Isolation of osteogenic progenitors from human amniotic fluid using a single step culture protocol
© Antonucci et al; licensee BioMed Central Ltd. 2009
Received: 23 April 2008
Accepted: 16 February 2009
Published: 16 February 2009
Stem cells isolated from amniotic fluid are known to be able to differentiate into different cells types, being thus considered as a potential tool for cellular therapy of different human diseases. In the present study, we report a novel single step protocol for the osteoblastic differentiation of human amniotic fluid cells.
The described protocol is able to provide osteoblastic cells producing nodules of calcium mineralization within 18 days from withdrawal of amniotic fluid samples. These cells display a complete expression of osteogenic markers (COL1, ONC, OPN, OCN, OPG, BSP, Runx2) within 30 days from withdrawal. In order to test the ability of these cells to proliferate on surfaces commonly used in oral osteointegrated implantology, we carried out cultures onto different test disks, namely smooth copper, machined titanium and Sandblasted and Acid Etching titanium (SLA titanium). Electron microscopy analysis evidenced the best cell growth on this latter surface.
The described protocol provides an efficient and time-saving tool for the production of osteogenic cells from amniotic fluid that in the future could be used in oral osteointegrated implantology.
Amniotic Fluid Cells (AFCs) can be classified in epitheloid E-type cells, amniotic fluid specific AF-type cells and fibroblastic F-type cells (1). In recent years, different reports have demonstrated that presence in human amniotic fluid of stem cells (AFS) able to differentiate into multiple lineages [1–8]. Very recently, the ability of clonal AFS to produce cell types inclusive of all embryonic germ layers was demonstrated [9, 10]. Unlike embryonic stem cells, AFS have been showed to be not tumorigenic after transplantation in mice . As a consequence, several studies have suggested the usefulness of these cells for therapeutic purposes [11–16]. Osteoblastic cells derived from AFS could be useful for bone regeneration after traumatic or degenerative damage [17, 18]. In fact, osteoblastic progenitors obtained from amniotic fluid could be used to engineer the craniofacial structures whose natural development is regulated by mesenchymal cells originating from the neural crest, avoiding long and difficult therapies of bone augmentation with intra-oral or extra oral donor site [19, 20]. In order to obtain the best results in craniofacial tissue engineering, great relevance is assumed by the use of scaffolds able to accommodate cell growth and tissue genesis. To date, implants with different surface treatments are investigated to define the best surface morphology for a good osteoblastic cell proliferation and osseointegration around implant [21–25].
The aim of the present study is to evaluate the ability of human AFS to differentiate into osteogenic cells using a novel single step culture protocol, and to test their growth ability on different implant surfaces.
Osteoblastic differentiation was obtained in the present study using two different culture protocols of amniotic fluid cells. In the first protocol (Protocol 1), Amniotic Fluid Mesenchymal Stem cells (AFMSCs) were transferred in osteogenic medium at passage 6, while in the second protocol (Protocol 2), pellets of amniotic fluid samples were directly resuspended in osteogenic medium without the selection of AFMSCs.
In order to evaluate the mitotic stability of cells, cytogenetic investigation was carried out on Protocol 2 cultures at day 30, showing normal diploid karyotype in all the investigated metaphases.
Different protocols have been reported in literature for the differentiation of osteogenic cells starting from amniotic fluid. Some authors reported the use of immunoselection with c-Kit specific antibodies in order to isolate AFS starting from confluent human amniocentesis cultures, followed by proliferation of AFMSCs under appropriate culture conditions, and finally osteoblastic differentiation after several days of culture [8, 9]. On the other hand, other groups cultured unselected amniotic fluid cells in media allowing the proliferation of AFMSCs, and subsequently induced their differentiation in osteoblastic cells [1, 4, 5].
In the present study, we demonstrated the ability of human AFS to differentiate into osteogenic cells using a single step culture procedure, allowing a 20 days reduction of the culture time as compared to previously reported protocols. This could represent an important point in the view of a possible therapeutic application of these cells. Amniotic fluid samples, directly resuspended in osteogenic medium without the selection of AFMSCs, were able to produce osteogenic cells after 18 days from the withdrawal as demonstrated by Alizarin Red staining. RT-PCR analysis showed the full expression of all osteogenic markers typical of late stage osteoblasts after 30 days of colture, while the same expression pattern is showed after 50 days from withdrawal by cells obtained using conventional protocols. Cytogenetic investigation, carried out at day 30 on cells obtained by the single step protocol, showed normal diploid karyotype in all the investigated samples, thus confirming the mitotic stability of cells obtained using this procedure. Cell count performed on cultures carried out with protocol 2 showed the presence at day 30 from withdrawal of about 9,7 × 106 cells starting from as little as 2–3 ml of amniotic fluid. Although the direct culture of AFS cells in osteogenic medium likely induces a complete cell differentiation within 30 days, with arrest of cell proliferation, the amount of cells obtained with this protocol fits well with the cell number required for preclinical studies in animal models and for local transplant in human. Since this latter approach would likely represent the gold standard for a future clinical application in odontoiatric and orthopaedic implantology, the cell number obtained using our direct protocol appears to be sufficient for future local therapeutic purposes.
In order to test the ability of osteoblastic cells obtained from amniotic fluid to proliferate onto surfaces commonly used for craniofacial implantology, and to evaluate their usefulness for tissue engineering, we tested these cells on disks with machined titanium and SLA titanium surfaces. Electron microscopy observation showed a good growth and adherence of osteoblastic cells on this latter surface. This result indicates the excellent biocompatibility of osteoblastic cells obtained from amniotic fluid with SLA titanium scaffolds currently utilized in dental implant.
The protocol described in the present study shows the ability of producing osteoblastic cells from amniotic fluid samples in a very short time, being these cells fully differentiated within one month from withdrawal. Although osteoblastic progenitors can be successfully obtained from bone marrow stromal cells, the use of amniotic fluid as a source of these cells is of relevance since AFS can be easily obtained from routine clinical amniocentesis specimens that would otherwise be discarded. Thus, it is possible to suggest that banking of these stem cells will provide in the future a relevant source both for autologous therapy in the adulthood and for the transplant in HLA matched recipients.
Isolation and culture of mesenchymal stem cells from amniotic fluid (AFMSCs)
Amniotic fluid samples were obtained from 11 women undergoing amniocentesis for prenatal diagnosis at 16–19 weeks of pregnancy after written informed consent. The study has been approved by the Ethics Committee for Biomedical Research of the "G. d'Annunzio" University, Chieti. For each sample, 2–3 ml of amniotic fluid, corresponding to a cell number ranging from 2 × 103 to 2 × 106  were centrifuged for 10 minutes at 1800 rpm. Pellets were resuspended in Iscove's modified Dulbecco's medium supplemented with 20% FBS, 100 U/ml penicillin, 100 μg/ml streptomycin (Sigma), 2 mM L-glutamine, 5 ng/ml basic fibroblast growth factor (FGF2) and incubated at 37°C with 5% humidified CO2. After 7 days, non-adherent cells were removed and the adherent cells allowed to growth in the same medium, which was changed each 4 days. When culture reached confluence (about 20 days after the primary culture), cells were treated with 0,05% trypsin and 0,02% EDTA, then counted and replaced in 25 cm2 culture flasks.
Two different culture protocols were used for osteogenic differentiation of amniotic fluid cells. In the first protocol (Protocol 1), AFMSCs cells at passage 6 were transferred in osteogenic medium consisting of the above described medium with the addition of 150 μg/ml β Glycerophosphate, 50 μg/ml ascorbic acid, and 10-8 M dexamethasone. In the second protocol (Protocol 2), pellets of amniotic fluid samples were directly resuspended in osteogenic medium in 75 cm2 flasks without the selection of AFMSCs. At day 8 from withdrawal, colony forming cells were counted, showing a number ranging from 20 to 20.000 in the different cultures. To visualize calcium sediments, cells treated with Protocol 2 were stained at different times (19, 22 and 30 days from withdrawal) with Alizarin Red S solution, according to Gregory et al. . Mineralization was demonstrated by the presence of red depositions. All reagents used for cells culture and staining were purchased by Sigma-Aldrich (Milano, Italy)
Culture on different surfaces
Three test disks (diameter 10 mm, thickness 5 mm) for each different surface, namely smooth copper, machined titanium and Sandblasted and Acid Etching titanium (SLA titanium), were used in this study Geometric surface morphology of Machined Titanium test disks was obtained with turning machined treatment with formation of titanium micro-parallel walls. Sandblasted and Acid Etching titanium (SLA titanium) test disks were obtained by TiO2 particles being applied to the surface and two phases of etching with fluoridric acid followed by a second acid attack by sulphuric-hydrochloric acid with irregular distributed porosity structure of micro-deep valleys alternated to elevated sharp crests. To preliminary characterize the surface morphology, test disks have been evaluated by means of Scanning Electron Microscopy (SEM) imaging (LEO 435 VP, Cambridge, UK) at about 15–20 kV, high vacuum mode. The surface roughness of the specimens were measured with a stylus profilometer (ANSI/ASME B46.1 1–2002) and a gloss meter (45°-90° sensor angle, 1–10 range, DIN 16537). Differences between treatment groups were evaluated using an analysis of variance at the 95% confidence level and parametric Newman-Keuls multiple comparison test at p = 0.05 significance level. After differentiation, at day 15, osteoblastic cells obtained using protocol 2 were divided in three groups and 3,7 × 104 cells were seeded onto each of the three different test disks. When 70% confluence was observed (after 2–3 days of culture), cells were prepared and analysed by SEM. The entire culture protocol on test disks was repeated two times.
Scanning Electron microscopy
For SEM analysis, specimens cells were fixed in 2% gluteraldehyde in 0.1 M cacodylate buffer (pH 7.4). To preserve the lipid structures, specimens were gently washed in 0.2 M cacodylate buffer (pH 7.4) with the addition of 0.15 M saccharose for three changes every 20 minutes, post-fixed in 1% osmium tetroxide at room temperature for 1 hours, then given two quick changes of the previous buffer and gradually dehydrated in increasing ethanol concentrations (from 25 to 100%, 15% steps). Samples were then carried through critical point drying (CPD) according to standard procedure using liquid carbon dioxide, mounted on aluminium stubs, gold-sputtered and observed with a Philips XL20 Scanning Electron Microscope (SEM Philips XL 20; FEI, Eindhoven, The Netherlands) at 20 KV, high vacuum mode. Images were stored in TIF format with 1024 × 768 Grid of Pixels
Total RNA was isolated using the SV Total RNA Isolation System Kit (Promega, Milano, Italy) from: a) AFMSCs cells after 20 days culture in standard medium (protocol 1); b) differentiated cells after 30 days in osteogenic medium (protocols 1 and 2). RNA from fresh amniotic fluid was also used as a control. One μg of total RNA was reverse transcribed using RETROscript Kit (Ambion, Milano, Italy).
Genes analyzed in RT-PCR experiments, primer sequences and annealing temperature.
Stromal cell-derived factor-1
F – gacccgcgctcgtccgcc
R – cgggtcaatgcacacacttgtcta
Chemokine (C-X-C motif) receptor 4
F – agctgttggctgaaaaggtgg
R – gcgcttctggtggcccttgga
Octamer-binding transcription factor 4
F – cgt gaa gct gga gaa gga gaa gct g
R – caa ggg ccg cag ctc aca cat gtt c
Stem cell factor
F – cca ttg atg cct tca agg ac
R – ctt cca gta taa ggc tcc aa
GATA binding protein 4
F – ttc ctc ttc cct cct caa at
R – tca gcg tgt aaa ggc atc tg
F – tca gcg tgt aaa ggc atc tg
R – cct tcg tga ata cca cg acct gc
Fibroblast growth factor 5
F – gct gtg tct cag ggg att gta gga ata
R – tat cca aag cga aac ttg agt ctg ta
Paired box 6
F – aga ttc aga tga ggc tca aa
R – aat tgg ttg gta gac act gg
Neural cell adhesion molecule
F – gag ggg gaa gat gcc gtg atg tg
R – ata ttc tgc ctg gcc cgg atg gta g
Bone morphogenetic protein 2
F – ttg cgg ctg ctc agc atg tt
R – ttg cga gaa cag atg caa gat g
F – gtg ctg cac ttc ttc ata tgc
R – tga cag cct caa gtt gtt cc
Type I collagen
F – ttcctttgcattcatctctca
R – caagtggaccaagcttcctt
F – gtctcactggctgtgttgga
R – aagacttgccatgtgggttc
F – aggaggaggcagagcaca
R – ctggtatggcacaggtgatg
F – catgagagccctcaca
R – agagcgacaccctagac
F – tgctgttcctacaaagttttacg
R – ctttgagtgctttagtgcgtg
F – ctatggaaggacgccacgcct
R – catagccatcgtagccttgtcc
Runt-related transcription factor 2
F – gacagaagcttgatgactctaaacc
R – tctgtaatctgactctgtccttgt
F – ccatggagaaggctggg
R – caaagttgtcatggatgacc
For cytogenetic analysis, cultures carried out using protocol 2 were treated at day 30 with trypsin and 36–48 hours colcemid. Metaphase chromosomes were stained with GTG-banding and Giemsa. At least 20 metaphases were examined for each sample.
Authors wish to thank Daniela Di Tizio and Tonia di Giovacchino for their technical assistance.
This study has been supported by a grant of the G. d'Annunzio University to LS and GP.
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