Autorzy
Skontaktuj Się z Nami

Zaloguj się

Aby wyświetlić tę treść, wymagana jest subskrypcja JoVE. Zaloguj się lub rozpocznij bezpłatny okres próbny.

W tym Artykule

  • Podsumowanie
  • Streszczenie
  • Wprowadzenie
  • Protokół
  • Wyniki
  • Dyskusje
  • Ujawnienia
  • Podziękowania
  • Materiały
  • Odniesienia
  • Przedruki i uprawnienia

Podsumowanie

The present protocol describes an improved methodology for ADSC isolation resulting in a tremendous cellular yield with time gain compared to the literature. This study also provides a straightforward method for obtaining a relatively large number of viable cells after long-term cryopreservation.

Streszczenie

Human mesenchymal stem cells derived from adipose tissue have become increasingly attractive as they show appropriate features and are an accessible source for regenerative clinical applications. Different protocols have been used to obtain adipose-derived stem cells. This article describes different steps of an improved time-saving protocol to obtain a more significant amount of ADSC, showing how to cryopreserve and thaw ADSC to obtain viable cells for culture expansion. One hundred milliliters of lipoaspirate were collected, using a 26 cm three-hole and 3 mm caliber syringe liposuction, from the abdominal area of nine patients who subsequently underwent elective abdominoplasty. The stem cells isolation was carried out with a series of washes with Dulbecco's Phosphate Buffered Saline (DPBS) solution supplemented with calcium and the use of collagenase. Stromal Vascular Fraction (SVF) cells were cryopreserved, and their viability was checked by immunophenotyping. The SVF cellular yield was 15.7 x 105 cells/mL, ranging between 6.1-26.2 cells/mL. Adherent SVF cells reached confluence after an average of 7.5 (±4.5) days, with an average cellular yield of 12.3 (± 5.7) x 105 cells/mL. The viability of thawed SVF after 8 months, 1 year, and 2 years ranged between 23.06%-72.34% with an average of 47.7% (±24.64) with the lowest viability correlating with cases of two-year freezing. The use of DPBS solution supplemented with calcium and bag resting times for fat precipitation with a shorter time of collagenase digestion resulted in an increased stem cell final cellular yield. The detailed procedure for obtaining high yields of viable stem cells was more efficient regarding time and cellular yield than the techniques from previous studies. Even after a long period of cryopreservation, viable ADSC cells were found in the SVF.

Wprowadzenie

Human mesenchymal stem cells are advantageous in both basic and applied research. The use of this adult cell type overpasses ethical issues-compared to the use of embryonic or other cells-being one of the most promising areas of study in autologous tissue regeneration engineering and cell therapy1, such as the neoplastic area, the treatment of degenerative diseases, and therapeutic applications in the reconstructive surgery area2,3,4,5. It has been previously reported that there is an abundant source of mesenchymal multipotent and pluripotent stem cells in the stromal vascular cell fraction of adipose tissue6,7. These ADSC are considered great candidates for use in cell therapy and transplantation/infusion since a considerable number of cells with a strong capacity for expansion ex vivo can be easily obtained with a high yield from a minimal invasive procedure5,8.

It was also demonstrated that adipose tissue presents a greater capacity to provide mesenchymal stem cells than two other sources (bone marrow and umbilical cord tissue)9. Besides being poorly immunogenic and having a high ability to integrate into the host tissue and to interact with the surrounding tissues4,10, ADSC has a multipotent capacity of differentiation into cell lines, with reports of chondrogenic, osteogenic, and myogenic differentiation under appropriate culture conditions11,12,13, and into cells, such as pancreatic, hepatocytes, and neurogenic cells14,15,16.

The scientific community agrees that the mesenchymal stem cells' immunomodulatory effect is a more relevant mechanism of action for cell therapy17,18,19 than their differentiation property. One of the most significant merits of the ADSC use is the possibility of autologous infusion or grafting, becoming an alternative treatment for several diseases. For regenerative medicine, ADSC have already been used in cases of liver damage, reconstruction of cardiac muscle, regeneration of nervous tissue, improvement of skeletal muscle function, bone regeneration, cancer therapy, and diabetes treatment20,21.

To this date, there are 263 registered clinical trials for the evaluation of ADSC's potential, listed on the website of the United States National Institutes of Health22. Different protocols to harvest adipose tissue have been established, but there is no consensus in the literature about a standardized method to isolate ADSC for clinical use23,24. Lipoaspirate processing methods during and after surgery can directly affect cell viability, the final cellular yield25, and the quality of the ADSC population20. Regarding the surgical pre-treatment, it is not well established which surgical pre-treatment technique yields a more significant number of viable cells after isolation or whether the anesthetic solution injected into adipose tissue affects cell yield and its functions26. Similarly, the difference between techniques for obtaining adipose cells can lead to as much as a 70% decrease in the number of viable ADSC20. According to the literature, mechanical treatments to obtain cell populations with high viability-including ultrasound-should be avoided, for they can break down the adipose tissue20. However, the manual fat aspiration method with syringes is less harmful, causing less cell destruction, with tumescent liposuction yielding a significant number of cells with the best quality26.

This technique uses a saline solution with lidocaine and epinephrine that is injected into the liposuction area. For each 3 mL volume of solution injected, 1 mL is aspirated. In this study, the wet liposuction technique was performed, in which for each 1 mL of adrenaline and saline solution injected, 0.2 mL of adipose tissue is aspirated. The use of digestive enzymes, especially collagenase, is common for the process of isolating ADSC.

After the first isolation step in the laboratory, the final pellet is called stromal vascular fraction (SVF). It contains different cell types27, including endothelial precursor cells, endothelial cells, macrophages, smooth muscle cells, lymphocytes, pericytes, pre-adipocytes, and ADSCs, which are capable of adhesion. Once the final isolation is concluded from in vitro cultures, cells that did not adhere to the plastic are eliminated in medium exchanges. After eight weeks of expansion, medium changes, and passages, ADSCs represent most of the cell population in the flasks20. One of the most significant advantages of using isolated adipose-derived stem cells for a possible future therapy is the possibility of cryopreservation. It was demonstrated that cryopreserved lipoaspirate is a potential source of SVF cells even after 6 weeks of freezing28, with biological activity even after 2 years of cryopreservation29, and full capability to grow and differentiate in culture30. However, during the thawing process, a considerable percentage of cells is usually lost31. Therefore, the lipoaspirate removal process and the following methods of cell isolation must ensure the highest cell yield.

This study describes a faster methodology for collecting and isolating ADSC, demonstrating high cellular yield and viability for better efficiency of cellular therapeutics. Furthermore, the effect of this improved technique after long-term SVF cryopreservation was evaluated.

Protokół

The present study is approved by the Ethics Committee of the UNIFESP (protocol number: 0029/2015 CAAE: 40846215.0.0000.5505), performed after obtaining written informed consent from the patients according to the Declaration of Helsinki (2004). The sample of the present study is composed of nine female patients, aged 33-50 years (average age 41.5) and average initial body mass index (BMI) of 24.54 (ranging between 22.32-26.77) (Table 1) who underwent aesthetic abdominoplasty due to excess of skin after pregnancies, at the Division of Plastic Surgery of the Universidade Federal de São Paulo (UNIFESP), Brazil. To reduce bias, the patients were selected as a homogeneous group considering sex, age, and BMI. The datasets used and/or analyzed during this study are available from the corresponding author upon reasonable request.

1. Collection of lipoaspirate

NOTE: This step needs to be performed in the surgery center.

  1. Use 4% chlorhexidine gluconate (see Table of Materials) for skin preparation and asepsis.
    1. Perform a 2 mm subcutaneous skin incision (between the sub-dermis and aponeurosis). Insert a Klein cannula of 26 mm 3 G three-hole and 3 mm caliber and a syringe to inject a total volume of 500 mL of an adrenaline solution (1 mg/mL) (see Table of Materials) diluted in saline (1:1,000,000) in the infraumbilical area.
  2. Connect a 60 mL syringe to a 26 mm 3 G three-hole and 3 mm caliber liposuction cannula and insert it through the skin incision, locking the plunger to create a vacuum.
    1. Make pushing and pulling movements so that, with the vacuum created, the lipoaspirate remains in the 60 mL syringe.
  3. Using a sterile connector with a valve, transfer the 100 mL of the collected lipoaspirate to a 150 mL polyvinyl chloride transfer bag (see Table of Materials).
    1. Pack the transfer bag in a polystyrene box at room temperature (~25 °C) and take it immediately to the laboratory. Do not take longer than 30 min to start the tissue processing.

2. Processing of lipoaspirate

NOTE: This step is to be performed in the laboratory.

  1. First, weigh the bag, gauge the temperature with a digital non-contact infrared clinical thermometer, and leave the bag resting for 5 min inside the laminar flow chamber for precipitation of the greasier layers (bubbles) and tissue separation containing the cells of interest.
    1. Perform a series of tissue washes. First wash: inject 100 mL of DPBS with calcium (1x) into the transfer bag and mix it with the hands.
    2. Let it stand for 5 min and remove most of the basal liquid that precipitates.
    3. Discard the basal liquid with a 60 mL syringe attached to the bag adapter. This process must be repeated twice.
  2. Add 100 mL of digestion solution to the bag (93 mL of calcium-free DPBS + 60 µL of calcium chloride (1 g/L) + 7 mL of 0.075% sterile collagenase, see Table of Materials) and leave at 37 °C for 30 min under slow stirring.
  3. Transfer all the bag's content to four conical tubes of 50 mL and centrifuge them at 400 x g at 22 °C for 10 min.
    1. Remove and discard the supernatant and add 5 mL of Dulbecco's modified Eagle's medium (DMEM) low glucose supplemented with 20% FBS (Fetal bovine serum) to the cell pellet (Figure 1).

3. Counting of the SVF cells

  1. Mix a fresh solution of 10 µL of trypan blue at 0.05% in distilled water with 10 µL of cellular suspension for 5 min.
  2. Count viable cells in a Neubauer cell counting chamber32 using an inverted light microscope (see Table of Materials) at 20x magnification.
  3. Resuspend the cell pellet in a cryoprotective medium (5 mL of FBS + 10% of Dimethyl Sulfoxide - DMSO) at a concentration of 1 x 106 cells/mL.
  4. Place 1 mL of this mix in cryovials. Use a freezing container (see Table of Materials) with a cooling rate of (1 °C/min to -80 °C).
    1. Store at -80 °C for 1 year.
    2. After this time, store in standard cassette boxes immersed in the liquid nitrogen vapor phase (-165 °C).

4. Thawing process of the cells

  1. Remove the vials from liquid nitrogen and place them immediately in the 37 °C water bath for 1 min.
  2. Place the SVF cells in a conical tube with 4 mL of DMEM (low glucose supplemented with 20% FBS) preheated at 37 °C.
  3. Centrifuge at 400 x g at 22 °C for 5 min.
  4. Remove the supernatant and add 1 mL of DMEM (low glucose) + 10% FBS. Perform immunophenotyping following the steps below.

5. Flow cytometry technique (immunophenotype multiple labeling)

  1. Place 1 mL of cell pellet (concentration of 1,000 cells/µL) in five cytometry tubes (200 µL each).
  2. Centrifuge at 400 x g at 22 °C for 5 min and discard the supernatant with a pipette.
  3. Add 300 µL of Phosphate-Buffered Saline (PBS) (10x), centrifuge at 400 x g at 22 °C and discard the supernatant with a pipette.
  4. Prepare five tubes for different marker combinations as follows: 5 µL of CD11B/5 µL of CD19/20 µL of CD45; 5 µL of CD73/20 µL of CD90/5 µL of CD105/20 µL of CD45; 20 µL of CD34/5 µL of HLA-DR/20 µL of CD45; Cell viability assay-5 µL of fluorescent reactive dye. (see Table of Materials) and a tube with unstained cells and PBS as the negative control. Homogenize in a vortex and incubate at 4 °C for 30 min.
    1. Centrifuge at 400 x g at 22 °C for 5 min, discard the supernatant with a pipette, add 500 µL of PBS (10x), and proceed with cell sorting.
      ​NOTE: Five thousand events are acquired per antibody set in the Flow Cytometer of four colors and five parameters and analyzed with CellQuest software.

6. Seeding of passage 1 (P1)

  1. Seed 2 x 105 cells in a 75 cm2 culture flask.
  2. Add 12 mL of DMEM low glucose + 20% of FBS + 10% antibiotic/antimycotic (with 10,000 units penicillin, 10 mg streptomycin, and 25 µg amphotericin B per mL, 0.1 µm).
  3. When the cells reach between 80%-90% confluence, perform trypsinization of adherent cells with 2 mL of 0.25% EDTA-trypsin for 3 min.
  4. Count cells again (as mentioned in step 3).
  5. Perform immunophenotyping again (as mentioned in step 5).

7. Statistical analysis

  1. Use Spearman's Rho Calculator33 to measure the strength of association between the following variables with P < 0.05, as mentioned below.
    1. Select SVF cellular yield and the number of days SVF stays in culture in the first passage (P1) until 80%-90% confluence (days to P1).
    2. Select SVF cellular yield before and after going to P1.
    3. Consider days to P1 and cellular yield before going to P1.
    4. Select SVF cellular yield with the average percentage of confirmed ADSC.
    5. Calculate the percentage of confirmed ADSC and the cellular yield before going to P1.
    6. Determine the BMI and SVF cellular yield.

8. Differentiation assay

  1. Perform the differentiation assay following a differentiation kit protocol (see Table of materials). Figure 4 demonstrates the results for Case 1.

Wyniki

The characterization of the nine individuals studied, including their age, weight, height, and BMI, are shown in Table 1.

According to the cellular yield initially presented, the cell volume inoculated in culture was calculated to be as close as possible to the capacity of the 75 cm2 culture flask. The sample volume seeded in each case is described in Table 2. Then, according to the initial cellular yield, a variable volume of cells for each sample ...

Dyskusje

Isolation yield
It is well established that the cryopreservation process, frequently required in cellular therapy, results in significant cell loss, sometimes greater than 50%29,30,35. Thus, a technical improvement for obtaining high initial cellular yield in isolation is fundamental. The collecting method of lipoaspirate and the isolation method of the cells must focus on preserving a greater number of ce...

Ujawnienia

The authors declare no competing financial interests.

Podziękowania

We thank the patients who volunteered to participate and the medical and nursing staff of the Hospital São Paulo. This study was supported by the Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq), Brazil.

Materiały

NameCompanyCatalog NumberComments
1.8 mL cryovialsNunc Thermo Fisher Scientific340711
150 mL polyvinyl chloride transfer bagJP FARMA80146150059
2% Alizarin Red S Solution, pH 4.2Sigma AldrichA5533
Adrenaline (1 mg/mL)HipolaborNA
Alcian Blue solutionSigma Aldrich1,01,647
Antibiotic-Antimycotic 100xGibco15240062
BD FACSCalibur Flow Cytometer using BD CellQues Pro AnalysisBD BioSciencesNA
Calcium chloride 10%Merck102379
Chlorhexidine gluconate 4%VIC PHARMANA
Collagenase, Type I, powderGibco17018029
DMEM (Dulbecco's modified Eagle's medium)Gibco11966025
DPBS no calcium, no magnesium (Dulbecco's Phosphate Buffered Saline Gibco Cell Therapy Systems)GibcoA1285801
DPBS with calcium (Dulbecco's Phosphate Buffered Saline Gibco Cell Therapy Systems)GibcoA1285601
Fetal bovine serumGibco10500056
Formaldehyde 4%Sigma Aldrich1,00,496
Inverted light microscopeNikon Eclipse TS100NA
Live and Dead Cell AssayThermofisher01-3333-41 | 01-3333-42
Monoclonal antibody: CD105BD BioSciences745927
Monoclonal antibody: CD11BBD BioSciences746004
Monoclonal antibody: CD19BD BioSciences745907
Monoclonal antibody: CD34BD BioSciences747822
Monoclonal antibody: CD45DAKOM0701
Monoclonal antibody: CD73BD BioSciences746000
Monoclonal antibody: CD90BD BioSciences553011
Monoclonal antibody: HLA-DRBD BioSciences340827
Mr. Frosty Freezing ContainerThermo Fisher Scientific5100-0001
PBS (phosphate buffered saline) 1x pH 7.4Gibco 10010023
StemPro Adipogenesis Differentiation KitGibcoA1007001
StemPro Chondrogenesis Differentiation KitGibcoA1007101
StemPro Osteogenesis Differentiation KitGibcoA1007201
Sterile connector with one spike with needle injection siteOrigen Biomedical Connector, USANACode mark: IBS
Trypan blue solution 0.4%Sigma Aldrich93595
Trypsin-EDTA 0.25% 1x, phenol redGibco25200056

Odniesienia

  1. Frese, L., Dijkman, P. E., Hoerstrup, S. P. Adipose tissue-derived stem cells in regenerative medicine. Transfusion Medicina and Hemotherapy. 43 (4), 268-274 (2016).
  2. Alperovich, M., et al. Adipose stem cell therapy in cancer reconstruction: a critical review. Annals of Plastic Surgery. 73, 104-107 (2014).
  3. Forcales, S. V. Potential of adipose-derived stem cells in muscular regenerative therapies. Frontiers in Aging Neuroscience. 7, 123 (2015).
  4. Bateman, M. E., Strong, A. L., Gimble, J. M., Bunnell, B. A. Concise review: using fat to fight disease: a systematic review of nonhomologous adipose-derived stromal/stem cell therapies. Stem Cells. 36 (9), 1311-1328 (2018).
  5. Gentile, P., Cervelli, V. Adipose-derived stromal vascular fraction cells and platelet- rich plasma: basic and clinical implications for tissue engineering therapies in regenerative surgery. Methods in Molecular Biology. 1773, 107-122 (2018).
  6. Schäffler, A., Büchler, C. Concise review: adipose tissue-derived stromal cells- basic and clinical implications for novel cell-based therapies. Stem Cells. 25 (4), 818-827 (2007).
  7. Witkowska-Zimny, M., Walenko, K. Stem cells from adipose tissue. Cellular & Molecular Biology Letters. 16, 236-257 (2011).
  8. Liew, L. J., Ong, H. T., Dilley, R. J. Isolation and culture of adipose-derived stromal cells from subcutaneous fat. Methods in Molecular Biology. 1627, 193-203 (2017).
  9. Fazzina, R., et al. Potency testing of mesenchymal stromal cell growth expanded in human platelet lysate from different human tissues. Stem Cell Research & Therapy. 7 (1), 122 (2016).
  10. Yarak, S., Okamoto, O. K. Human adipose-derived stem cells: Current challenges and clinical perspectives. Brazilian Annals of Dermatology. 85 (5), 647-656 (2010).
  11. Mizuno, H. Adipose-derived stem cells for tissue repair and regeneration: ten years of research and a literature review. Journal of Nippon Medical School. 76 (2), 56-66 (2009).
  12. Makarov, A. V., Arutyunyan, I. V., Bol'Shakova, G. B., Volkov, A. V., Gol'Dshtein, D. V. Morphological changes in paraurethral area after introduction of tissue engineering constructo on the basis of adipose tissue stromal cells. Bulletin of Experimental Biology and Medicine. 148 (4), 719-724 (2009).
  13. Simonacci, F., Bertozzi, N., Raposio, E. Off-label use of adipose-derived stem cells. Annals of Medicina and Surgery. 24, 44-51 (2017).
  14. Scanarotti, C., et al. Neurogenic-committed human pre-adipocytes express CYP1A isoforms. Chemico-Biological Interactions. 184 (3), 474-483 (2010).
  15. Aluigi, M. G., et al. Pre-adipocytes commitment to neurogenesis 1: Preliminary localisation of cholinergic molecules. Cell Biology International. 33 (5), 594-601 (2009).
  16. Coradeghini, R., et al. A comparative study of proliferation and hepatic differentiation of human adipose-derived stem cells. Cells Tissues Organs. 191 (6), 466-477 (2010).
  17. Xishan, Z., et al. Jagged-2 enhances immunomodulatory activity in adipose derived mesenchymal stem cells. Scientific Reports. 5, 14284 (2015).
  18. Debnath, T., Chelluri, L. K. Standardization and quality assessment for clinical grade mesenchymal stem cells from human adipose tissue. Hematology, Transfusion and Cell Therapy. 41 (1), 7-16 (2019).
  19. El-Sayed, M., et al. Immunomodulatory effect of mesenchymal stem cells: Cell origin and cell quality variations. Molecular Biology Reports. 46 (1), 1157-1165 (2019).
  20. Harasymiak-Krzyzanowska, I., et al. Adipose tissue-derived stem cells show considerable promise for regenerative medicine applications. Cellular & Molecular Biology Letters. 18 (4), 479-493 (2013).
  21. Lindroos, B., Suuronen, R., Miettinen, S. The potential of adipose stem cells in regenerative medicine. Stem Cell Reviews and Reports. 7 (2), 269-291 (2011).
  22. . US National Institutes of Health Website Available from: https://clinicaltrials.gov/ (2019)
  23. Baer, P. C., Geiger, H. Adipose-derived mesenchymal stromal/stem cells: Tissue localization, characterization, and heterogeneity. Stem Cells International. 2012, 812693 (2012).
  24. Raposio, E., Bertozzi, N. Isolation of ready-to-use adipose-derived stem cell (ASC) pellet for clinical applications and a comparative overview of alternate methods for ASC isolation. Current Protocols in Stem Cell Biology. 41, 1-12 (2017).
  25. Cucchiani, R., Corrales, L. The effects of fat harvesting and preparation, air exposure, obesity, and stem cell enrichment on adipocyte viability prior to graft transplantation. Aesthetic Surgery Journal. 36 (10), 1164-1173 (2016).
  26. Muscari, C., et al. Comparison between stem cells harvested from wet and dry lipoaspirates. Connective Tissue Research. 54 (1), 34-40 (2013).
  27. Bora, P., Majumdar, A. S. Adipose tissue-derived stromal vascular fraction in regenerative medicine: a brief review on biology and translation. Stem Cell Research & Therapy. 8 (1), 145 (2017).
  28. Zanata, F., et al. Effect of cryopreservation on human adipose tissue and isolated stromal vascular fraction cells: in vitro and in vivo analyses. Plastic and Reconstructive Surgery. 141 (2), (2018).
  29. Harris, D. Long-term frozen storage of stem cells: challenges and solutions. Journal of Biorepository Science for Applied Medicina. 4, 9-20 (2016).
  30. Minonzio, G., et al. Frozen adipose-derived mesenchymal stem cells maintain high capability to grow and differentiate. Cryobiology. 69 (2), 211-216 (2014).
  31. Devitt, S. M., et al. Successful isolation of viable adipose-derived stem cells from human adipose tissue subject to long-term cryopreservation positive implications for adult stem cell-based therapeutics in patients of advanced age. Stem Cells International. 2015, 146421 (2015).
  32. Freshney, R. I. . Culture of animal cells: a manual of basic technique. 3rd Ed. , (2001).
  33. . Spearman's Rho Calculator Available from: https://www.socscistatistics.com/tests/spearman/ (2020)
  34. Dominici, M., et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy. 8 (4), 315-317 (2006).
  35. Irioda, A. C., et al. Human adipose-derived mesenchymal stem cells cryopreservation and thawing decrease α4-integrin expression. Stem Cells International. 2016, 2562718 (2016).
  36. Aust, L., et al. Yield of human adipose-derived adult stem cells from liposuction aspirates. Cytotherapy. 6 (1), 7-14 (2004).
  37. Pérez, L. M., et al. Altered metabolic and stemness capacity of adipose tissue-derived stem cells from obese mouse and human. PLoS One. 10 (4), 0123397 (2015).
  38. Yoshimura, K., et al. Characterization of freshly isolated and cultured cells derived from the fatty and fluid portions of liposuction aspirates. Journal of Cellular Physiology. 208 (1), 64-76 (2006).
  39. Mojallal, A., et al. Influence of age and body mass index on the yield and proliferation capacity of adipose-derived stem cells. Aesthetic Plastic Surgery. 35 (6), 1097-1105 (2011).
  40. Tevlin, R., et al. A novel method of human adipose-derived stem cell isolation with resultant increased cell yield. Plastic and Reconstructive Surgery. 138 (6), (2016).
  41. Tsekouras, A., et al. Comparison of the viability and yield of adipose-derived stem cells (ASCs) from different donor areas. In Vivo. 31 (6), 1229-1234 (2017).
  42. Varghese, J., Griffin, M., Mosahebi, A., Butler, P. Systematic review of patient factors affecting adipose stem cell viability and function: implications for regenerative therapy. Stem Cell Research & Therapy. 8 (1), 45 (2017).
  43. Mitchell, J. B., et al. Immunophenotype of human adipose-derived cells: temporal changes in stromal-associated and stem cell-associated markers. Stem Cells. 24 (2), 376-385 (2006).
  44. Huss, R. Perspectives on the morphology and biology of CD34-negative stem cells. Journal of Hematotherapy and Stem Cell Research. 9 (6), 783-793 (2000).
  45. Bourin, P., et al. Stromal cells from the adipose tissue-derived stromal vascular fraction and culture expanded adipose tissue-derived stromal/stem cells: a joint statement of the International Federation for Adipose Therapeutics (IFATS) and Science and the International Society for Cellular Therapy (ISCT). Cytotherapy. 15 (6), 641-648 (2013).
  46. Scherberich, A., Di Maggio, N., Mcnagny, K. M. A familiar stranger: CD34 expression and putative functions in SVF cells of adipose tissue. World Journal of Stem Cells. 5 (1), 1-8 (2013).
  47. Yu, G., et al. Yield and characterization of subcutaneous human adipose-derived stem cells by flow cytometric and adipogenic mRNA analyzes. Cytotherapy. 12 (4), 538-546 (2010).
  48. Meyerrose, T. E., et al. In vivo distribution of human adipose-derived mesenchymal stem cells in novel xenotransplantation models. Stem Cells. 25 (1), 220-227 (2007).
  49. Pilgaard, L., Lund, P., Rasmussen, J. G., Fink, T., Zachar, V. Comparative analysis of highly defined proteases for the isolation of adipose tissue-derived stem cells. Regenerative Medicina. 3 (5), 705-715 (2008).
  50. Browne, S. M., Al-Rubeai, M. Defining viability in mammalian cell cultures Defining viability in mammalian cell cultures. Biotechnolology Letters. 33, 1745-1749 (2011).
  51. Pogozhykh, D., et al. Influence of temperature fluctuations during cryopreservation on vital parameters, differentiation potential, and transgene expression of placental multipotent stromal cells. Stem Cell Research & Therapy. 8 (1), 66 (2017).
  52. Shaik, S., Wu, X., Gimble, J., Devireddy, R. Effects of decade long freezing storage on adipose derived stem cells functionality. Scientific Reports. 8 (1), 8162 (2018).
  53. Weissbein, U., Benvenisty, N., Ben-David, U. Genome maintenance in pluripotent stem cells. Journal of Cell Biology. 204 (2), 153-163 (2014).
  54. Francis, M. P., Sachs, P. C., Elmore, L. W., Holt, S. E. Isolating adipose-derived mesenchymal stem cells from lipoaspirate blood and saline fraction. Organogenesis. 6 (1), 11-14 (2010).
  55. Palumbo, P., et al. Methods of isolation, characterization and expansion of human Adipose-Derived Stem Cells (ASCs): An overview. International Journal of Molecular Sciences. 19 (7), 1897 (2018).

Przedruki i uprawnienia

Zapytaj o uprawnienia na użycie tekstu lub obrazów z tego artykułu JoVE

Zapytaj o uprawnienia

Przeglądaj więcej artyków

Mesenchymal Stem CellsAdipose TissueStromal Vascular FractionCryopreservationCell IsolationPluripotent CellsCytogenomic Stability TestingDrug TestingLaminar Flow ChamberDPBSDigestion SolutionCentrifugationDMEMFetal Bovine SerumTrypan BlueNeubauer Counting ChamberCryo protective Medium

This article has been published

Video Coming Soon

JoVE Logo

Prywatność

Warunki Korzystania

Zasady

Skontaktuj Się z Nami

Poleć Bibliotece

BIULETYNY JoVE

Badania

Edukacja

Autorzy

Bibliotekarze

O JoVE

Copyright © 2025 MyJoVE Corporation. Wszelkie prawa zastrzeżone