Isolation of Stem Cells from Fatty Tissue
as SVF or
Adipose-Derived Stem Cells (ADSCs)

Gimble et al. have suggested that an ideal stem cell source for clinical applications should meet the following requirements:

  • Can be found in abundant quantities (millions to billions of cells)
  • Can be harvested by a minimally invasive procedure
  • Can be differentiated along multilineage pathways in a reproducible manner
  • Can be safely and effectively transplanted to either autologous or allogeneic host
  • Can be manufactured in accordance with the current Good Manufacturing Practices (cGMP).

Adipose tissue can fulfill all these criteria, since it is abundant and can be easily obtained in large quantities to isolate adequate numbers of adipose-derived stem cells (ASCs) with little donor site morbidity and patient discomfort. In vitro and in vivo, ASCs can differentiate along multiple pathways, including osteogenic, adipogenic, chondrogenic, endothelial, epithelial, hepatic, myogenic, and neuronal-like lineages. They can also act through the paracrine mechanism of releasing growth factors to accelerate tissue repair and regeneration. Furthermore, large-scale production of ASCs required for cellular therapies can be achieved under cGMP condition, and ASCs’ application has been proven safe and efficacious in preclinical and clinical studies.

History of Isolation of Stem Cells from Fatty Tissue

It is very short history, started just in 2001, by Zuk et al.  were the first to isolate Adult Mesenchymal Stem Cells from adipose tissue after a liposuction procedure by means of existing enzymatic strategies. Since then, interest in ADSCs has grown dramatically & several groups working independently have developed procedures to isolate and characterize them. However, it has emerged that, depending on the isolation process, 1 g of adipose tissue can yield an inconsistent number of ADSCs. Mizuno et al. (2009) reported a yield of approximately 5 x 103 stem cells for each gram of adipose tissue, which is 500-fold greater than the number of BmSCs in 1 g of bone marrow; Bouquest et al. (2006) reported a yield of 1 x 107 ADSCs from 300 mL of lipoaspirate. Francis et al. (2010) described a rapid collagenase-free isolation protocol with a yield of  2.5 x 105 ASCs starting from 250 mL of lipoaspirate. Other studies have demonstrated that 1 g of adipose tissue can yield 2 x 106 SVF cells, with 510% more or less of these cells thought to be ADSCs.

About the Fatty Tissue (Adipose Tissue)

Adipocytes constitute almost 90% of adipose tissue volume and nearly 65% of the total cell number. When enzymatically digested, adipose tissue yields a heterogeneous population of many cell types (preadipocytes, fibroblasts, vascular smooth muscle cells, endothelial cells, resident monocytes/macrophages, and lymphocytes), which upon isolation are termed the Stromal Vascular Fraction (SVF). The SVF includes ADSCs, which account for up to 30% of the total SVF cells.

Adipocytes constitute almost 90% of adipose tissue volume & nearly 65% of the total cell number 

Ideal Methods
to Separate the “Stem Cells” from Adipose Tissue

In general, the most efficient methods that can isolate about 500,000–1,000,000 cells per gram of lipoaspirate tissue with a >80 % viability.

The number of viable cells required for treatment of a particular condition is unknown because there is insufficient data to establish a reliable dose vs effect relationship.

Technical Factors Influencing ADSCs Characteristics:

Various factors need to be taken into consideration for optimal stem cells from fatty tissue (ADSCs), these include;

  1. The anatomical location of the donor and the recipient
  2. The donor’s age and gender
  3. The tissue-harvesting technique
  4. Cell isolation procedures
  5. Maintenance of cells under good tissue manufacturing practices.

 Adipose Tissue Type & Anatomical Location:

The aim of the tissue engineering scientist is to identify the variation of stem characteristic according to the harvesting areas, which could help to identify best site of stem cells collection according to structural and functional need. Their findings highlight the importance of the specific anatomical location as a source of stem cells.

  • Stem cells (ADSCs) isolated from subcutaneous adipose tissue may be more suited to differentiate into mature adipocytes (fat cells) than visceral or omental tissue (Intra-abdominal fat).
  • No significant differences in terms of the amount or viability of stem cells collected from fat of the abdomen, hip or thigh donor areas was observed according to (Oedayrajsingh-Varma 2006).

In contrast, (Jurgens et al 2008, Iyyanki et al. 2015) found that the amount of stem cells collected from fat is dependent on the specific fatty tissue-harvesting site. The abdominal area yielded significantly more ADSCs when compared to the hip and thigh regions although no differences in the proliferation & differentiation capacity (function).

  • Upper layer of fatty tissue (superficial layer) give higher amount of stem cells with increase the  multipotency and differentiation capacity.

 Patient Age & Gender:

Age & gender are also important factors to consider when isolating ADSCs from adipose tissue. The younger patients demonstrated significantly higher cell proliferation rates & higher lipolysis activity. There is still controversy with regards to what causes aging of mesenchymal stem cells MSCs, whether it is related to intrinsic or extrinsic factors, possibility, both.

 Harvesting Techniques:

The process starts with fat collection which is technically is a simple and safe procedure, fat can be harvested in an easy outpatient session. The fat excision or liposuction of fat has minimal ethical considerations and limited pain and discomfort to the patient and demonstrated an increased ADSCs yield and viability.

Types of Fat Harvesting Techniques

Although there is unanimous agreement on the harvesting procedure for adipose tissue, there are many methods to get the fat from patient’s body plus  various protocols for adipose-derived stem cell isolation:

Liposuction Versus Biopsy / Resection:

A recent study compared ADSCs obtained through liposuction and the one from fat excision & clearly demonstrated that adipose tissue extraction by liposuction does not damage the stem cells & they went further and suggested that liposuction is the better method for harvesting ADSCs due to increased proliferation and differentiation capacity of ADSCs obtained from liposuction, interfering that liposuction produces a more homogenous population of stem cells than the ADSCs obtained from resection of fatty tissue. Interestingly, although the ADSCs obtained from liposuction and resection were collected from the same patient at the same donor site, differences in gene expression profile were observed, (Gnanasegaran et al. 2014)

Dry Liposuction Versus Wet Technique:

Local anaesthetics have a marked influence on the quality and quantity of viable preadipocytes and ADSCs. All local anaesthetic-exposed ADSCs demonstrated impaired adipocyte differentiation as determined by adiponectin expression.

Effect of Different Harvesting Technique on the
Quality of ADSCs

Most plastic surgeon world-wide currently in practice, with experience in different types of liposuction techniques.

  • It was confirmed that laser-assisted liposuction (LAL), negatively impacts the biology of ADSCs and is therefore NOT preferable for tissue engineering purposes.
  • Adipose tissue harvested by both biopsy (fat excision) and wet liposuction provided high yields of rapidly growing ADSCs, whereas
  • Adipose tissue obtained by Ultrasonic-Assisted Liposuction (UAL) provided a low yield of ADSCs exhibiting a low proliferative capacity.
  • Recently it was shown that water-jet assisted liposuction (WAL) yields more viable ASCs in the SVF compared to conventional liposuction SAL & ultrasonic-assisted liposuction UAL, but less viable ADSCs compared to the Coleman harvesting technique (syringe technique) (Myer et al. 2015).
  • Coleman harvesting technique (syringe technique) using 10,20,60 ml syringes were the negative pressure of; 275, 394, & 549 mmHg, respectively, and no significant differences in ADSCs integrity and viability were observed (Charles-de-Sa et al. 2015)

Coleman Technique: a Coleman blunt tip harvesting cannula with outer diameter of 4 mm, and inner diameter of 2.5 mm. The harvesting cannula is inserted through a puncture wound into the subcutaneous adipose layer of the abdominal door site. The plunger of the syringe is withdrawn 1-3 ml at a time to create the low negative pressure vacuum within the barrel of the harvesting cannula and syringe. The negative pressure decreases within the suction system as the barrel of the syringe fills with adipose tissue. The plunger is then drawn again creating vacuum to allow more adipose tissue to be suctioned. As the syringe fills with lipoaspirate, the negative pressure decreases until the suction vacuum is insufficient to allow for further harvesting of adipose tissue.

This method is successful in obtaining large volume of ADSCs from the SVF & does not negatively affect cellular expansion and differentiation.   

Using Coleman dry needle aspiration technique with an average of 1,019,129 cells/ ml. based on what is reported in the literature the above mentioned harvesting technique to be the preferred technique for isolation of adipose derived SVF cultured ADSCs.

 Cell isolation procedures

The number of ADSCs that  witll be isoalted from lipoaspirate tissue depend on which protocol of isolation & preparation that we are going to use.    

1. Non-expanded, Ready-to-Use Pellet

"Crude (SVF)"

Lipoaspirate is washed 2–3 times using a sterile  aqueous salt solution such as PBS, Lactated Ringer’s solution, or Hank’s Balanced Salt Solution (HBSS) to to remove blood cells, saline and local. The isolation is by one of 2 ways, depend on the case demand & availability where the treatment as the legislation is different from one country to other. 

Enzymatic protocol; the washed lipoaspirate is then incubated with a collagenase Solution of variable concentration & composition, depending on the method and tissue dissociation enzyme product used.

Mechanical protocol; alternative non-enzymatic means of removing SVF cells from the adipose tissue & tend to be focused around washing and shaking/vibrating lipoaspirate followed by centrifugation in order to concentrate the SVF cells. SVF isolation report from mechanical isolation protocol is significantly lower yields of nucleated cells/cc of lipoaspirate processed. Cell yields are reported from 10,000 nucleated cells/cc of lipoaspirate to 240,000 nucleated cells/cc of lipoaspirate

The outcome of the process is called Stromal Vascular fraction (SVF). SVF is a complex fraction that contain ADSCs (5%-10% more or less) but cannot be defined as stem cells or Mesenchymal Stem Cells MSCs fraction. It is highly heterogenous and contain many cells types including; mature adipocytes, a mixture of endothelial cells, smooth cells, fibroblasts, pericytes, and reservoir of immature stromal cells, & hematopoietic cells (up to 20% of total fraction).

SVF is more effective if the extracted adipose tissue treated by collagenase digestion to obtain the ADSCs which can be 1000 times more effective than the SVF collected by mechanical methods.. But, the gathered data by Rasposio el al, published 2017 showing a total of 100 ml of lipoaspirate allowed to isolate a mean of 9.06 x 105 , a greater amount (25.9%) of ADSCs by enzymatic protocol compared to a mean of 6.25 x 105 , (5%) of ASCs by mechanical protocol

Despite the enzymatic procedure was superior as regards the number of ASCs isolated, the time required for the entire isolation process was greater (80 min) compared to the mechanical protocol (15 min). The use of collagenase as an injectable pharmaceutical is associated with the risk of serious side effects, such as cutaneous ulcers, nerve injury, tendon or ligament damage, and allergic reactions. However, No study has been carried out reporting the residual collagenase activity in adipose-derived stem cell samples inoculated in humans.

History of Ready-to-Use SVF Preparationd
"Enzymatic Method"

The isolation process described by Zuk et al. has come to be the most commonly published method for ADSC isolation. Briefly, the freshly harvested lipoaspirate is washed with sterile phosphate buffered solution to remove blood cells, saline and local anaesthetics. Then, it is enzymatically digested using 0.075% collagenase type I, which subsequently must be inactivated by means of addition of an equal volume of Dulbecco’s modified Eagle’s medium containing 10% fetal bovine serum. Finally, red blood cell lysis is performed, and the pellet is separated from the infranatant by centrifugation. Although the procedure has proven effective, it can be complex, expensive & time-consuming for clinical application, since the entire process takes an average of 2 h.

In 2016, Raposio et al. described a method that was specifically designed for clinical application, which appeared easy, safe & fast (80 min), allowing collection of a Ready-to-Use SVF pellet. Just like previous isolation processes, first the adipose tissue was harvested by a standard liposuction procedure, & then the isolation process was carried out through both mechanical (centrifugation) & enzymatic (collagenase) means. The entire process was performed in a closed-circuit system, which guaranteed sterility and the safety. Once an SVF pellet was obtained in this manner, Raposio and his colleagues were able to inject it into the skin at the wound edges, as well as at the bottom of chronic skin ulcers, to promote wound-healing processes. Based on a flow cytometric assay, 9.06 x 105 (SD ± 6.6 x 105) ADSCs were isolated from 100 mL of adipose tissue. ADSCs accounted for 25.9% of the pelleted cells, which is a considerably higher yield compared with those reported by Mizuno (2012).

History of ready-to-use SVF Pellet by
"Mechanical Method"

Several alternative isolation methods have been proposed which avoid enzymatic digestion completely, in keeping with the findings of Yoshimura & co-workers as they have found that “a significant number of ADSCs could be isolated from what they called the liposuction aspirate fluid, and although cell numbers were less than those obtained with enzymatic digestionthey were  still enough to be used clinically without cell expansion”. In the light of these findings, Francis et al. (2010) isolated cells capable of trilineage differentiation by mechanically separating the SVF directly from the lipoaspirate. Soon after, Bianchi et al. (2013) described a non-expanded, ready-to-use fat product obtained by pushing aspirated fat through size reduction filters while allowing waste products to exit in a closed system.

Raposio et al. (2014) also described an effective alternative procedure by mechanical separation using a laminar airflow bench. Once the adipose tissue was harvested during a conventional tumescent liposuction, it was placed in a vibrating shaker at 600 vibrations per minute for 6 min and then centrifuged at 1600 rpm for 6 min so that the previously detached ADSCs were collected into the resulting pellet at the bottom of each 10-mL test tube containing the lipoaspirate (crude SVF), then collected using a pipette and added to a 10-mL Luer Lock syringe, ready to be injected. The entire isolation process lasted approximately 15 min and it yielded a mean of 5 x 105 (SD: ±1 x 105) ADSCs with 97% cell viability.

The surge in clinical applications for ADSCs increases the need for clear & reliable information about the efficiency, cost & safety of automated equipment & manual techniques which facilitate separation of the stromal vascular fraction (SVF) from adipose tissue.

Effect of Fat Processing Technique on the
Quality of "ADSCs"

A prospective cross-sectioned study evaluated three widely used fat processing techniques in plastic surgery for the viability and number of adipocytes and ADSCs isolated from collected lipoaspirate (Conde-Green et al. 2010).

  1. Decantation
  2. Washing
  3. Centrifugation

Flow cytometric analysis has revealed various differences in ADSCs comparing the middle firm tissue layers of all three different processing techniques.

  • The firm layer of decantation process contained large amount of blood contaminants and very few ADSCs & endothelial cells.
  • The firm layer of washing process contained few blood contaminants & more endothelial cells & ADSCs, compared to the decantation process.
  • The firm layer of centrifuged sample contained the least number of ADSCs, blood contaminants, and endothelial cells, whereas the pellet of the centrifuged contained the greatest number of ADSCs, blood contaminants, and endothelial cells. In addition, the firm tissue layer did not expand and proliferate in vitro, while the pellet of the centrifuged samples demonstrated extensive proliferation and expansion (Conde-Green et al. 2010). This finding has been confirmed recently by Iyyanki et al (2015) comparing centrifuged and non-centrifuged sample collected from subcutaneous adipose in the abdominal area using the Coleman technique and revealed that the centrifuged samples contained significantly greater SVF & ADSCs yield. The result of this study also confirmed the finding of Tommaso et al. (2012) that the ADSCs are tougher cells than adipocytes and can withstand centrifugal process of up to 3000 rpm (Conde-Green et al. 2010).
  • Centrifugation of adipose tissue separates fat cells from lipid, blood cells, water, and water-soluble ingredients such as proteases and lipases, but does not shift ADSCs between the adipose and fluid portions, possibly due to the strong adherence to adipose tissue or since they are resident with the adipose tissue. It is shown also that the increased centrifugal forces compacted the adipose portion more and therefore concentrated the red cell within the adipose portion rather than shifting the red blood cells into the fluid portion.
  • It was found that the yield of ADSCs in culture for 1 week was consistent up to 300 g but decrease with centrifugal forces of more than 3000 g (Kurita et al. 2008), Dickens et al (2009) revealed that gentle centrifugation produced the highest cell viability, whereas long period of centrifugation resulted in the selection of most proliferate ADSCs subpopulation.

2. Cultured-Adipocyte-derived Stem Cells

Cultured- ADSCs

After taking fat from connective tissue underneath the belly fat or any area of the body where there is unwanted extra fat tissue, send it to lab for isolation and incubation and multiplying of stem cells to a number sufficient to inject and treat the affected area. The cultured-ADSCs re-inject it to the same person for area of treatment.

The cells called adipose-derived stromal cells (ADSCs) must be distinguished from the crude stromal vascular fraction (SVF) obtained after digestion of adipose tissue. ADSCs share many features with mesenchymal stem cells derived from bone marrow, including paracrine activity, but they also display some specific features, including a greater angiogenic potential.

ADSCs are generally considered to be stable throughout long-term culture, as it has been reported that even ADSCs that had passed through more than 100 population doublings had a normal diploid karyotype (2008). The proliferation of ADSCs can be stimulated by various growth factors via different signalling pathways e.g. Izadpanah, (2002)

  • Fibroblast growth factor-2 (FGF-2)
  • Epithelial growth factor (EGF)
  • Insulin-like growth factor-1 (IGF-1),
  • Platelet-derived growth factor (PDGF), and
  • Tumour necrosis factor-a (TNF-a)

Factors influencing ADSCs Isolation & Expansion
“Cultured ADSCs”

Studies have suggested that ADSCs exhibit an average population doubling time of 60 hours or generally 2-4 days, depending on the below factors: (Fossett et al. 2012, Gimble et al. 2007, Misuno 2009)

  1. Donor’s age
  2. The type of fat WAT or BAT
  3. Location of adipose tissue; subcutaneous or visceral
  4. Type of surgical procedure
  5. Culture conditions
  6. Growth factors
  7. Plating or seeding densities
  8. Passage number
  9. Medial formulation
  10. Other factor to consider in the isolation process is the effect of seeding density on cell proliferation. Fossett et al. (2012) showed that low seeding densities increase the proliferation density on the MSC proliferation was demonstrated with BmMSCs that were seeded at 100 cells/ cm2 and reached their target of 200X106 cells 4.1 days faster than cells seeded at 5000 cells / cm2  (Both et al.2007)

Characteristics of Cultured- ADSCs

Like other Mesenchymal Stem Cells (MSCs), undifferentiated ADSCs must be identified using a panel of surface markers on which no clear agreement has been reached. However, based on current literature, markers that are uniformly reported to have strong positive expression are CD13, CD29, CD34 (a well-established stem cell marker), CD44, CD73, CD90, CD105 (a mesenchymal stem cell-associated marker). 

CD166 and MHC I, CD14, CD31, CD45, and CD133 (all markers of the hematopoietic & angiogenic lineages) have been reported to show low or no expression in ADSCs.

Cytometric analysis of adipose-derived stem cells (ASCs) has shown that these cells do not express CD31 and CD45, but do express CD34, CD73, CD105, & the mesenchymal stem cell marker CD90. ASCs have a differentiation potential similar to that of other mesenchymal stem cells as well as a higher yield upon isolation and a greater proliferative rate in culture than bone marrow-derived stem cells.

The International Society for Cellular Therapy (ISCT) has proposed minimum criteria to define MSCs. These cells should:

  1. Exhibits plastic adherence
  2. Express cell surface markers like cluster of differentiation (CD)29, CD44, CD73, CD90, CD105& lack expression of CD14, CD34, CD45 and human leucocyte antigen-DR (HLA-DR) and
  3. Have the ability to differentiate in vitro into adipocyte (fat tissue), chondrocyte (cartilage tissue) and osteoblast (bone tissue).

These characteristics are valid for all MSCs, although few differences exist in MSCs isolated from various tissue origins.

Read More- References:

  1. A. Salibian, A.D. Widgerow, M. Abrouk, G.R. Evans, Stem cells in plastic surgery: a review of current clinical and translational applications, Arch. Plast. Surg. 40 (2013) 666675. Click here
  2. Higuci, C.W. Chuang, Q.D. Ling, et al., Differentiation ability of adiposederived stem cells separated from adipose tissue by a membrane filtration method, J. Memb. Sci. 366 (2011) 286294. Clickhere
  3. J. Katz, R. Llull, M.H. Hedrick, J.W. Futrell, Emerging approaches to the tissue engineering of fat, Clin. Plast. Surg. 26 (1999) 587603. Click here
  4. M. Altman, F.J. Abdul Khalek, E.U. Alt, C.E. Butler, Adipose tissue-derived stem cells enhance bioprosthetic mesh repair of ventral hernias, Plast. Reconstr. Surg. 126 (2010) 845854. Click here
  5. Al Battah, J. De Kock, E. Ramboer, et al., (2011) Evaluation of the multipotent character of human adipose tissue-derived stem cells isolated by Ficoll gradient centrifugation and red blood cell lysis treatment, Tox Vitro 25, 1224-1230. Click Here for PDF
  6. Aronowitz, J. A., Lockhart, R. A., & Hakakian, C. S. (2015). Mechanical versus enzymatic isolation of stromal vascular fraction cells from adipose tissue. Springerplus, 4, 713. Click Here for PDF
  7. Aust, B. Devlin, S.J. Foster, et al., Yield of human adipose-derived adult stem cells from liposuction aspirates, Cytotherapy 6, (2004) 7-14. Click Here for PDF
  8. Baptista, R.F.J.C. Amaral, R.B.V. Carias, M. Aniceto, C.C. Silva, R. Borojevic, (2009) An alternative method for the isolation of mesenchymal stromal cells derived from lipoaspirate samples, Cytotherapy 11, 706-715.Click Here for PDF
  9. Banyard, A.A. Salibian, A.D. Widgerow, G.R. Evans, (2015) Implications for human adipose-derived stem cells in plastic surgery, J. Cell Mol. Med. 19, 21-30. Click Here for PDF
  10. Bellei, B., Migliano, E., Tedesco, M., Caputo, S., & Picardo, M. (2017). Maximizing non-enzymatic methods for harvesting adipose-derived stem from lipoaspirate: technical considerations and clinical implications for regenerative surgery. Sci Rep, 7(1), 10015. Click here for PDF
  11. Bertolini F, Lohsiriwat V, Petit JY et al (2012) Adipose tissue cells, Lipotransfer and cancer: a challenge for scientists, oncologists and surgeons. Biochim Biophys Acta 1862:209–214 Click Here for PDF
  12. Bertozzi, N., Simonacci, F., Grieco, M. P., Grignaffini, E., & Raposio, E. (2017). The biological and clinical basis for the use of adipose-derived stem cells in the field of wound healing. Ann Med Surg (Lond), 20, 41-48. Click Here for PDF
  13. Bianchi, M. Maioli, E. Leonardi, et al., (2013) A new nonenzymatic method and device to obtain a fat tissue derivative highly enriched in pericyte-like elements by mild mechanical forces from human lipoaspirates, Cell Transpl. 22, 2063-2077. Click Here for PDF
  14. Bielli A, Scioli MG, Gentile P et al (2014) Adult adipose-derived stem cells and breast cancer: a controversial relationship. Springerplus 3:345 Click Here for PDF
  15. Bourin P, Bunnell BA, Casteilla L et al (2013) 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 and Science (IFATS) and the International Society for Cellular Therapy (ISCT). Cytotherapy 15:641–648 Click Here for PDF
  16. Bunnell BA, Flaat M, Gagliardi C, Patel B, Ripoll C (2008) Adipose-derived stem cells: isolation, expansion and differentiation. Methods 45(2):115–120 Click here for PDF
  17. Bouquest, A. Shahdadfar, J.E. Brinchmann, et al., (2006) Isolation of stromal stem cells from human adipose tissue, Methods Mol. Biol. 325, 35-46. Click Here for PDF
  18. M. Cowan, Y.Y. Shi, O.O. Aalami, et al., Adipose-derived adult stromal cells heal critical-size mouse calvarial defects, Nat. Biotechnol. 22 (2004) 560567. Click here
  19. Caruana, N. Bertozzi, E. Boschi, M.P. Grieco, E. Grignaffini, E. Raposio, (2015) Role of adipose-derived stem cells in chronic cutaneous wound healing, Ann. Ital. Chir. 86, 1-4. Click Here for PDF
  20. Chen YW, Wang JR, Liao X, Li SH, Xiao LL, Cheng B, Xie GH, Song JX, Liu HW. Effect of suction pressures on cell yield and functionality of the adipose-derived stromal vascular fraction.J Plast Reconstr Aesthet Surg. 2017 Feb;70(2):257-266.
  21. Cheriyan T, Kai Kao H, Qiao X, Guo L (2014) Low harvest pressure enhances autologous fat graft viability. Plast Reconstr Surg 133:1365–1368 Click Here for PDF
  22. Chung MT, Paik KJ, Atashroo DA, Hyun JS, McArdle A, Senarath-Yapa K, Zielins ER, Tevlin R, Duldulao C, Hu MS, Walmsley GG, Parisi-Amon A, Momeni A, Rimsa JR, Commons GW, Gurtner GC, Wan DC, Longaker MT (2014) Studies in fat grafting: part I. Effects of injection technique on in vitro fat viability and in vivo volume retention. Plast Reconstr Surg 134:29–38 Plast Reconstr Surg 132:35–46 Click Here for PDF
  23. Committee for Advanced Therapies (CAT), Overview of Comments Received on ‘Reflection Paper on Classification of Advanced Therapy Medicinal Products’ (EMA/CAT/600280/2010 rev. 1), 2015.
  24. Conde-Green A, Rodriguez RL, Slezak S et al (2014) Enzymatic digestion and mechanical processing of aspirated adipose tissue. Plast Recons Surg 134:54 Click Here for PDF
  25. Conde´-Green A, de Amorim NF, Pitanguy I (2010) Influence of decantation, washing and centrifugation on adipocyte and mesenchymal stem cell content of aspirated adipose tissue: a comparative study. J Plast Reconstr Aesthet Surg 63:1375–1381 Click Here for PDF
  26. CPadoin AV, Braga-Silva J, Martins P, Rezende K, Rezende AR, Grechi B, Gehlen D, Machado DC. Sources of processed lipoaspirate cells: influence of donor site on cell concentration. Plast Reconst Surg. 2008;122(2):614-8 Click Here for PDF
  27. A. Banyard, A.A. Salibian, A.D. Widgerow, G.R. Evans, Implications for human adipose-derived stem cells in plastic surgery, J. Cell Mol. Med. 19 (2015) 2130.
  28. D.C. Wu, A.S. Byod, K.J. Wood, Embryonic stem cell transplantation: potential applicability in cell replacement therapy and regenerative medicine, Front. Biosci. 12 (2007) 45254535.
  29. Daher, B.H. Johnstone, D.G. Phinney, et al., (2008) Adipose stromal/stem cells:basic and translational advances: the IFATS collection, Stem Cell 26 Click Here for PDF
  30. Dahl, S. Duggal, N. Coulston, et al., (2008) Genetic and epigenetic instability of human bone marrow mesenchymal stem cells expanded in autologous serum or fetal bovine serum, Int. J. Dev. Biol. 52 1033-1042. Click Here for PDF
  31. Dominici M, Le Blanc K, Mueller I et at. (2006) Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8(4):315-317 Click Here for PDF
  32. Dubey, N., et al. (2018). “Revisiting the Advances in Isolation, Characterization and Secretome of Adipose-Derived Stromal/Stem Cells.” International Journal of Molecular Sciences 19(8): 2200. Click here for PDF
  33. Duscher, Zeshaan N. Maan, Anna Luan, Matthias M. Aitzetmüller, Elizabeth A. Brett, David Atashroo, Alexander J. Whittam, Michael S. Hu, Graham G. Walmsley, Khosrow S. Houschyar, Arndt F. Schilling, Hans-Guenther Machens, Geoffrey C. Gurtner, Michael T. Longaker, Derrick C. Wan. Ultrasound-assisted liposuction provides a source for functional adipose-derived stromal cells Dominik, Cytotherapy. Sep 2017
  34. Raposio, C. Guida, I. Baldelli, F. Benvenuto, M. Curto, L. Paleari, et al., Characterization and induction of human pre-adipocytes, Toxicol. In Vitro 21 (2007) 330334.
  35. Raposio, C. Guida, R. Coradeghini, C. Scanarotti, A. Parodi, I. Baldelli, et al., In vitro polydeoxyribonucleotide effects on human pre-adipocytes, Cell Prolif. 41 (2008) 739754.
  36. Raposio, G. Caruana, M. Petrella, S. Bonomini, M.P. Grieco, A standardized method of isolating adipose-derived stem cells for clinical applications, Ann. Plast. Surg. 76 (2016) 124126.
  37. Raposio, G. Caruana, S. Bonomini, G. Libondi, A novel and effective strategy for the isolation of adipose-derived stem cells: minimally manipulated adipose-derived stem cells for more rapid and safe stem cell therapy, Plast. Reconstr. Surg. 133 (2014) 14061409.
  38. Raposio, N. Bertozzi, S. Bonomini, et al., Adipose-derived stem cells added to platelet-rich plasma for chronic skin ulcer therapy, Wounds 28 (2016) 126131.
  39. Simonacci, N. Bertozzi, M.P. Grieco, E. Grignaffini, E. Raposio, Autologous fat transplantation for breast reconstruction: a literature review, Ann. Med. Surg. (Lond) 12 (2016) 94100.
  40. Fadel, B.R. Viana, M.L.T. Feitosa, et al., (2011) Protocols for obtainment and isolation of two mesenchymal stem cell sources in sheep, Acta Cir. Bras. 26, 267-273. Click Here for PDF
  41. Fogarty WM, Griffin PJ (1973) Production and purification of metalloprotease of Baccilus polymyxa. Appl Microbiol 26(2):185–190 Click Here for PDF
  42. Fossett E, Khan WS, Longo UG et al (2012) Effect of age and gender on cell proliferation and cell surface characterization of synovial fat pad derived mesenchymal stem cells J Orthop Res 30:1013-1018 Click Here for PDF
  43. Francis, P.C. Sachs, L.W. Elmore, S.E. Holt, (2010) Isolating adipose- derived mesenchymal stem cells from lipoaspirate blood and saline fraction, Organogenesis 6, 11-14. Click Here for PDF
  44. Fraser JK, Wulur I, Alfonsno Z (2006) Fat tissue:an underappreciated source of stem cells for biotechnology. Trends Biotechnol 24:150-154 Click Here for PDF
  45. Caruana, N. Bertozzi, E. Boschi, M. Pio Grieco, E. Grignaffini, E. Raposio, Role of adipose-derived stem cells in chronic cutaneous wound healing, Ann. Ital. 86 (2015) 14.
  46. D. Yancopoulos, S. Davis, N.W. Gale, J.S. Rudge, S.J. Wiegand, J. Holash, Vascular-specific growth factors and blood vessel formation, Nature 407 (2000) 242248.
  47. Rigotti, A. Marchi, M. Galie, et al., Clinical treatment of radiotherapy tissue damage by lipoaspirate transplant: a healing process mediated by adipose derived adult stem cells, Plast. Reconstr. Surg. 119 (2007) 14091422.
  48. Gimble, F. Guilak, (2003) Adipose-derived adult stem cells: isolation, characterization, and differentiation potential,Cytotherapy, 2003;5(5):362-9.Click Here for PDF
  49. Gimble JM, Nuttall ME (2011) Adipose-derived stromal/stem cells (ASC) in regenerative medicine: pharmaceutical applications. Curr Pharm Des 17(4):332–339
  50. Gimble JM, Bunnell BA, Guilak F (2012) Human adipose-derived cells: an update on the transition to clinical translation. Regen Med 7(2):225–235
  51. Gimble JM, Nuttall ME (2011) Adipose-derived stromal/stem cells (ASC) in regenerative medicine: pharmaceutical applications. Curr Pharm Des 17(4):332–339
  52. Gimble JM, Adam JK, Bunnell BA (2007) Adipose-derived stem cells for regenerative medicine. Circ Res 100:1249-1260 Click Here for PDF
  53. Gimble, F. Guilak, (2003) Adipose-derived adult stem cells: isolation, characterization, and differentiation potential, Cytotherapy 5 Click Here for PDF
  54. Gimble JM, Bunnell BA, Guilak F (2012) Human adipose-derived cells: an update on the transition to clinical translation. Regen Med 7(2):225–235
  55. Gnanasegaran N, Govindasamy V, Musa S et al (2014) Different isolation methods alter the gene expression profiling of adipose derived stem cell. Int J Med Sci 11:391-403 Click Here for PDF
  56. Gonzalez-Rey, M.A. Gonzalez, N. Varela, et al., (2010) Human adipose-derived mesenchymal stem cells reduce inflammatory and T cell responses and induce regulatory T cells in vitro in rheumatoid arthritis, Ann. Rheum. Click Here for PDF
  57. Griffin PJ, Fogarty WM (1973) Physiochemical properties of the native, zinc-and manganese-prepared metalloprotease of Bacillus polymyxa. Appl Microbiol 26(2):191–195 Click Here for PDF
  58. Chang, B.R. Do, J.H. Che, et al., Safety of adipose-derived stem cells and collagenase in fat tissue preparation, Aesthetic Plast. Surg. 37 (2013) 802808.
  59. Han J, Koh YJ, Moon HR, Ryoo HG, Cho CH, Kim I et al (2010) Adipose tissue is an extramedullary reservoir for functional hematopoietic stem and progenitor cells. Blood 115:957–964 Click Here for PDF
  60. Hoppe DL, Ueberreiter K, Surlemont Y, Peltoniemi H, Stabile M, Kauhanen S (2013) Breast reconstruction de novo by water-jet assisted autologous fat grafting–a retrospective study. Ger Med Sci 11:Doc17 Click Here for PDF
  61. Porro, A. Schenone, M. Fato, E. Raposio, E. Molinari, F. Beltrame, An integrated environment for plastic surgery support: building virtual patients, simulating interventions, and supporting intraoperative decisions, Comput. Imaging Graph 29 (2005) 385394.
  62. Iyyanki T, Hubenak J, Liu U et at (2015) Harvesting technique affects adipose-derived stem cell yield. Aesthet Surg J 35(4):467-476 Click Here for PDF
  63. Izadpanah, D. Kaushal, C. Kriedt, et al., (2008) Long-term in vitro expansion alters the biology of adult mesenchymal stem cells, Cancer Click Here for PDF
  64. Fan, E. Raposio, J. Wang, R.E. Nordstrom, Development of the inframammary fold and ptosis in breast reconstruction with textured tissue expanders, Aesthetic Plast. Surg. 26 (2002) 219222.
  65. Jurgens WJFM, Oedayrajsingh-Varma MJ, Helder MN et. (2008) Effect of tissue-harvesting site on yield of stem cells derived from adipose tissue: implications for cell -based therapies. Cell Tissue Res 332:415-426 Click Here for PDF
  66. Schenke-Layland, B.M. Strem, M.C. Jordan, et al., Adipose tissue-derived cells improve cardiac function following myocardial infarction, J. Surg. Res. 153 (2009) 217e223.
  67. H. Cheng, T.L. Kuo, K.K. Kuo, et al., Human adipose-derived stem cells: isolation, characterization and current application in regeneration medicine, GMBHS 3 (2011) 5362.
  68. K. Masuoka, T. Asazuma, H. Hattori, et al., Tissue engineering of articular cartilage with autologous cultured adipose tissue-derived stromal cells using atelocollagen honeycomb-shaped scaffold with a membrane sealing in rabbits, J. Biomed. Mater Res. B 79 (2006) 2534.
  69. L. Gould, P. Abadir, H. Brem, et al., Chronic wound repair and healing in older adults: current status and future research, Wound Rep. Reg. 23 (2015) 113.
  70. Kaufman MR, Bradley JP, Dickinson B, Heller JB, Wasson K, O’Hara C, Huang C, Gabbay J, Ghadjar K, Miller TA (2007) Autologous fat transfer national consensus survey: trends in techniques for harvest, preparation, and application, and perception of short- and long-term results. Plast Reconstr Surg 119:323–331 Click Here for PDF
  71. Ke Q, Chen A, Minoda M et al (2013) Safety evaluation of a thermolysin enzyme from Geobacillus stearothermophilus. Food Chem Toxicol 59:541–548 Click Here for PDF
  72. Kling RE, Mehrara BJ, Pusic AL, Young VL, Hume KM, Crotty CA, Rubin JP (2013) Trends in autologous fat grafting to the breast: a national survey of the American society of plastic surgeons. Plast Reconstr Surg 132:35–46 Click Here for PDF
  73. Kokai, K. Marra, J.P. Rubin, (2014) Adipose stem cells: biology and clinical applications for tissue repair and regeneration, Trans. Click Here for PDF
  74. Kretlow JD, Jin YQ, Liu W, Zhang WJ, Hong TH, Zhou G, Baggett LS, Mikos AG, Cao Y (2008) Donor age and cell passage affects differentiation potential of murine bone marrowderived stem cells. BMC Cell Biol 9:60
  75. L. Reoseti, M. Serra, D. Tigani, et al., M. Cell manipulation in autologous chondrocyte implantation: from research to cleanroom, Chir. Organi Mov. 91 (2008) 147151.
  76. Li K, Gao J, Zhang Z, Li J, Cha P, Liao Y, Wang G, Lu F (2013) Selection of donor site for fat grafting and cell isolation. Aesthet Plast Surg 37:153–158 Click Here for PDF
  77. E. Kokai, K. Marra, J.P. Rubin, Adipose stem cells: biology and clinical applications for tissue repair and regeneration, Trans. Res. 163 (2014) 399408.
  78. Li J, Curley JL, Floyd ZE, Wu X, Halvorsen YDC, Gimble JM. Isolation of Human Adipose-Derived Stem Cells from Lipoaspirates. Methods Mol Biol. 2018;1773:155-165. 
  79. Liu G, Chen X. Isolating and Characterizing Adipose-Derived Stem Cells.Methods Mol Biol. 2018;1842:193-201. doi: 10.1007/978-1-4939-8697-2_13
  80. E. Kokai, K. Marra, J.P. Rubin, Adipose stem cells: biology and clinical applications for tissue repair and regeneration, Trans. Res. 163 (2014) 399408.
  81. M.G. Aluigi, R. Coradeghini, C. Guida, et al., Pre-adipocytes commitment to neurogenesis 1: preliminary localisation of cholinergic molecules, Cell Biol. Int. 33 (2009) 594601.
  82. M. Klinger, F. Caviggioli, F.M. Klinger, et al., Autologous fat graft in scar treatment, J. Craniofac. Surg. 24 (2013) 16101615.
  83. Markarian FM, Frey GZ, Silveira MD et al (2014) Isolation of adipose-derived stem cells: a comparison among different methods. Biotechnol Lett 36:693–702 Click Here for PDF
  84. Matsumoto D, Shigeura , Sato T, Inoue K, Suga H,Kato H, Aoi N, Murase S, Gonda K, Yoshimura K, Influence of preservation at various temperature on liposuction aspirates.Plast Reconst Surg 2007;120(6):1510-7 Click Here for PDF
  85. McIntosh K, Zvonic S, Garrett S et al (2006) The immunogenicity of human adipose derived cells: temporal changes in vitro. Stem Cells 24:1245–1253 Click Here for PDF
  86. Meyer J, Salamon A, Herzmann N et al (2015) Isolation and differentiation potential of human mesenchymal stem cells form adipose tissue harvested by water jet-assisted liposuction. Aesth Surg J. doi:101093/asj/sjvo75 Click Here for PDF
  87. Meza-Zepeda, A. Noer, J.A. Dahl, et al., (2008) High-resolution analysis of genetic stability of human adipose tissue stem cells cultured to senescence, J. Cel. Mol. Med. 12 553-563. Click Here for PDF
  88. Mizuno H (2009) Adipose-derived stem cells for tissue repair and regeneration: ten years of research and a literature review. J Nippon Med Sch 76 (2):56-66 Click Here for PDF
  89. Mizuno, M. Tobita, A.C. Uysal, (2012) Concise review: adipose-derived stem cells as a novel tool for future regenerative medicine, Stem Cell 30, 804-810.Click Here for PDF
  90. Mizuno, (2009) Adipose-derived stem cells for tissue repair and regeneration: ten years of research and a literature review, J. Nippon. Med. Sch. 76, 56-66.Click Here for PDF
  91. Mizuno H, Tobita M, Uysal AC (2012) Adipose-derived stem cells as a novel tool for future regenerative medicine. Stem Cells 30(5):804–810
  92. Nie, D. Yang, J. Xu, et al., Locally administered adipose-derived stem cells accelerate wound healing through differentiation and vasculogenesis, Cell 20 (2011) 205216.
  93. Oedayrajsingh-Varma MJ, Van Ham SM, Knippenberg M et al. (2006) Adipose tissue-derived mesenchymal stem cell yield and growth characteristics are affected by the tissue-harvesting procedure. Cytotherapy 8(2):166-177 Click Here for PDF
  94. A. Zuk, Tissue engineering craniofacial defects with adult stem cells? Are we ready yet? Ped Res. 63 (2008) 478486.
  95. R. Coradeghini, C. Guida, C. Scanarotti, et al., A comparative study of proliferation and hepatic differentiation of human adipose-derived stem cells, Cells Tissues Organs 191 (2010) 466477.
  96. Raposio, G. Caruana, S. Bonomini, G. Libondi, (2014) A novel and effective strategy for the isolation of adipose-derived stem cells: minimally manipulated adipose-derived stem cells for more rapid and safe stem cell therapy, Plast. Reconstr. Surg. 133, 1406-1409. Click Here for PDF
  97. Raposio, G. Caruana, M. Petrella, S. Bonomini, M.P. Grieco, (2016) A standardized method of Isolating adipose-derived stem cells for clinical applications, Ann. Plast. Surg. 76, 124-126. Click Here for PDF
  98. Raposio, N. Bertozzi, S. Bonomini, et al., (2016) Adipose-derived stem cells added to platelet-rich plasma for chronic skin ulcer therapy, Wounds 28, 126-131. Click Here for PDF
  99. Raposio E, Caruana G, Bronomini S, Libondi G (2014) A novel strategy for the isolation of adipose-derived stem cells: minimally manipulated adipose-derived stem cells for more rapid and safe stem cell therapy. Plast Reconstr Surg 133(6):1406–1409 Click Here for PDF
  100. Raposio E, Simonacci F, Perrotta RE. Adipose-derived stem cells: Comparison between two methods of isolation for clinical applications.Ann Med Surg (Lond). 2017 Jul 8;20:87-91.
  101. Raposio, V. Belgrano, P. Santi, C. Chiorri, Which is the ideal breast Size?: some social clues for plastic surgeons, Ann. Plast. Surg. 76 (2016) 340345.
  102. Raposio, S. Cicchetti, M. Adami, R.G. Ciliberti, P.L. Santi, Computer planning for breast reconstruction by tissue expansion: an update, Plast. Reconstr. Surg. 113 (2004) 20952097.
  103. Raposio, P. Caregnato, P. Barabino, et al., Computer-based preoperative planning for breast reconstruction in the woman with unilateral breast hypoplasia, Minerva Chir. 57 (2002) 711714.
  104. E. Raposio, N. Bertozzi
    How to isolate a ready-to-use adipose-derived stem cells pellet for clinical application. Eur Rev Med Pharmacol Sci. (2017);Vol. 21 – N. 18, P.: 4252-4260

  105. Ray R, Novotny NM, Crisostomo PR et al (3008) sex steroids and stem cells function. Mol Med 14:493:501 Click Here for PDF
  106. Reoseti, M. Serra, D. Tigani, et al., M., (2008) Cell manipulation in autologous chondrocyte implantation: from research to clean room, Chir. Organi Mov. 91, 147-151. Click Here for PDF
  107. Rohrich RJ, Sorokin ES, Brown SA. In search of improved fat transfer viability: a quantitative analysis of the role of centrifugation and harvest site. Plast Reconst Surg. 2004:113(1):391-5 Click Here for PDF
  108. Rohrich RJ, Sorokin ES, Brown SA (2004) In search of improved fat transfer viability: a quantitative analysis of the role of centrifugation and harvest site. Plast Reconstr Surg 113:391–397 Click Here for PDF
  109. Rubin JP,Marra KG. Adipose stem cell therapy for soft tissue reconstruction. Lancet. 2013;382(9898):1077-9 Click Here for PDF
  110. S. Brongo, G.F. Nicoletti, S. La Padula, et al., Use of lipofilling for the treatment of severe burn outcomes, Plast. Reconstr. Surg. 130 (2012) 374e376.
  111. R. Daher, B.H. Johnstone, D.G. Phinney, et al., Adipose stromal/stem cells: basic and translational advances: the IFATS collection, Stem Cell 26 (2008) 26642665.
  112. Santos FD, Andrade PZ, Abecasis MM, Gimble JM, Chase LG, Campbell AM, Boucher S, Vemuri MC, Silva CL, Cabral JM (2011) Toward a clinical-grade expansion of mesenchymal stem cells from human sources: a microcarrier-based culture system under xenofree conditions. Tissue Eng Part C Methods 17(12):1201–1210
  113. Shiffman MA, Mirrafati S (2001) Fat transfer techniques: the effect of harvest and transfer methods on adipocyte viability and review of the literature. Dermatol Surg 27:819–826 Click Here for PDF
  114. Salibian, A.D. Widgerow, M. Abrouk, et al., (2013) Stem cells in plastic surgery: a review of current clinical and translational applications, Arch. Plast. Surg. 40 666e675. Click Here for PDF
  115. Santos FD, Andrade PZ, Abecasis MM, Gimble JM, Chase LG, Campbell AM, Boucher S, Vemuri MC, Silva CL, Cabral JM (2011) Toward a clinical-grade expansion of mesenchymal stem cells from human sources: a microcarrier- based culture system under xeno-free conditions. Tissue Eng Part C Methods 17(12):1201–1210
  116. Scanarotti, A.M. Bassi, M. Catalano, et al., Neurogenic-committed human pre-adipocytes express CYP1A isoforms, Chem. Biol. Interact. 184 (2010) 474483.
  117. Sethe S, Scutt A, Stolzing A. Aging of mesenchymal stem cells. Ageing Res Rev. 2006;5:91–116. 83.
  118. Small K, Choi M, Petruolo O, Lee C, Karp N (2014) Is there an ideal donor site of fat for secondary breast reconstruction? Aesthet Surg J 34:545–550 Click Here for PDF
  119. Scherberich A, Di Maggio N, McNagny KM. A familiar stranger: CD34 expression and putative functions in SVF cells of adipose tissue. World J Stem Cells 2013; 5(1): 1-8
  120. Tambasco D, Arena V, Grussu F, Cervelli D (2013) Adipocyte damage in relation to different pressures generated during manual lipoaspiration with a syringe. Plast Reconstr Surg 131:645–646 Click Here for PDF
  121. Tjabringa GS,Zandieh-Doulabi B, Helder MN el at (2008) The polymine spermine regulates osteogenic differentiation in adipose stem cells J Cell Mol Med 12:1710-1717 Click Here for PDF
  122. U.D. Wankhade, M. Shen, R. Kolhe, S. Fulzele, Advances in adipose-derived stem cells isolation, characterization, and application in regenerative tissue engineering, Stem Cells Int. 2016 (2016).
  123. Ueberreiter K, von Finckenstein JG, Cromme F, Herold C, Tanzella U, Vogt PM (2010) BEAULITM–a new and easy method for large-volume fat grafts. Handchir Mikrochir Plast Chir 42:379–385 Click Here for PDF
  124. S. Food and Drug Administration, CFR- Code of Federal Regulations Title wq. Part 1271: Human Cells, Tissues and Cellular and Tissue-based Products, U.S.
  125. Von Hamburg D, Hemmrich K, Haydarlioglu S et al (2004) Comparison of viable cell yield from excised verssu aspirated adipose tissue. Cells Tissues Organs 178(2):87-92.Click Here for PDF
  126. H. Hassan, U. Udo Greiser, W. Wang, Role of adipose-derived stem cells in wound healing, Wound Rep. Reg. 22 (2014) 313325
  127. Witort EJ, Pattarino J, Papucci L et at. (2007) Autologous lipofilling: coenzyme Q10 can rescue adipocytes from stress-induced apoptotic death. Plast Reconst Surg 119:11911199 Click Here for PDF
  128. Wei, L. Niklason, D.M. Steinbacher, (2013) The effect of age on human adipose-derived stem cells, Plast. Reconstr. Surg. 131, 27-37. Click Here for PDF
  129. Kitagawa, M. Koroby, K. Toriyama, et al., History of discovery of humanadipose-derived stem cells and their clinical application, Jpn. J. Plast. Reconstr. Surg. 49 (2006) 10971104
  130. Y. Miyahara, N. Nagaya, M. Kataoka, et al., Monolayered mesenchymal stem cells repair scarred myocardium after myocardial infarction, Nat. Med. 12 (2006) 459465.
  131. Yoshimura, T. Shiguera, D. Matsumoto, et al., (2006) Characterization of freshly isolated and cultured cells derived from the fatty and fluid portions of liposuction aspirates, J. Cell Phys. 208, 64-76. Click Here for PDF
  132. Zhu, T. Liu, K. Song, et al., (2008) Adipose-derived stem cell: a better stem cell than BMSC, Cell Biochem. Funct. 26, 664-675. Click Here for PDF
  133. Zhu M, Heydarkhan-Hagvall S, Hedrick M, Benhaim P, Zuk P. Manual Isolation of Adipose-derived Stem Cells from Human Lipoaspirates. Journal of Visualized Experiments : JoVE. 2013;(79):50585. doi:10.3791/50585.
  134. Zimmerlin L, Donnenberg VS, Pfeifer ME et al (2010) Stromal vascular progenitors in adult human adipose tissue. Cytometry 77(1):22–30 Click Here for PDF
  135. Zuk PA, Zhu M, Mizuno H et al (2001) Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng 7:211–229 Click Here for PDF
  136. Zuk PA (2010) The adipose-derived stem cell: looking back and looking ahead. Mol Biol Cell 21(11):1783–1787
  137. Zuk, (2013) Adipose-derived stem cells in tissue regeneration: a review, ISRN Stem Cell, 1-35. Click Here for PDF
  138. Zuk, (2008) Tissue engineering craniofacial defects with adult stem cells? Are we ready yet? Pediatr. Click Here for PDF
  139. Zuk, M. Zhu, H. Mizuno, et al., (2001) Multilineage cells from human adipose tissue: implications for cell-based therapies, Tissue Eng. 7, 211-228 Click Here for PDF
  140. Zuk, M. Zhu, P. Ashjian, et al., (2002) Human adipose tissue is a source of multipotent stem cells, Mol. Biol. Cell 13 4279-4295. Click Here for PDF
  141. Zuk P.A, M. Zhu, H. Mizuno, et al., Multilineage cells from human adipose tissue:implications for cell-based therapies, Tissue Eng. 7 (2001) 211228.
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