load, energy absorption, stiffness; ↓ no. of capillaries in neotendon, COLL III protein than group 1
[55]

In vivo:
36 adult NZW rabbits (2.5-3 kg) with a complete transaction through the central one third of flexor tendon
Allogenic SVF () from inguinal region
Allogenic BMSCs () from iliac crest
Group 1: PBS (0.2 ml)
Group 2: SVF
Group 3: BMSCs
Biomechanics
3 and 8 wks
Groups 2 and 3: ↑ energy absorption, ultimate load, ultimate stress, yield load, stiffness than group 1
[56]

In vivo:
36 adult NZW rabbits (2-2.5 kg) with supraspinatus tendon severed from the great trochanter
Autologous SVF from inguinal region
Group 1: FG (1 ml)
Group 2: SVF+FG (1 ml)
Radiography
Biomechanics
1 mo
Group 2: ↑ maximum load, maximum strength than group 1
2 mo
Group 2: ↑ maximum load, maximum strength, stiffness than group 1
3 mo
Group 2: ↑ SNQ, stiffness than group 1
[57]

In vivo:
36 NZW rabbits (2-2.5 kg) with bilateral supraspinatus tendon severed from the great trochanter
Autologous SVF from inguinal region
Group 1: FG (1 ml)
Group 2: SVF+FG (1 ml)
Histology
IHC (COLL I, COLL III, BMP2)
Histomorphometry
Biomechanics
2 mo
Group 2: ↑ tendon-bone healing maturity, COLL I, COLL III, COLL I/COLL III, BMP2 protein, maximum load, maximum strength, stiffness than group 1
[58]

Clinical trial:
43 pz (29-55 yrs) with unilateral or bilateral NIAT
Allogenic SVF (4 ml) from abdomen
Group 1: PRP (4 ml)
Group 2: SVF
Radiography
Clinical evaluation
6 mo
Groups 1 and 2: ↓ VAS; ↑ MR size, US size, PD than presurgery
Group 1: ↓ MR-Si than group 2
[59]

Clinical trial:
44 pz (18-55 yrs) with unilateral or bilateral chronic tendinopathy of the Achilles tendon
Allogenic SVF (4 ml) from abdomen
Group 1: PRP (4 ml)
Group 2: SVF
Clinical evaluation
2 wks, 1 mo, 4 mo, and 6 mo
Groups 1 and 2: ↓ VAS pain scale; ↑ VISA-A score, AOFAS score, SF-36 than presurgery
2 wks
Group 2: ↓ VAS pain scale; ↑ VISA-A score, AOFAS score than group 1
1 mo
Group 2: ↓ VAS pain scale than group 1
[60]

SVF was injected in the complete transection of the DDF tendon, and after 2 months, fibrillar linearity and continuity, COLL I production, ultimate load, energy absorption, and stiffness increased, while the number of capillaries and COLL III production decreased [55].

SVF and BMSCs, injected in the complete transection of the flexor tendon, increased energy absorption, ultimate load, ultimate stress, yield load, and stiffness after 3 and 8 weeks [56].

After 1, 2, and 3 months from SVF injection into the supraspinatus tendon severed from the great trochanter, maximum load, maximum strength, stiffness, and signal-to-noise quotient (SNQ) increased [57], as well as tendon-bone healing, COLL I, and bone morphogenetic protein 2 (BMP2) after 2 months [58].

3.3.2. Clinical Studies

The two clinical studies were performed. One study included 45 patients with noninsertional Achilles tendinopathy (NIAT) [59], and the other study included 44 patients with chronic tendinopathy of the Achilles tendon [60]. SVF was allogenic and harvested from the abdomen [59, 60] (Table 3).

In the first study, both PRP and SVF reduced VAS and increased MR size, US size, and peri- and intratendinous flow (PD), while only SVF increased MR signal intensity (MR-Si) after 6 months [59].

Similarly, PRP and SVF were compared in Achilles tendon tendinopathy. Both PRP and SVF reduced VAS and increased the Victorian Institute of Sports Assessment—Achilles (VISA-A) questionnaire score, AOFAS score, and SF-36 score after 2 weeks, 1, 2, 4, and 6 months, showing better results with SVF [60].

3.4. AECs and Bone

3.4.1. In Vitro Studies

Four in vitro studies were conducted with human AECs (hAECs) [61–64] (Table 4).

Table 4: In vitro, in vivo, and clinical studies on AECs in bone regeneration.

Experimental model
Treatment groups
Evaluations
Results
Ref

In vitro:
hAECs
Purchased hADSCs
Group 1: hAECs () cultured in OM
Group 2: hADSCs () cultured in OM
Group 3: hAECs ( per pellet) cultured in CM
Group 4: hADSCs ( per pellet) cultured in CM
RT-PCR (RUNX2, SOX9)
Alizarin red staining
Alcian blue staining
IHC (OCN, AGG, and COLL II)
3, 7, and 14 days
Group 1: ↑ RUNX2 gene expression, OCN protein than group 2
Group 3:↑ AGG and COLL II proteins, SOX9 gene expression, than group 4
2 and 4 wks
Group 1: ↑ mineralization during time
[61]

In vitro:
hAECs
, 1 mT for 30 min each time, 2 for days with an interval of 12 hours
Group 1: hAECs in NM
Group 2: hAECs in OM
Group 3: hAECs in NM+PEMF
Group 4: hAECs in OM+PEMF
RT-PCR (ALP, OCN, BMP2, RUNX2, NRF2, and KEAP1)
ALP activity
IHC (OCN)
Calcium deposition
3, 7, and 11 days
Group 4: ↑ ALP, OCN gene expression than groups 1-3
Group 3: ↑ ALP, OCN than groups 1 and 2
Group 2: ↑ ALP, OCN than group 1
7 days
Group 3: ↑ ALP activity, calcium deposition than group 1
7, 11, and 21 days
Group 2: ↑ ALP activity than groups 1 and 3
Group 4: ↑ ALP activity, OCN protein, calcium deposition than groups 1-3
Group 3: ↑ OCN protein than group 1
21 days
Group 2: ↑ OCN protein, calcium deposition than groups 1 and 3
3, 11, and 21 days
Group 4: ↑ BMP2, RUNX2, NRF2, and KEAP1 gene expression than groups 1-3
[62]

In vitro:
Conditioned medium from hAECs after 24 hrs of culture
Purchased hFOB1.19 ()
Group 1: hFOB1.19 in NM
Group 2: hFOB1.19 in conditioned medium
Group 3: hFOB1.19 in conditioned medium+TGFβ1 antibody (5 μg/ml)
Cell proliferation
Cell migration
ALP activity
RT-PCR (ALP, OCN, OPN, and RUNX2)
2 hrs
Group 2: ↑ hFOB1.19 migration than group 1
1, 2, and 3 days
Group 2: ↑ hFOB1.19 proliferation than group 1
6 days
Group 2: ↑ hFOB1.19 ALP activity, ALP, OCN, OPN, RUNX2 gene expression than group 1
Group 3: ↓ hFOB1.19 ALP activity, ALP, OCN gene expression than group 2
6 hrs
Group 3: ↓ hFOB1.19 migration than group 2
[63]

In vitro:
hAECs (3rd-5th passage) from 6 patients ()
Group 1: hAECs in NM
Group 2: hAECs+mechanical stretch (maximum uniaxial stretched length (7.35 cm) for 2, 6, 12, and 24 hrs with 5% elongation at a frequency of 0.5 Hz
Group 3: hAECs in OM for 21 days
Group 4: hAECs+OM for 21 days+mechanical stretch for 2, 6, 12, and 24 hrs
Group 5: hAECs+Runx2 shRNA+OM for 24 hrs
Group 6: hAECs+Runx2 shRNA+mechanical stretch for 24 hrs
Group 7: hAECs+Runx2 shRNA+OM+mechanical stretch for 24 hrs
RT-PCR (ALP, OCN, RUNX2, β-catenin, and Cyclin D1)
WB (ALP, OCN, RUNX2, β-catenin, and Cyclin D1)
12 hrs
Groups 3 and 4: ↑ OCN protein than group 1
6, 12, and 24 hrs
Group 3: ↑ RUNX2, ALP, and OCN gene expression than group 1
6 and 12 hrs
Group 4: ↑ RUNX2, ALP, OCN gene expression and protein than groups 2 and 3
12 hrs
Group 3: ↑β-catenin, RUNX2, Cyclin D1 gene expression and protein than group 1
Group 4: ↑β-catenin, RUNX2, Cyclin D1 gene expression and protein than groups 2 and 3
24 hrs
Groups 5-7: ↓ RUNX2, OCN gene expression, and protein than groups 2-4
[64]

In vivo:
6 adult sheep (2 yrs) that need bilateral sinus augmentation
Allogenic oAECs () from 3 slaughtered sheep (3 mo of pregnancy)
CaP (HA-βTCP: 30/60) scaffold
Group 1: scaffold
Group 2: scaffold+AECs
Micro-CT
Histology
Histomorphometry
IHC (VEGF)
45 days
Group 2: ↑ VEGF, VA; ↑ BA than group 1
3 mo
Group 2: ↓ VEGF, VA; ↑ BA than group 1
[65]

In vivo:
16 SD rats (7 wks) with unilateral maxillary alveolar defect
hAECs
βTCP scaffold ()
Group 1: scaffold
Group 2: scaffold+AECs
Micro-CT
Histology
Histomorphometry
IHC (CD68, VEGF)
1 mo
Group 2: ↓ Tb.Sp, CD68 than group 1
2 mo
Group 2: ↑ BMD, BV/TV, Tb.N, BA, VEGF area, CD68 than group 1
[66]

In vivo:
3 nude mice with subcutaneous pouches
hAECs, hBMSCs, hAFMSCs
βTCP scaffold ()
Group 1: scaffold
Group 2: scaffold+hAEC
Group 3: scaffold+hBMSCs
Group 4: scaffold+hAFMSCs
Histology
IHC (OPN, OCN)
1 mo
All groups: no well-mineralized islands
Groups 2-4: viable cells; ↑ OPN, OCN proteins than group 1
[67]

In the first study, hAECs and hADSCs were compared in terms of osteogenic and chondrogenic differentiation. After 3, 7, and 14 days of culture, hAECs showed higher RUNX2 and SOX9 gene expression, OCN, aggrecan, and COLL II protein than hADSCs. After 14 and 28 days, mineralization was more pronounced in hAECs than in hADSCs [61].

Keeping in terms of osteogenic differentiation, hAECs, cultured in OM and stimulated with pulsed electromagnetic fields (PEMFs), increased ALP, BMP2, RUNX2, nuclear factor erythroid 2-related factor 2 (NRF2), Kelch-like ECH-associated protein 1 (KEAP1) and OCN gene expression, ALP activity, and OCN protein and calcium deposition in comparison to cells in NM or in OM, without PEMFs, after 3, 7, 11, and 21 days. The addition of PEMFs in NM also induced ALP and OCN gene expression, ALP activity, and OCN protein and calcium deposition, after 3, 7, 11, and 21 days [62].

The effect of conditioned medium from hAECs, produced after 24 hours of culture, on human fetal osteoblast cell line (hFOB1.19) was evaluated in one study. Conditioned medium increased hFOB1.19 cell migration and proliferation; ALP activity; and ALP, OCN, OPN, and RUNX2 gene expression after 2 hours and 1, 2, 3, and 6 days. The addition of the antibody against transforming growth factor β (TGFβ1) in the culture medium reduced ALP activity, ALP and OCN gene expression, and cell migration after 6 hours and 6 days [63].

Finally, a study showed how mechanical stretch (with a maximum uniaxial stretched length of 7.35 cm, 5% of elongation, and frequency of 0.5 Hz) increased OCN, RUNX2, ALP, β-catenin, and Cyclin D1 gene expression and protein expression 6, 12, and 24 hrs after stimulation, with a synergic effect when OM was added [64].

3.4.2. In Vivo Studies

Three studies were conducted in sheep submitted to sinus augmentation [65], in rats with maxillary alveolar defect [66], and in subcutaneous pouches of nude mice [67]. AECs were allogenic [65] or xenogenic [66, 67] (Table 4).

Ovine AECs, seeded onto a calcium phosphate (CaP) scaffold and implanted into alveolar defects, increased vascular endothelial growth factor (VEGF), vascular area, and new bone more than the scaffold alone after 3 months [65]. Similarly, a βTCP scaffold, seeded with hAECs, reduced Tb.Sp and CD68 cells and increased BMD, BV, trabecular number (Tb.N), BA, and VEGF more than the scaffold alone after 1 and 2 months [66].

Three types of cells, namely, hAECs, hBMSCs, and hAFMSCs, were seeded onto the βTCP scaffold and implanted into subcutaneous pouches, showing higher viable cells and higher OPN and OCN protein production than the scaffold alone, regardless of cells, after 1 month [67].

3.5. AECs and Cartilage

3.5.1. In Vitro Studies

There were two in vitro studies conducted with hAECs [68, 69] (Table 5).

Table 5: In vitro, in vivo, and clinical studies on AECs in cartilage regeneration.

Experimental model
Treatment groups
Evaluations
Results
Ref

In vitro:
Cartilage samples with defect of 2 mm
Human chondrocytes from femoral heads or knee ()
hBMSCs from femoral heads of donors (50-70 yrs)
hAECs
hAMSC
HAM scaffold
Group 1: scaffold seeded with chondrocytes+chondrocyte pellet ()
Group 2: scaffold seeded with BMSCs+BMSC pellet ()
Group 3: scaffold seeded with AECs+AEC pellet ()
Group 4: scaffold seeded with AMSCs+AMSC pellet ()
Histology
IHC (COLL I, COLL II)
2 mo
Group 1: =repair of lesioned area, ICRS score than groups 2, 3, and 4
Group 4: ↑ COLL II protein than group 1
Group 2: ↑ COLL I protein than groups 1, 3, and 4
[68]

In vitro:
hAECs in micromasses ()
Group 1: micromasses+TGFβ1 (1 ng/ml)
Group 2: micromasses+BMP7 (100 ng/ml)
RT-PCR (SOX9, COLL II, PG)
3 days
Group 2: ↑ SOX9, COLL II gene expression than group 1
3 wks
Group 1: ↑ SOX9, COLL II gene expression than group 2
Groups 1 and 2: ↑ PG gene expression during time
[69]

In the first study, cartilage samples, with a defect of 2 mm, were ex vivo treated with a human amniotic membrane (HAM) scaffold seeded with human chondrocytes, hBMSCs, hAECs, or hAMSCs and added with the respective cell pellets. After 2 months of culture, all treatments showed the same degree of repair and International Cartilage Repair Society (ICRS) score [68].

In the second study, micromasses of hAECs were cultured with TGFβ1 or BMP7 for 3 days and 3 weeks. The addition of BMP7 increased SOX9 and COLL II gene expression at 3 days, while TGFβ1 increased it to 3 weeks. Both treatments increased proteoglycan gene expression during time [69].

3.6. AECs and Tendon

3.6.1. In Vitro Studies

One in vitro study was conducted with ovine AECs alone or cocultured with fetal or adult ovine tenocytes or tendons (Table 6). After 7, 14, and 28 days of culture, fetal tendons increased AEC migration, while adult tenocytes or tendons reduced AEC proliferation. Fetal tenocytes or tendons increased AEC proliferation and OCN, tenomodulin (TNMD), and scleraxis (SCX) protein production. In addition, fetal tendons reduced the telomere area (TEA) and feret maximum (TEF) and mean densitometric (MEAND) values, while adult tendons or tenocytes reduced COLL III gene expression and TNMD protein [70].

Table 6: In vitro, in vivo, and clinical studies on AECs in tendon regeneration.

Experimental model
Treatment groups
Evaluations
Results
Ref

In vitro:
oAECs ()
Tenocytes () or tendons from heel of adult sheep (2-3 yrs) or fetus sheep (2-3 mo)
Group 1: AECs
Group 2: AECs+fetal tenocytes
Group 3: AECs+adult tenocytes
Group 4: AECs+fetal tendons
Group 5: AECs+adult tendons
AEC proliferation
AEC migration
AEC telomere analysis
RT-PCR (COLL I, COLL III)
IHC (OCN, TNMD, and SCXB)
7 days
Group 4: ↑ AEC migration than group 1
2 wks
Groups 3 and 5: ↓ AEC proliferation than group 1
1 mo
Groups 2 and 4: ↑ AEC proliferation, OCN, TNMD, and SCXB proteins than group 1
Group 4: ↓ TEA, TEF, and MEAND than group 1
Groups 2-5: ↑ COLL I gene expression than group 1
Group 3: ↓ COLL III gene expression than group 1
Groups 3 and 5: ↓ TNMD protein than group 1
[70]

In vivo:
15 horses with acute SDFT monolateral injuries
oAECs () from slaughtered sheep at 3 mo of pregnancy
Group 1: healthy tendons
Group 2: AECs
Histology
IHC (COLL III, COLL I, and Ki-67)
2 mo
Group 2: ↓ COLL III protein; ↑ COLL I, Ki-67 proteins than group 1, no immunological reaction
[71]

In vivo:
6 adult horses (3-7 yrs) with acute or chronic SDFT tendinopathy in the midmetacarpal region;
oAECs () from slaughtered sheep at 3 months of pregnancy
Group 1: AECs
Ultrasonography
Histology
6 mo
Group 1: cross-sectional area similar to an healthy tendon, collagen fibers parallel to the longitudinal axis of the tendon, no neovascularization than presurgery
[72]

In vivo:
23 sheep (40 kg) with a bilateral full-thickness FDST hole (3 mm diameter)
Allogenic oAECs () from slaughtered sheep at 3 months of pregnancy
Group 1: FG
Group 2: AECs+FG
Histology
RT-PCR (COLL I, COLL III)
Histomorphometry
Biomechanics
2 wks
Group 2: ↑ tendon microarchitecture than group 1
1 mo
Group 2: ↑ tendon microarchitecture, proliferation, COLL I gene expression, maximum failure load, stiffness; ↓ VA, leukocyte infiltration, cellularity, COLL III gene expression than group 1
[73]

In vivo:
3 adult sheep (50 kg) with bilateral defect in the Achilles tendon middle portion
Allogenic oAECs () from slaughtered sheep at 60-80 days of pregnancy
Group 1: no treatment
Group 2: AECs
Histology
1 mo
Group 2: absent inflammation-proliferating cells, collagen fibers started to be oriented along a longitudinal axis
Group 1: no tendon healing, no proliferating cells, nonorganized collagen fibers
[74]

In vivo:
18 Appenninica breed sheep (2 yrs) with a full-thickness hole in FDST (3 mm diameter)
Allogenic oAECs ()
Group 1: FG
Group 2: FG+oAECs
Histology
IHC (COLL I, CD45, CD68, CD86, and MMR/CD206)
RT-PCR (CD86, CD206, YM1, IL10, and IL12b)
7, 14, and 28 days
Group 2: ↑ tissue organization; ↓ leukocyte infiltration, IL12 b gene expression than group 1
14 and 28 days
Group 2: ↓ macrophage infiltration, CD86 gene expression, CD86 positive cells; ↑ IL10 gene expression than group 1
28 days
Group 2: ↑ COLL I; ↓ cellularity, YM1 gene expression, CD206-positive cells than group 1
[75]

In vivo:
29 adult sheep (2 yrs) with bilateral Achilles tendon defects (5 mm diameter)
hAECs ()
Group 1: FG
Group 2: FG+hAECs
RT-PCR (COLL I, COLL III, TNMD, THBS4, CD206, IL10, VEGF, CD68, IL12b)
IHC (COLL I, COLL III, VEGF)
Histology
Biomechanics (maximum load, stiffness)
1 mo
Group 2: rapid recovery of the tissue; sporadic edema; regular longitudinal profile; ↑ maximum failure load; stiffness; regularly arranged fibers; COLL I gene expression and protein, TNMD, THBS4, CD206, and IL10 gene expression; ↓ COLL III and VEGF gene expression and protein, CD86, and IL12b gene expression than group 1
[76]

Abbreviations: hSVF = human stromal vascular fraction; hBMSCs = human bone marrow mesenchymal stem cells; yrs = years; hrs = hours; wks = weeks; BMSCs = bone marrow mesenchymal stem cells; SVF = stromal vascular fraction; ALP = alkaline phosphatase; RT-PCR = reverse transcriptase-polymerase chain reaction; RUNX2 = runt-related transcription factor 2; COLL = collagen; OSX = Osterix; hADSCs = human adipose-derived mesenchymal stem cells; ADSCs = adipose-derived mesenchymal stem cells; mm = millimeters; BA/TA = bone area/tissue area; PLCL = poly(L-lactide-co-caprolactone); NM = normal medium; CM = chondrogenic medium; OM = osteogenic medium; SOX = sex-determining region Y boxes; OCN = osteocalcin; ACAN = aggrecan; mo = months; IHC = immunohistochemistry; Micro-CT = microcomputed tomography; HA = hydroxyapatite; cm = centimeters; BV/TV = bone volume/tissue volume; CD = cluster of differentiation; βTCP = beta-tricalcium phosphate; CFU = colony forming units; ml = milliliters; BA = bone area; OPN = osteopontin; PRP = platelet-rich plasma; mg = milligrams; PCL-DCB = polycaprolactone-decellularized bovine bone extracellular matrix; BV = bone volume; VA = vascular area; PLA = polylactic acid; DBM = demineralized bone matrix; SD = Sprague-Dawley; BVF = bone volume fraction; Tb.Th = trabecular thickness; PLGA = poly(lactide-co-glycolide); NZW = New Zealand White; BMD = bone mineral density; Tb.Sp = trabecular separation; PMN = polymorphonuclear; ONJ = osteonecrosis of the jaw; pz = patients; BCP = biphasic calcium phosphate; OV/TV = osteoid volume/tissue volume; GAG = glycosaminoglycans; kg = kilograms; SB = subchondral bone; OA = osteoarthritis; kDa = kilodaltons; SDF = stromal cell-derived factor; PBS = phosphate-buffered solution; NOD/SCID = nonobese diabetic/severe combined immunodeficiency; JKOM score = Japanese Knee Osteoarthritis Measure; VAS score = visual analog scale score; WOMAC score = the Western Ontario and McMaster Universities Osteoarthritis score; SF = synovial fluid; KOOS score = Knee Injury and Osteoarthritis Outcome Score; ADL = Activities of Daily Living; QOL = quality of life; BM = bone marrow; ROM = range of motion; MOCART = magnetic resonance observation of cartilage repair tissue; MSCs = mesenchymal stem cells; AOFAS score = American Orthopedic Foot and Ankle Score; IKDC = International Knee Documentation Committee; EQ-VAS = EuroQol-visual analog scale; CT = cartilage thickness; DDFT = deep digital flexor tendon; no. = number; FG = fibrin glue; SNQ = signal-to-noise quotient; BMP = bone morphogenetic protein; NIAT = noninsertional Achilles tendinopathy; PD = peri- and intratendinous flow; MR-Si = MR signal intensity; VISA-A score = Victorian Institute of Sports Assessment—Achilles questionnaire; SF-36 = Short Form 36 Health Survey; AECs = amniotic endothelial cells; hAECs = human amniotic endothelial cells; NRF2 = nuclear factor erythroid 2-related factor 2; KEAP1 = Kelch-like ECH-associated protein 1; PEMF = pulsed electromagnetic field; Hz = Hertz; min = minutes; hFOB = human fetal osteoblastic; TGFβ1 = transforming growth factor β1; μg = micrograms; VEGF = vascular endothelial growth factor; oAECs = ovine AECs; CaP = calcium phosphate; Tb.N = trabecular number; hAFMSCs = human amniotic fluid mesenchymal stem cells; hAMSCs = human amniotic membrane mesenchymal stem cells; HAM = human amniotic membrane; ICRS score = International Cartilage Repair Society; PG = polygalacturonase; TNMD = tenomodulin; SCXB = scleraxis; TEA = telomere area; TEF = feret maximum; MEAND = mean densitometric value; SDFT = superficial digital flexor tendon; FDST = flexor digitorum superficialis tendon.

3.6.2. In Vivo Studies

Six in vivo studies were conducted in horses [71, 72] with monolateral acute superficial digital flexor tendon (SDFT) injuries [71] and acute or chronic SDFT injury in the midmetacarpal region [72] or in sheep [73–76] with a bilateral full-thickness hole of flexor digitorum superficialis tendon (FDST) [73, 75] or bilateral defect of the middle portion of the Achilles tendon [74, 76]. AECs were xenogenic [71, 72, 76] or allogenic [73–75] (Table 6).

A reduction in COLL III and an increase in COLL I and Ki-67 proteins after 2 months from ovine AEC injection into SDFT injuries were observed [71].

Ovine AECs induced a cross-sectional area similar to the healthy tendon, parallel collagen fibers, and no vascularization in SDFT tendinopathy after 6 months [72].

Ovine AECs, injected into the hole of FDST, induced greater improvement in tendon microarchitecture, proliferation index, COLL I gene expression, maximum failure load, and stiffness, as well as greater reduction in vascular area (VA); leukocyte infiltration; macrophage infiltration; and CD86, IL12b, and COLL III gene expression than fibrin glue (FG) after 7, 14, and 28 days [73, 75].

Finally, 1 month after ovine AEC injection, an increase in collagen fibers and no inflammation were observed in defects of the Achilles tendon [74], with a rapid recovery; higher biomechanics; higher TNMD, THBS4, CD206, and IL10 gene expression; and lower CD86 and IL12b gene expression in comparison to FG [76].

4. Discussion

Treatments for the regeneration of musculoskeletal tissue disorders of bone, cartilage, and tendon could be conservative or surgical. Rehabilitation, anti-inflammatory drugs, biophysical therapy, platelet derivatives (i.e., PRP) or cell therapy [1, 77], microfracture, arthroscopy [1], and the implantation of grafts or scaffolds or hydrogels [78] are the main treatment options alone or in combination. Moreover, due to the lesion type and site or the patient characteristics (i.e., age, comorbidities, and lifestyles), some treatments reportedly do not have the ability to regenerate the original healthy tissue or have a short-term or incomplete effect.

Preclinical research has always been focused on finding increasingly innovative and less invasive therapies. In this regard, MSCs, in particular ADSCs and BMSCs, but also MABs and FAPs, have been used in musculoskeletal tissue regeneration, even if they show some limitations: the necessity of two surgical steps (one for the harvesting and one for the implantation after in vitro expansion), patient morbidity, and the necessity to obtain a huge number of cells through in vitro expansion with associated risks.

Nowadays, a novel source of MSCs and new techniques to isolate such cells are being studied. Both SVF and AECs are spreading as new protagonists in the field of cell-based therapies for regenerative medicine, starting from the first applications in cardiology, and it is interesting to note that the only study in which both were used was aimed at cardiac repair [79].

In the field of regenerative medicine, the use of SVF has a relatively recent history. One of the pivotal studies in the introduction of the employment of SVF for clinical purposes dates to 2001, when Zuk et al. described the characterization of multilineage cells harvested from human adipose tissue and called them “processed lipoaspirate.” In this study, it was demonstrated that the collected SVF contained not only adipose cells but also mesodermal or mesenchymal origin cells, as well as pericytes, endothelial cells, and smooth muscle cells. The evidence of the cells’ capability to differentiate in vitro towards adipogenic, chondrogenic, myogenic, and osteogenic lineages opened the possibility of exploiting this as a new source for tissue regeneration [12]. The protocol described in this paper became a reference to isolate SVF and study its composition; however, until about ten years ago, the investigation was limited to in vitro evaluation of SVF derived from mainly rodent models and tested for characterization and regeneration, with particular reference to cardiac muscle [80, 81]. One of the first applications for a musculoskeletal system in vivo available in literature compared the efficacy of SVF and BMSCs in the treatment of OA in an equine model, without however obtaining any evident results in favour of neither treatment [82]. Since then, the number of studies has increased, while the application of SVF in clinical trials is fairly recent, as evidenced by this review.

As for AECs, despite the extensive literature history, it was only after the First International Workshop on Placenta-Derived Stem Cells when it was finally pointed out the field of application of mesenchymal stromal cells isolated from various parts of the placenta or epithelial cells isolated from the amniotic membrane in regenerative medicine. The findings suggested that the main applications were oriented mainly towards hepatic and cardiac repair and neurological disorders [83].

This review focuses on preclinical and clinical studies on an innovative source of cells (amniotic membrane) and a one-step surgical technique not based on in vitro expanded cells (SVF transplantation) for musculoskeletal tissue regeneration.

In the last 10 years, 48 preclinical and clinical studies employed SVF (73%) [7, 17, 28–60] or AECs (27%) [61–76] in musculoskeletal tissue regeneration.

n general, in the studies of these cell sources, researchers prevalently performed in vivo studies [7, 29, 31–39, 42–45, 56–58, 65–67, 71–76], followed by clinical ones [40–42, 46–54, 59, 60] and in vitro studies [17, 28–30, 43, 61–64, 68–70]. As for AECs, no clinical studies have yet been conducted, and the only two studies that analysed AEC behavior in cartilage were in vitro [68, 69]. For both cell types, most of the studies regarded bone regeneration [17, 28–42, 61–67], followed by cartilage [7, 43–54, 68, 69] and tendon [55–60, 70–76]. SVF was prevalently implanted with scaffolds for bone regeneration: HA [31, 32], βTCP [32, 40, 41], devitalized hypertrophic cartilage pellet [33], deproteinized sterilized bovine bone [34], PCL-DCB [35], DBM [36], PLA [36], PLGA [37], or fibrin hydrogel added with porous silicate HA microgranules [42]. Conversely, in cartilage regeneration, SVF was prevalently injected without scaffolds [43, 45], especially in the clinical studies [46–54], even if two studies employed COLL I/COLL III [44] or hyaluronic acid [7]. All the studies on tendon employed SVF without scaffolds [55–60].

In vivo AECs were seeded onto CaP [65] or βTCP [66, 67] scaffolds in bone defects, while in tendon defects the authors injected AECs without scaffolds [70–76].

For both SVF and AECs, the results of the preclinical studies were obtained prevalently with histology and/or histomorphometry [29, 31–38, 42–45, 55, 58, 65–68, 71–76], protein production detection through immunohistochemistry (IHC) [7, 17, 31, 34, 36, 38, 42, 55, 58, 60, 65–68, 71, 75, 76], and micro-CT [17, 31, 33, 35, 37, 38, 44, 65, 66] and RT-PCR for gene expression analysis [23, 30, 34, 61–63, 69, 70, 73, 75, 76].

The preclinical results showed that SVF increased BMSCs’ osteogenic differentiation, ALP activity, and the calcium content of THP1 cells [28, 29], and it was able to differentiate towards osteogenic, chondrogenic, or tenogenic lineages both in vitro and in heterotopic sites [17, 30–35, 43, 55]. SVF also increased BV, defect filling, Tb.Th, and cartilage or tendon regeneration in animal models of calvaria [35, 36], ulnar [37], ONJ-like [38], lameness [39], osteochondral [44], OA [7], cartilaginous [45], flexor tendon [55, 56], and supraspinatus tendon [57, 58] defects.

AECs have an osteogenic [61, 62, 64, 67], chondrogenic [69], or tenogenic [70] differentiation ability, in vitro and in heterotopic sites; increased BA, vessel formation, BMD in sinus defects [65, 66], and regenerated ex vivo cartilage defects [69]; and acute or chronic SDFT, FDST, and Achilles [71–74] tendon lesions.

Clinical studies were conducted in edentulous patients [40, 41] or in patients affected by humerus fractures [42], OA [46–52], full-thickness chondral defects [53], osteochondral lesions [54], NIAT [59], and Achilles tendon lesions [60]. The measurements were performed with micro-CT [41, 42], histology and histomorphometry [40–42], clinical scores (WOMAC, KOOS, VAS, Lysholm, JKOM, IKDC, PD, VISA-A, AOFAS and EQ-VAS scores, and ROM) [45–54, 59, 60] and radiography (MOCART score, MR-Si, and MR size) [47, 52, 53, 59].

In these studies, SVF could be employed not only as autologous cells but also as allogenic ones, as observed in some studies in which allogenic SVF was used in rabbits [37, 55, 56], mice [38, 43], and in patients affected by OA [46, 48] or tendinopathy [59, 60]. One of the major potential advantages of AECs is that, unlike other cells, they can be used as allogeneic or xenogenic cells. Allogenic AECs were implanted into sheep [65, 73–75] while xenogenic ones in rats [66], horses [71, 72], and sheep [76]. To obtain an idea on other ongoing clinical trials, a further search was carried out on http://www.clinicaltrial.gov/. From this search, six clinical trials were found using SVF in OA patients, 2 in bone and 1 in tendon defects. However, the absence of clinical trials regarding AECs underlines that much remains to be explored on the potential of these cells.

More precisely, in OA patients, randomized clinical studies, single or triple blind in phase I, II, or III, are evaluating the effects of SVF injections into knees, also comparing SVF with corticosteroids, umbilical cord MSCs, BMSCs, or activated PRP. The enrolled patients are 20, 30, 40, 52, 200, and 480, and the evaluations are performed with clinical scores, ROM, adverse event recording, radiography, and arthroscopic cartilage repair assessment after 1, 3, 6, 9, 12, 18, and 24 months.

In 8 and 5 patients, two clinical trials in phase II have evaluated the effect of SVF, seeded or not onto composite HA microgranules after embedding in a fibrin gel, in the treatment of proximal humeral fractures in osteoporotic patients or in craniofacial injuries. Radiological and clinical scores and average tissue thickness are evaluated at 6 and 12 weeks and 1, 3, 6, 9, and 12 months. [42]

One randomized, quadruple blind, phase II clinical trial has been performed in 52 patients to evaluate the ability of autologous SVF in improving the repair of chronic rotator cuff tears, and the evaluations of clinical measurements, change in muscle stiffness, and change in fatty infiltration are carried out at 6 weeks and at 6, 12, and 24 months [58].

The overview of the state-of-the-art in the study of these cell sources highlights some limitations in the comparison of SVF or AECs with other cell types, which is represented by the paucity of preclinical studies and the insufficient number of clinical trials conducted. However, despite these limits, some considerations about the comparison of AECs and SVF vs. other cell types can be made. For example, a superiority of SVF over ADSCs was found in inducing ALP activity and calcium content in THP1 cells [29], in osteogenic differentiation ability [17, 35], in chondrogenic differentiation [43], and in osteochondral or OA defect regeneration [7, 44]. In addition, SVF regenerated biomechanical properties of flexor tendon defects in a similar manner to BMSCs [56]. On the other hand, AECs showed higher osteogenic differentiation ability than ADSCs [61], but similar osteogenic differentiation to BMSCs and AFMSCs [67] and similar chondral defect regeneration to BMSCs and AMSCs [68].

5. Conclusions

SVF and AECs are two promising cells for regenerating bone, cartilage, and tendon. Both show advantages in terms of application in a one-step approach, which has become one of the main goals for streamlining and reducing bias in surgical procedures. In addition, harvesting procedures are easy and less invasive in comparison to other cell sources. In addition, as for AECs, many ethical concerns have been overcome when dealing with waste material; at the same time, the harvesting technique of SVF might make this source exploitable for autologous use also in patients with a particular pathology or undergoing therapies. Several preclinical studies affirm the regenerative ability of both SVF and AECs; less clinical studies on SVF exist, while no studies on AECs exist. Most of the clinical studies deal with SVF in patients affected by OA. Additionally, even if there are only a few studies that compare SVF or AECs with other cell types, they showed that SVF and AECs behave in a better or similar way to in vitro culture-expanded ADSCs or BMSCs, without showing the now known limitations linked to the cells expanded in culture. An advantage of AECs, compared to other cells, is its use in the form of allogeneic or xenogenic cells, although, in order to affirm this, clinical studies are necessary.

Data Availability

The data supporting this systematic review are from previously reported studies and datasets, which have been cited.

Conflicts of Interest

The authors declare that they have no conflict of interests.

Acknowledgments

This work was partially supported by the Ministry of Health-Ricerca Corrente through the Rizzoli Orthopaedic Institute and by PRIN: Progetti di Ricerca di Rilevante Interesse Nazionale—Bando 2015 (Amniotic epithelial stem cells (AECs) vs. adipose-derived mesenchymal stem cells (ADSCs): translational potential as biological injective treatment for osteoarthritis).

References

G. A. Brown, “AAOS clinical practice guideline: treatment of osteoarthritis of the knee: evidence-based guideline, 2nd edition,” The Journal of the American Academy of Orthopaedic Surgeons, vol. 21, no. 9, pp. 577–579, 2013.

S. Deng, Z. Sun, C. Zhang, G. Chen, and J. Li, “Surgical treatment versus conservative management for acute achilles tendon rupture: a systematic review and meta-analysis of randomized controlled trials,” The Journal of Foot and Ankle Surgery, vol. 56, no. 6, pp. 1236–1243, 2017.

S. Meena, A. Hooda, P. Sharma, S. Mittal, J. Sharma, and B. Chowdhury, “Operative versus non operative treatment of displaced intraarticular fracture of calcaneum: a meta-analysis of randomized controlled trials,” Acta Orthopaedica Belgica, vol. 83, no. 1, pp. 161–169, 2017.

A. I. Caplan and D. Correa, “The MSC: an injury drugstore,” Cell Stem Cell, vol. 9, no. 1, pp. 11–15, 2011.

S. A. Rodeo, “Cell therapy in orthopaedics: where are we in 2019?” Bone Joint J., vol. 101-B, no. 4, pp. 361–364, 2019.

M. Strioga, S. Viswanathan, A. Darinskas, O. Slaby, and J. Michalek, “Same or not the same? Comparison of adipose tissue-derived versus bone marrow-derived mesenchymal stem and stromal cells,” Stem Cells and Development, vol. 21, no. 14, pp. 2724–2752, 2012.

X. Lv, J. He, X. Zhang et al., “Comparative efficacy of autologous stromal vascular fraction and autologous adipose-derived mesenchymal stem cells combined with hyaluronic acid for the treatment of sheep osteoarthritis,” Cell Transplantation, vol. 27, no. 7, pp. 1111–1125, 2018.

C. H. Jo, Y. G. Lee, W. H. Shin et al., “Intra-articular injection of mesenchymal stem cells for the treatment of osteoarthritis of the knee: a proof-of-concept clinical trial,” Stem Cells, vol. 32, no. 5, pp. 1254–1266, 2014.

N. Yucel and H. M. Blau, “Skeletal muscle stem cells,” in Principles of Regenerative Medicine, Academic Press, 3rd Ed edition, 2019.

B. Biferali, D. Proietti, C. Mozzetta, and L. Madaro, “Fibro-adipogenic progenitors cross-talk in skeletal muscle: the social network,” Frontiers in Physiology, vol. 10, p. 1074, 2019.

R. N. Judson, M. Low, C. Eisner, and F. M. Rossi, “Isolation, culture, and differentiation of fibro/adipogenic progenitors (FAPs) from skeletal muscle,” Methods in Molecular Biology, vol. 1668, pp. 93–103, 2017.

P. A. Zuk, M. I. N. Zhu, H. Mizuno et al., “Multilineage cells from human adipose tissue: implications for cell-based therapies,” Tissue Engineering, vol. 7, no. 2, pp. 211–228, 2001.

Y. H. Lee, A. P. Petkova, and J. G. Granneman, “Identification of an adipogenic niche for adipose tissue remodeling and restoration,” Cell Metabolism, vol. 18, no. 3, pp. 355–367, 2013.

J. Guo, A. Nguyen, D. A. Banyard et al., “Stromal vascular fraction: a regenerative reality? Part 2: mechanisms of regenerative action,” Journal of Plastic, Reconstructive & Aesthetic Surgery, vol. 69, no. 2, pp. 180–188, 2016.

D. A. Upchurch, W. C. Renberg, J. K. Roush, G. A. Milliken, and M. L. Weiss, “Effects of administration of adipose-derived stromal vascular fraction and platelet-rich plasma to dogs with osteoarthritis of the hip joints,” American Journal of Veterinary Research, vol. 77, no. 9, pp. 940–951, 2016.

M. O. Sullivan, W. J. Gordon-Evans, L. P. Fredericks, K. Kiefer, M. G. Conzemius, and D. J. Griffon, “Comparison of mesenchymal stem cell surface markers from bone marrow aspirates and adipose stromal vascular fraction sites,” Frontiers in Veterinary Science, vol. 2, p. 82, 2016.

I. Roato, D. C. Belisario, M. Compagno et al., “Adipose-derived stromal vascular fraction/xenohybrid bone scaffold: an alternative source for bone regeneration,” Stem Cells International, vol. 2018, Article ID 4126379, 11 pages, 2018.

Y. Jang, Y. G. Koh, Y. J. Choi et al., “Characterization of adipose tissue-derived stromal vascular fraction for clinical application to cartilage regeneration,” In Vitro Cellular & Developmental Biology - Animal, vol. 51, no. 2, pp. 142–150, 2015.

M. T. Rodrigues, S. J. Lee, M. E. Gomes, R. L. Reis, A. Atala, and J. J. Yoo, “Amniotic fluid-derived stem cells as a cell source for bone tissue engineering,” Tissue Engineering. Part A, vol. 18, no. 23-24, pp. 2518–2527, 2012.

R. Keeley, N. Topoluk, and J. Mercuri, “Tissues reborn: fetal membrane-derived matrices and stem cells in orthopedic regenerative medicine,” Critical Reviews in Biomedical Engineering, vol. 42, no. 3-4, pp. 249–270, 2014.

Q. Zhou, X. Y. Liu, Y. X. Ruan et al., “Construction of corneal epithelium with human amniotic epithelial cells and repair of limbal deficiency in rabbit models,” Human Cell, vol. 28, no. 1, pp. 22–36, 2015.

M. Magatti, M. Caruso, S. De Munari et al., “Human amniotic membrane-derived, mesenchymal and epithelial cells exert different effects on monocyte-derived dendritic cell differentiation and function,” Cell Transplantation, vol. 24, no. 9, pp. 1733–1752, 2015.

F. Veronesi, G. Giavaresi, M. Tschon, V. Borsari, N. Nicoli Aldini, and M. Fini, “Clinical use of bone marrow, bone marrow concentrate, and expanded bone marrow mesenchymal stem cells in cartilage disease,” Stem Cells and Development, vol. 22, no. 2, pp. 181–192, 2013.

L. Cavinatto, B. B. Hinckel, R. E. Tomlinson, S. Gupta, J. Farr, and A. R. Bartolozzi, “The role of bone marrow aspirate concentrate for the treatment of focal chondral lesions of the knee: a systematic review and critical analysis of animal and clinical studies,” Arthroscopy, vol. 35, no. 6, pp. 1860–1877, 2019.

J. M. Cottom and B. S. Plemmons, “Bone marrow aspirate concentrate and its uses in the foot and ankle,” Clinics in Podiatric Medicine and Surgery, vol. 35, no. 1, pp. 19–26, 2018.

M. A. Imam, J. Holton, S. Horriat et al., “A systematic review of the concept and clinical applications of bone marrow aspirate concentrate in tendon pathology,” SICOT-J, vol. 3, p. 58, 2017.

S. I. Wolbank, A. Peterbauer, M. Fahrner et al., “Dose-dependent immunomodulatory effect of human stem cells from amniotic membrane: a comparison with human mesenchymal stem cells from adipose tissue,” Tissue Engineering, vol. 13, no. 6, pp. 1173–1183, 2007.

J. W. Choi, S. Shin, C. Y. Lee et al., “Rapid induction of osteogenic markers in mesenchymal stem cells by adipose- derived stromal vascular fraction cells,” Cellular Physiology and Biochemistry, vol. 44, no. 1, pp. 53–65, 2018.

Y. Zhang, E. C. Grosfeld, W. A. Camargo, H. Tang, A. M. P. Magri, and J. J. P. van den Beucken, “Efficacy of intraoperatively prepared cell-based constructs for bone regeneration,” Stem Cell Research & Therapy, vol. 9, no. 1, p. 283, 2018.

W. J. Jurgens, R. J. Kroeze, R. A. Bank, M. J. P. F. Ritt, and M. N. Helder, “Rapid attachment of adipose stromal cells on resorbable polymeric scaffolds facilitates the one-step surgical procedure for cartilage and bone tissue engineering purposes,” Journal of Orthopaedic Research, vol. 29, no. 6, pp. 853–860, 2011.

T. Ismail, R. Osinga, T. Todorov Jr. et al., “Engineered, axially-vascularized osteogenic grafts from human adipose-derived cells to treat avascular necrosis of bone in a rat model,” Acta Biomaterialia, vol. 63, pp. 236–245, 2017.

A. M. Müller, A. Mehrkens, D. J. Schäfer et al., “Towards an intraoperative engineering of osteogenic and vasculogenic grafts from the stromal vascular fraction of human adipose tissue,” European Cells & Materials, vol. 19, pp. 127–135, 2010.

A. Todorov, M. Kreutz, A. Haumer et al., “Fat-derived stromal vascular fraction cells enhance the bone-forming capacity of devitalized engineered hypertrophic cartilage matrix,” Stem Cells Translational Medicine, vol. 5, no. 12, pp. 1684–1694, 2016.

S. J. Najman, V. J. Cvetković, J. G. Najdanović et al., “Ectopic osteogenic capacity of freshly isolated adipose-derived stromal vascular fraction cells supported with platelet-rich plasma: a simulation of intraoperative procedure,” Journal of Cranio-Maxillo-Facial Surgery, vol. 44, no. 10, pp. 1750–1760, 2016.

E. Nyberg, A. Farris, A. O'Sullivan, R. Rodriguez, and W. Grayson, “Comparison of stromal vascular fraction and passaged adipose-derived stromal/stem cells as point-of-care agents for bone regeneration,” Tissue Engineering. Part A, vol. 25, no. 21-22, pp. 1459–1469, 2019.

S. C. Rhee, Y. H. Ji, N. A. Gharibjanian, E. S. Dhong, S. H. Park, and E. S. Yoon, “In vivo evaluation of mixtures of uncultured freshly isolated adipose-derived stem cells and demineralized bone matrix for bone regeneration in a rat critically sized calvarial defect model,” Stem Cells and Development, vol. 20, no. 2, pp. 233–242, 2011.

A. Kim, D. H. Kim, H. R. Song et al., “Repair of rabbit ulna segmental bone defect using freshly isolated adipose-derived stromal vascular fraction,” Cytotherapy, vol. 14, no. 3, pp. 296–305, 2012.

S. Kuroshima, M. Sasaki, K. Nakajima, S. Tamaki, H. Hayano, and T. Sawase, “Transplantation of noncultured stromal vascular fraction cells of adipose tissue ameliorates osteonecrosis of the jaw-like lesions in mice,” Journal of Bone and Mineral Research, vol. 33, no. 1, pp. 154–166, 2018.

P. Tyrnenopoulou, M. Karayannopoulou, S. Angelopoulou et al., “Successful management of an equine carpal chip fracture by intra-articularly injected adipose-derived stromal vascular fraction after arthroscopic removal,” Iranian Journal of Veterinary Research, vol. 17, no. 1, pp. 59–61, 2016.

E. Farré-Guasch, N. Bravenboer, M. N. Helder, E. A. J. M. Schulten, C. M. ten Bruggenkate, and J. Klein-Nulend, “Blood vessel formation and bone regeneration potential of the stromal vascular fraction seeded on a calcium phosphate scaffold in the human maxillary sinus floor elevation model,” Materials, vol. 11, no. 1, p. 161, 2018.

H. J. Prins, E. A. J. M. Schulten, C. M. ten Bruggenkate, J. Klein-Nulend, and M. N. Helder, “Bone regeneration using the freshly isolated autologous stromal vascular fraction of adipose tissue in combination with calcium phosphate ceramics,” Stem Cells Translational Medicine, vol. 5, no. 10, pp. 1362–1374, 2016.

F. Saxer, A. Scherberich, A. Todorov et al., “Implantation of stromal vascular fraction progenitors at bone fracture sites: from a rat model to a first-in-man study,” Stem Cells, vol. 34, no. 12, pp. 2956–2966, 2016.

L. Wu, H. J. Prins, J. Leijten et al., “Chondrocytes cocultured with stromal vascular fraction of adipose tissue present more intense chondrogenic characteristics than with adipose stem cells,” Tissue Engineering. Part A, vol. 22, no. 3-4, pp. 336–348, 2016.

W. J. F. M. Jurgens, R. J. Kroeze, B. Zandieh-Doulabi et al., “One-step surgical procedure for the treatment of osteochondral defects with adipose-derived stem cells in a caprine knee defect: a pilot study,” Biores Open Access, vol. 2, no. 4, pp. 315–325, 2013.

P. van Pham, K. Hong-Thien Bui, D. Quoc Ngo, L. Tan Khuat, and N. Kim Phan, “Transplantation of Nonexpanded Adipose Stromal Vascular Fraction and Platelet- Rich Plasma for Articular Cartilage Injury Treatment in Mice Model,” Journal of Medical Engineering, vol. 2013, Article ID 832396, 7 pages, 2013.

N. Yokota, M. Yamakawa, T. Shirata, T. Kimura, and H. Kaneshima, “Clinical results following intra-articular injection of adipose-derived stromal vascular fraction cells in patients with osteoarthritis of the knee,” Regenerative Therapy, vol. 6, pp. 108–112, 2017.

I. Roato, D. C. Belisario, M. Compagno et al., “Concentrated adipose tissue infusion for the treatment of knee osteoarthritis: clinical and histological observations,” International Orthopaedics, vol. 43, no. 1, pp. 15–23, 2019.

H. Bansal, K. Comella, J. Leon et al., “Intra-articular injection in the knee of adipose derived stromal cells (stromal vascular fraction) and platelet rich plasma for osteoarthritis,” Journal of Translational Medicine, vol. 15, no. 1, p. 141, 2017.

N. Gibbs, R. Diamond, E. O. Sekyere, and W. D. Thomas, “Management of knee osteoarthritis by combined stromal vascular fraction cell therapy, platelet-rich plasma, and musculoskeletal exercises: a case series,” Journal of Pain Research, vol. 8, pp. 799–806, 2015.

P. D. Nguyen, T. D.-X. Tran, H. T.-N. Nguyen et al., “Comparative clinical observation of arthroscopic microfracture in the presence and absence of a stromal vascular fraction injection for osteoarthritis,” Stem Cells Translational Medicine, vol. 6, no. 1, pp. 187–195, 2017.

T. D. X. Tran, C. M. Wu, N. K. Dubey et al., “Time- and Kellgren-Lawrence grade-dependent changes in intra-articularly transplanted stromal vascular fraction in osteoarthritic patients,” Cell, vol. 8, no. 4, p. 308, 2019.

Z. Hong, J. Chen, S. Zhang et al., “Intra-articular injection of autologous adipose-derived stromal vascular fractions for knee osteoarthritis: a double-blind randomized self-controlled trial,” International Orthopaedics, vol. 43, no. 5, pp. 1123–1134, 2019.

Y. S. Kim, H. J. Lee, Y. J. Choi, Y. I. Kim, and Y. G. Koh, “Does an injection of a stromal vascular fraction containing adipose-derived mesenchymal stem cells influence the outcomes of marrow stimulation in osteochondral lesions of the talus? A clinical and magnetic resonance imaging study,” The American Journal of Sports Medicine, vol. 42, no. 10, pp. 2424–2434, 2014.

R. Z. Salikhov, R. F. Masgutov, M. A. Chekunov et al., “The stromal vascular fraction from fat tissue in the treatment of osteochondral knee defect: case report,” Frontiers in Medicine, vol. 5, p. 154, 2018.

M. Behfar, F. Sarrafzadeh-Rezaei, R. Hobbenaghi, N. Delirezh, and B. Dalir-Naghadeh, “Adipose-derived stromal vascular fraction improves tendon healing in rabbits,” Chinese Journal of Traumatology, vol. 14, no. 6, pp. 329–335, 2011.

M. Behfar, S. Javanmardi, and F. Sarrafzadeh-Rezaei, “Comparative study on functional effects of allotransplantation of bone marrow stromal cells and adipose derived stromal vascular fraction on tendon repair: a biomechanical study in rabbits,” Cell Journal, vol. 16, no. 3, pp. 263–270, 2014.

L. Y. Lu, M. Ma, J. F. Cai et al., “Effects of local application of adipose-derived stromal vascular fraction on tendon-bone healing after rotator cuff tear in rabbits,” Chinese Medical Journal, vol. 131, no. 21, pp. 2620–2622, 2018.

L. Y. Lu, C. Y. Kuang, and F. Yin, “Magnetic resonance imaging and biomechanical analysis of adipose-derived stromal vascular fraction applied on rotator cuff repair in rabbits,” Chinese Medical Journal, vol. 131, no. 1, pp. 69–74, 2018.

D. Albano, C. Messina, F. G. Usuelli et al., “Magnetic resonance and ultrasound in achilles tendinopathy: predictive role and response assessment to platelet-rich plasma and adipose-derived stromal vascular fraction injection,” European Journal of Radiology, vol. 95, pp. 130–135, 2017.

F. G. Usuelli, M. Grassi, C. Maccario et al., “Intratendinous adipose-derived stromal vascular fraction (SVF) injection provides a safe, efficacious treatment for achilles tendinopathy: results of a randomized controlled clinical trial at a 6-month follow-up,” Knee Surgery, Sports Traumatology, Arthroscopy, vol. 26, no. 7, pp. 2000–2010, 2018.

N. Topoluk, R. Hawkins, J. Tokish, and J. Mercuri, “Amniotic mesenchymal stromal cells exhibit preferential osteogenic and chondrogenic differentiation and enhanced matrix production compared with adipose mesenchymal stromal cells,” The American Journal of Sports Medicine, vol. 45, no. 11, pp. 2637–2646, 2017.

Q. Wang, W. Wu, X. Han et al., “Osteogenic differentiation of amniotic epithelial cells: synergism of pulsed electromagnetic field and biochemical stimuli,” BMC Musculoskeletal Disorders, vol. 15, no. 1, p. 271, 2014.

G. Wang, F. Zhao, D. Yang, J. Wang, L. Qiu, and X. Pang, “Human amniotic epithelial cells regulate osteoblast differentiation through the secretion of TGFβ1 and microRNA-34a-5p,” International Journal of Molecular Medicine, vol. 41, no. 2, pp. 791–799, 2018.

F. Luan, K. Ma, J. Mao, F. Yang, M. Zhang, and H. Luan, “Differentiation of human amniotic epithelial cells into osteoblasts is induced by mechanical stretch via the Wnt/β-catenin signalling pathway,” Molecular Medicine Reports, vol. 18, no. 6, pp. 5717–5725, 2018.

B. Barboni, C. Mangano, L. Valbonetti et al., “Synthetic bone substitute engineered with amniotic epithelial cells enhances bone regeneration after maxillary sinus augmentation,” Plos One, vol. 8, no. 5, article e63256, 2013.

S. Jiawen, Z. Jianjun, D. Jiewen et al., “Osteogenic differentiation of human amniotic epithelial cells and its application in alveolar defect restoration,” Stem Cells Translational Medicine, vol. 3, no. 12, pp. 1504–1513, 2014.

J. Si, J. Dai, J. Zhang et al., “Comparative investigation of human amniotic epithelial cells and mesenchymal stem cells for application in bone tissue engineering,” Stem Cells International, vol. 2015, Article ID 565732, 14 pages, 2015.

E. Muinos-Lopez, T. Hermida-Gomez, I. Fuentes-Boquete, J. de Toro-Santos, F. J. Blanco, and S. M. Dıaz-Prado, “Human amniotic mesenchymal stromal cells as favorable source for cartilage repair,” Tissue Engineering. Part A, vol. 23, no. 17-18, pp. 901–912, 2017.

J. Zhou, G. Yu, C. Cao, J. Pang, and X. Chen, “Bone morphogenetic protein-7 promotes chondrogenesis in human amniotic epithelial cells,” International Orthopaedics, vol. 35, no. 6, pp. 941–948, 2011.

B. Barboni, V. Russo, V. Curini et al., “Achilles tendon regeneration can be improved by amniotic epithelial cell allotransplantation,” Cell Transplantation, vol. 21, no. 11, pp. 2377–2395, 2012.

A. Muttini, L. Valbonetti, M. Abate et al., “Ovine amniotic epithelial cells: In vitro characterization and transplantation into equine superficial digital flexor tendon spontaneous defects,” Research in Veterinary Science, vol. 94, no. 1, pp. 158–169, 2013.

A. Muttini, V. Russo, E. Rossi et al., “Pilot experimental study on amniotic epithelial mesenchymal cell transplantation in natural occurring tendinopathy in horses. Ultrasonographic and histological comparison,” Muscles, Ligaments and Tendons Journal, vol. 5, no. 1, pp. 5–11, 2015.

B. Barboni, V. Curini, V. Russo et al., “Indirect co-culture with tendons or tenocytes can program amniotic epithelial cells towards stepwise tenogenic differentiation,” Plos One, vol. 7, no. 2, article e30974, 2012.

A. Muttini, M. Mattioli, L. Petrizzi et al., “Experimental study on allografts of amniotic epithelial cells in calcaneal tendon lesions of sheep,” Veterinary Research Communications, vol. 34, no. S1, pp. 117–S120, 2010.

A. Mauro, V. Russo, L. di Marcantonio et al., “M1 and M2 macrophage recruitment during tendon regeneration induced by amniotic epithelial cell allotransplantation in ovine,” Research in Veterinary Science, vol. 105, pp. 92–102, 2016.

B. Barboni, V. Russo, V. Gatta et al., “Therapeutic potential of hAECs for early Achilles tendon defect repair through regeneration,” Journal of Tissue Engineering and Regenerative Medicine, vol. 12, no. 3, pp. 1–15, 2017.

J. D. Rees, N. Maffulli, and J. Cook, “Management of tendinopathy,” The American Journal of Sports Medicine, vol. 37, no. 9, pp. 1855–1867, 2009.

J. Vamze, M. Pilmane, and A. Skagers, “Biocompatibility of pure and mixed hydroxyapatite and α-tricalcium phosphate implanted in rabbit bone,” Journal of Materials Science. Materials in Medicine, vol. 26, no. 2, p. 73, 2015.

C. Castellani, G. Vescovo, B. Ravara et al., “The contribution of stem cell therapy to skeletal muscle remodeling in heart failure,” International Journal of Cardiology, vol. 168, no. 3, pp. 2014–2021, 2013.

M. A. Vidal, G. E. Kilroy, M. J. Lopez, J. R. Johnson, R. M. Moore, and J. M. Gimble, “Characterization of equine adipose tissue-derived stromal cells: adipogenic and osteogenic capacity and comparison with bone marrow-derived mesenchymal stromal cells,” Veterinary Surgery, vol. 36, no. 7, pp. 613–622, 2007.

N. J. Palpant, S. Yasuda, O. MacDougald, and J. M. Metzger, “Non-canonical Wnt signaling enhances differentiation of Sca1+/c-kit+ adipose-derived murine stromal vascular cells into spontaneously beating cardiac myocytes,” Journal of Molecular and Cellular Cardiology, vol. 43, no. 3, pp. 362–370, 2007.

D. D. Frisbie, J. D. Kisiday, C. E. Kawcak, N. M. Werpy, and C. W. McIlwraith, “Evaluation of adipose-derived stromal vascular fraction or bone marrow-derived mesenchymal stem cells for treatment of osteoarthritis,” Journal of Orthopaedic Research, vol. 27, no. 12, pp. 1675–1680, 2009.

O. Parolini, F. Alviano, G. P. Bagnara et al., “Concise review: isolation and characterization of cells from human term placenta: outcome of the first international Workshop on Placenta Derived Stem Cells,” Stem Cells, vol. 26, no. 2, pp. 300–311, 2008.