This is a great question a wonderful patient asked me recently. The answer is that there are many ways to determine how many cells are in a milliliter of SVF with cell counters, but it is very difficult (outside of precise laboratory studies to my knowledge: see below **) to determine how many viable stem cells are actually present. Technology is almost there where automatic machines can give you such data, but they are still not readily available to doctors and their patients at this time (though I do have 2 calls into companies that are heading there...to see what the starting price tag is for such machines).
**See post http://eyedoc2020.blogspot.com/2018/02/what-is-best-source-of-stem-cells-bone.html
And search for "Isolation and Preparation of Adipose-derived Stem Cells."
It is not easy to prove viability of adipose derived stem cells. There are many cell counters, but these count all cells in a sample.
So the question is:
Should patients seek SVF injections or deployments as is currently formulated in its various forms (knowing enzymatic methods have more viable stem cells over manual methods)
OR
Risk further damage of tissue and decline of body function and wait for the technology to get better.
This is a very tough decision.
The technology is almost there, but there are not large scale, prospective randomized, double blinded studies to prove its use in every disease. And it is not cheap.
Why is that? Most feel it is because there are no drug companies involved that could do such a big study. They is no money for them if stem cells work as these cells are coming from inside the patient's own body.
Many MDs are using the manual method without enzyme use, but this has been proven to yield less viable stem cells. It is easier, though, and cheaper and does work in many studies noted below in certain diseases.
Enzymatic methods of isolating SVF cells from adipose tissue are better and used by the Cell Surgical Network, for instance, but they are more costly and time consuming.
This is a very confusing and frustrating time for patients and surgeons. Doctors want to heal their patients and do no harm. Patients want a guaranteed cure. We are so close, but still not there yet and no guarantee can be given or promises even with the best, purest stem cell isolate.
The first, double blinded study with stem cell injections was done and published by the Mayo Clinic: see here for post.
The below information hopes to educate all patients of where we stand today on Stromal Vascular Fraction which is a more accessible way to inject a patient's own stem cells into a diseased tissue.
For the dry eye issue, my plan is to use special, sterile filters to isolate out any further blood products and get further purity of the stem cell isolate.
All injection of SVF have risks. The biggest risk is the unknown. The second biggest risk is the risk of no improvement. The risk of infection, tumor growth, cancer appears to be very low if not zero in most situations. Still there is no 100% guarantee it will be a cure for the condition being treated.
Sandra Lora Cremers, MD, FACS
Below are 2 excellent articles about the status of SVF today.
The key points are:
Article 1:
1. The advantage of SVF over ADSCs is believed to be in two fundamental areas. Firstly, although similar in properties such as immunomodulation, anti-inflammatory, angiogenesis, and so forth, the distinctive, heterogeneous cellular composition of SVF may be responsible for the better therapeutic outcome observed in comparative animal studies [9–12].
Digestion of lipoaspirate is achieved by collagenase, and the presence of collagenase in the injectable product does not bode well with regulatory authorities such as the US Food and Drug Administration (FDA) [3]. Consequently, alternative methods are being explored with some encouraging outcomes [22–25].
The surge in clinical applications for ASCs increases the need for clear and reliable information about the efficiency, cost and safety of automated equipment and manual techniques which facilitate separation of the stromal vascular fraction (SVF) from adipose tissue. In clinical practice, adipose-derived stem cells are often not administered as a pure isolate but rather as one constituent of stromal vascular fraction, a heterogeneous mixture of cells resulting from the mechanical or enzymatic processing of aspirated adipose tissue. SVF contains a variety of cells including macrophages, various blood cells, pericytes, fibroblasts, smooth muscle cells, vascular endothelial progenitors and adipose-derived stem cells (Yoshimura et al. 2006; Bourin et al. 2013; Han et al. 2010; McIntosh et al. 2006; Bonab et al. 2006; Yoshimura et al. 2009). Stromal vascular fraction is one component of the heterogeneous mixture of adipose tissue fragments, stromal tissue, blood and tumescent fluid which constitutes lipoaspirate. The ASC content of SVF varies substantially depending on the method employed, with reports from less than 1 % of cells to over 15 % (Table 1). SVF cells can be safely isolated, quantified and characterized at the point of care in approximately 90 min. This is a timeframe which permits isolation and treatment to occur in the same surgical procedure, that is, at the point of care.
Article 2:
1.
Enzymatic methods
Enzymatic methods of isolating SVF cells from adipose tissue at the POC are based on a commonly used laboratory method of obtaining stem cells. The methods used to manually process adipose tissue using collagenase follow the same basic steps, but vary slightly in technique and reagents used. Lipoaspirate is washed 2–3 times using an aqueous salt solution such as PBS, Lactated Ringer’s solution, or Hank’s Balanced Salt Solution (HBSS). The washed lipoaspirate is then incubated with a collagenase solution of variable concentration and composition, depending on the method and tissue dissociation enzyme product used. Enzymatic digestion is typically carried out in a heated shaker to provide constant agitation at 37 °C for 30 min to 2 h. The digested adipose tissue is then centrifuged (speed/duration vary. See Table 1) which separates the processed lipoaspirate into three main layers, the oil/adipose tissue layer, the aqueous layer, and the pellet. The SVF is contained within the pellet, so the other layers are discarded, although SVF cells can be recovered from the aqueous layer (Yoshimura et al. 2006). The pellet is washed to remove any residual enzyme and filtered to remove tissue fragments and detritus. Collagenase-based enzymatic methods can be up to 1000 times more effective in SVF cell recovery than mechanical methods. Enzymatic methods are more efficient in isolating SVF cells because disruption of the collagen-based extracellular matrix (ECM) which binds together adipocytes and other cells of adipose tissue.
2.
Mechanical methods for SVF isolation report 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 (Table 1). Mechanical methods seek alternative non-enzymatic means of removing SVF cells from the adipose tissue and tend to be focused around washing and shaking/vibrating lipoaspirate followed by centrifugation in order to concentrate the SVF cells. All of the mechanical methods mentioned in this article contain a centrifugation step in order to concentrate the SVF cells. The composition of the cell populations recovered through simple centrifugation and other non-enzymatic methods have been shown to contain a greater frequency of peripheral blood mononuclear cells and a substantially lower number of progenitor cells (Conde-Green et al. 2014; Raposio et al. 2014; Shah et al. 2013)
3.
This a enclosed system to separate stem cells but waiting to hear from rep on how much it costs.
http://www.cytori.com/wp-content/uploads/2016/08/RM-045-LIT-EU_C-0615_SSCellBankBrochure_HR.pdf
Here is what the rep said:
Here is another automatic unit recommended in 2 papers below: waiting to hear from rep on cost.Here is what the rep said:
The Celution System will not be available in the U.S. for at least 18 months. In Europe the Celution device and single-use procedure set are $93,052 and $3,407, respectively.
Best regards,
Russ Havranek, MS MBA
VP, Global Marketing and Business Development
VP, Global Marketing and Business Development
http://www.tissuegenesis.com/icellator
4.
The main drawback of many of these devices is the cost of operation. The closed, enzymatic systems can be very expensive, with some costing over $50,000 for the system. In addition to purchasing the device, many require single-use disposable kits which can cost hundreds or thousands of dollars for a single disposable kit in some cases. A mechanical system like the StromaCell offers the benefit of a closed sterile system and tends to be more affordable, but does not provide the superior yield afforded by the enzymatic systems such as the Cytori Celution system or the Tissue Genesis Icellator system. All of the systems mentioned here can be operated by a single trained technician at the point of care. The processing times vary between systems, with mechanical systems being in the 15–30 min range and the enzymatic systems ranging from about 60–90 min depending on the amount of tissue processed.
5.
Despite a lack of reported clinical risk, in vitro studies have demonstrated potential oncological risks which clinicians should be cautious of when using SVF based therapies (Bertolini et al. 2012; Bielli et al. 2014). See full reference below.
These studies are also below and the 2014 study concludes:
"Preliminary data describe that SVF/ASCs enrichment did not show increased risk of new cancer or relapse compared with control group."
Stem Cell Res Ther. 2017; 8: 145.
Published online 2017 Jun 15. doi: 10.1186/s13287-017-0598-y
PMCID: PMC5472998
Adipose tissue-derived stromal vascular fraction in regenerative medicine: a brief review on biology and translation
Copyright © The Author(s). 2017
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
Background
Adipose-derived stem/stromal cells (ADSCs) were first characterised in 2001, and have since been widely studied and used as a major source of cells with regenerative potential, with characteristics similar to that of mesenchymal stem/stromal cells (MSCs) [1–4]. ADSCs are isolated as part of the aqueous fraction derived from enzymatic digestion of lipoaspirate (the product of liposuction). This aqueous fraction, a combination of ADSCs, endothelial precursor cells (EPCs), endothelial cells (ECs), macrophages, smooth muscle cells, lymphocytes, pericytes, and pre-adipocytes among others, is what is known as the stromal vascular fraction (SVF).
ADSCs, like MSCs, have shown promise in regenerative and reconstructive medicine [5–8]. Recent advances in the area of tissue regeneration have put SVF on a par and at times even above ADSCs [9–17]. For instance, in a study of erectile function in a rat model of cavernous nerve injury, SVF treatment showed superior statistically significant results compared to ADSC treatment alone, especially in smooth muscle/collagen ratio and in endothelial cell content [12]. The advantage of SVF over ADSCs is believed to be in two fundamental areas. Firstly, although similar in properties such as immunomodulation, anti-inflammatory, angiogenesis, and so forth, the distinctive, heterogeneous cellular composition of SVF may be responsible for the better therapeutic outcome observed in comparative animal studies [9–12]. Secondly, unlike ADSCs, SVF is much more easily acquired, without the need for any cell separation or culturing conditions. Thus, the therapeutic cellular product is instantaneously obtained and has minimal contact with reagents making it comparatively safer and subject to the fulfilment of lesser regulatory criteria. It should be noted that, whereas ADSCs find utility in both allogeneic and autologous treatments, SVF, owing to the presence of various cell types known to cause immunological rejection, is suitable for autologous treatments only.
Although almost all ADSCs are derived from the white adipose tissue (WAT), as covered in this review, the identification of progenitor cells in brown adipose tissue (BAT) of adult humans is fascinating and worth a mention [18, 19]. Termed as BADSCs (brown adipose-derived stem cells), these have been isolated from BAT deposits present in relatively inaccessible regions such as the mediastinum, and are capable of differentiating to metabolically active BA cells with differences in surface antigen expression as compared to WAT-originating ADSCs [18]. Current understanding of WAT and BAT define these cells with distinct functionalities, and thus translational avenues for ADSCs from either source should be compared to identify specific therapeutic targets and potential advantage of one over the other. Understanding of the molecular mechanisms behind either cell fate and the possibility of inter-conversion are interesting avenues of research with basic and translational implications [20, 21].
Despite the potential of SVF in regenerative medicine there are challenges to overcome. First is isolation of SVF, which needs a specialised infrastructure such as a clean room facility, equipment, reagents, and technical capabilities. These conditions limit the reach of SVF to only major hospitals in tier 1/2 cities, especially in a country such as India. In this regard, the up and coming point-of-care biomedical devices which can take lipoaspirate as their input and produce sterile, injectable SVF as output will be beneficial. Secondly, the method of isolating SVF is a vital roadblock in the approved use of SVF for therapeutic applications. Digestion of lipoaspirate is achieved by collagenase, and the presence of collagenase in the injectable product does not bode well with regulatory authorities such as the US Food and Drug Administration (FDA) [3]. Consequently, alternative methods are being explored with some encouraging outcomes [22–25]. Finally, characterisation of the regenerative cells of SVF has not reached a wide consensus. Organisations such as the International Federation for Adipose Therapeutics and Science (IFATS) and the International Society of Cellular Therapy (ISCT) have been updating the surface antigen-based definition of SVF cells, where CD34 antigen, primarily associated with haematopoietic stem cells (HSCs), became an important marker of regenerative, MSC-like cells of the SVF [1, 26, 27].
In this review, using the broader topics of isolation and characterisation of SVF, we will touch upon some of the challenges and innovations in the field and comment upon the future of SVF.
Isolation of SVF
Enzymatic isolation of SVF
The most widely used technique for the isolation of SVF from lipoaspirate is by digestion of the fatty portion of the lipoaspirate with collagenase, separating the contents into two distinct phases: the floating mature adipocytes fraction, and the cellular components of interest in the lower aqueous fraction [17, 28]. This separation can be enhanced by centrifugation; nevertheless, comparable separation can be achieved by gravity-based phase separation and filtration [29]. Although centrifugation is more efficient, it will also pellet down all the cells present, while filtration can be designed to capture only the important cell types based on size, thus enriching the specific cellular cocktail.
Non-enzymatic isolation of SVF
In view of the regulatory questions relating to enzymatic isolation, it is important to look into alternative methods for isolating SVF and compare these with the conventional methods [3, 24, 25]. Most of these techniques involve mechanical agitation which breaks down the adipose tissue and releases the stromal cells. As expected, the cellular yield from mechanical procedures are much lower compared to enzymatic methods, as cells of the adipose tissue tightly bound by collagen will not be easily released by mechanical action alone [24].
A novel method of mechanical agitation was recently defined by Tonnard et al. [23]. The injectable product, termed as “nanofat”, was obtained by emulsification and filtration of the lipoaspirate. Although termed as nanofat grafting, in effect no viable adipose cells survived the emulsification process, but the graft was rich in CD34+ ADSCs. The efficacy and properties of nanofat have been demonstrated in multiple case studies related to skin rejuvenation, scar healing, skin grafting for wound management, and treating vulvar lichen sclerosus (VLS), a chronic inflammatory disease of the anogenital area, and also by standard ADSC-related phenotypic and differentiation studies [23, 31, 32]. Owing to the simplicity of the technique, it might be amenable to scaling up by simply using the desired volume of syringe and/or using multiple syringes as required.
The effect of the emulsification process on other cells of interest, normally found in enzymatically processed SVF, remains to be seen. Combining such techniques with centrifugation or filtration can yield products highly concentrated with ADSCs, thus eliminating enzymatic digestion, reducing process time, cost, and respective regulatory constraints.
Automated devices for point-of-care isolation of SVF
The infrastructure, expertise, and consumables required for the conventional method of SVF isolation is not commonplace in most health-care facilities. Cosmetic surgery, being at the upper-end of medical expenditure, is the largest consumer of SVF and related products, but the actual scope is much wider [3]. Thus, it is unfortunate that the benefits of this very simple technology have not reached full potential. This gap can be overcome by automated, point-of-care biomedical devices, which can produce injectable SVF from lipoaspirate.
Such developments have been underway for quite some time, although mostly still in trial stages, with Cytori’s (San Diego, USA) Celution® being the first system [33]. Currently, about 30 different automated and semi-automated systems are under development [22]. The technologies and methodologies used vary, with most opting for the tried and tested enzymatic process. Stempeutics (Bangalore, India) has developed one such system, Stempeutron™, the proof-of-concept of which was reported in SundarRaj et al. [29]. Stempeutron™ uses the more efficient and conventional enzymatic digestion method and gravity-enabled separation of fatty and aqueous fraction followed by filtration of the aqueous fraction to achieve SVF isolation and concentration.
Since Stempeutron™ uses filtration we wanted to know the physical dimensions of SVF cells. As such, a list of cell sizes was not found while searching through the literature for this review and we resorted to mining for individual reports of cell size, surface area, and volume measurements. Table 1 summarises available cell diameter information accumulated from various reports [34–45]. The filtration system in Stempeutron™ is capable of capturing the majority of the therapeutically important cell types (Table 1) [3, 29]. Future developments might enable size-based enrichment of specific cellular populations, targeted towards specific diseases.
Characterisation of SVF
Criteria for characterising the cellular contents of SVF using surface antigen (cluster of differentiation (CD)) combinations is an evolving area of research as, within certain generally accepted norms, it differs between laboratories. A list of commonly used positive and negative markers identifying different cellular populations of SVF is provided in Table 1 [1, 26, 29]. Considering the variables present in isolation of SVF, such as the age of the patient, downstream processing, and so forth, the diversity observed between samples is quite understandable. However, if there is a relationship between the different ratios of cellular components present in SVF with its efficacy towards specific ailments, one might be able to come up with an optimum composition corresponding to the highest therapeutic efficacy. Traktuev et al. demonstrated that certain factors produced by ADSCs such as vascular endothelial growth factor (VEGF) help in migration, and that better survival of EPCs and correspondingly platelet-derived growth factors (PDGF)-BB produced by EPCs enable ADSCs to proliferate and migrate [46, 47]. They also provide proof of physical interaction between ADSCs and ECs in which ECs form a stable tubular, vasculature-like structure with support from ADSCs, both in vitro and in vivo [47]. This information along with some other articles has been used to draw up a schematic in Fig. 1 for the action of SVF, focussing on the interaction between ADSCs and EPCs [46–49].
Potential mechanism of action of ADSCs and ECs present in SVF towards angiogenesis. Breakdown of adipose tissue releases many cell types, which together are termed SVF. The cells of the SVF can produce several bioactive soluble factors. ADSCs and EPCs, ...
ADSCs in SVF are currently defined to be positive for classical MSC markers such as CD73 and CD90, and express CD34 but not the pan-haematopoietic lineage marker CD45. CD34 is expressed by progenitors of haematopoietic and endothelial lineages as well, and in ADSCs it is expressed transiently up to about 8–12 population doublings in culture [1].The case of CD34 is interesting since it is still largely considered to be a marker for HSCs owing to its historical association with the enrichment of such cells for bone marrow and umbilical cord blood transplantation. Even the pericytic theory related to MSCs and ADSCs has two sides [50]; whereas Crisen et al. attribute CD34– pericytes to be the progenitors of such stromal cells [51], Traktuev et al. demonstrated a CD34+ pericytic identity for ADSCs [46]. Maumus et al. tried to investigate this further but found that native CD34+ ADSCs did not exhibit in vivo pericytic markers, but they were rather observed over the course of the culture process [52]. Our data also show that both manually isolated and Stempeutron™-isolated SVF contains a CD146+ pericytic population that are mostly (>90%) CD34–[29], suggesting that freshly isolated SVF contains a pericytic population devoid of expressing both CD34 and CD31 markers. Whether the CD146+ cells observed within the SVF population subsequently become CD34+ ADSCs remains to be determined. Considerable evidence also exists in favour of CD34 expression in bone marrow-derived MSCs (BMMSCs), especially in the early stages of BMMSC research which included data on the disappearance of CD34 upon culturing [53]. Many aspects of this puzzle are yet to be solved, but it is probable that CD34 marks different progenitor cell types such as different MSCs and vascular endothelial progenitor cells.
In the course of preparing this review, it was also observed that reports of ADSC function and physiology in vitro is minimal and in vivo and/or in the native state is rare and in need of further investigation. Table 2summarises the observations about the characteristics of ADSCs in situ, in vivo, and in vitro that has been discussed within the review [1, 34–36, 46, 47, 52].
The curious case of CD34
ADSC research, being predominantly carried out using culture-expanded cells, has led to rather recent acceptance of CD34 as a marker for freshly isolated and native ADSCs. Thus, there remain interesting aspects of CD34 biology to be explored and understood. Firstly, CD34 expression has been associated with “stemness” in various systems including human ADSCs. A report by Suga et al. implied association of CD34 expression with naivety, angiogenic gene expression, and greater replicative capacity [54]. Similar to HSCs, reversal of CD34 expression has also been observed in MSCs with a change in culture conditions, thus hinting that CD34 expression might be reversible [53, 54]. Maumus et al. demonstrated an inverse relationship between CD34 expression and in vitro expansion of ADSCs and provided evidence for CD34 being a niche-specific marker of human ADSCs [52]. Interestingly, they commented on the morphological features of ADSCs in vivo, that is having up to 80-μm long protrusions, capable of forming networks surrounding mature adipocytes; however, the scientific and anatomical reason for these structural features are poorly understood. Taking these into account has led to speculation that CD34 is a physiological niche-specific marker of immature/early progenitor cells which is lost in in-vitro conditions [52–56]. Scherberich et al. review CD34 biology in general and with regards to ADSCs in detail [56].
The second interesting aspect is the relationship between CD34 and hypoxia. Since CD34 might be a niche-specific marker of progenitors, it can be speculated that hypoxic conditions might have something to do with its expression. Hypoxia is related to maintenance of adult stem cells such as those in bone marrow and neural stem cells [57]. In MSCs, and also recently in ADSCs, hypoxic pre-conditioning/culturing has shown improved results with regards to proliferation, retention of transplant, angiogenesis, and modulation of angiogenic factors such as VEGF and interleukin (IL)-6, homing, and mobilisation-related characteristics of MSCs/ADSCs, and so forth [58–63]. It is important to note that the ADSC study specifically selected for CD34– cells to begin with and subsequently did not find any significant expression of CD34 in their hypoxically cultured cells [63]. On the other hand, there was a study which speculated that the CD34 gene might be transcriptionally regulated by hypoxia inducible factor 1 (HIF1). The researchers observed that the concentration of oxygen in culture not only influenced the expression of CD34 but also that better maintenance of the antigen corresponded with more undifferentiated cells, which led them to hypothesise that CD34 and hypoxia play an important and inter-related function in maintenance of primitive stem cells of cord blood [64].
Such observations give a certain level of enigma; clearly CD34 and hypoxia are important factors in the maintenance of “stemness”, and it is also likely that CD34 expression is somehow related to hypoxic conditions in different stem or progenitor cell types. However, such a connection remains to be mechanistically studied in human ADSCs, or any other kind of MSCs for that matter. Such studies might provide evidence connecting CD34 with more naive/primitive stem cells, maintained in a hypoxic niche.
Current state in the clinic and laboratory
The first clinical applications of SVF were reported around 2007 to 2008 for cosmetic breast augmentation and also in the treatment of radiation injury post-radiotherapy in breast cancer patients [14, 65]. The Yoshimura group coined the term CAL, or cell-assisted lipotransfer, in 2008, where they enhanced fat grafts with SVF, demonstrating improved graft retention [14, 17]. Since these two early clinical reports from the last decade, there has been a many-fold increase in basic research and, consequently, many clinical trials are also now underway.
Searching www.ClinicalTrials.gov with keywords such as “SVF”, “Stromal vascular fraction”, “ADSC”, “Adipose stem cells”, and so forth, provides many hits. Although most of those studies are underway or recruiting at the time of this communication, interest has been rising with time. What is truly exciting is the breadth of conditions being targeted by SVF and ADSCs. Despite having properties like MSCs, the use of culture-expanded ADSCs has not reached similar consensus for allogeneic applications. However, ADSCs and SVF have been the preferred regenerative tools for use in autologous applications, and some of the major ones (along with case study references and/or ClinicalTrials.gov identification number) are listed in Table 3 [10, 14, 16, 23, 30, 31, 65–76]. Some other major ailments covered are pulmonary diseases, arterial and vascular diseases, graft versus host disease, Crohn’s disease, peripheral nerve regeneration, and so forth. Clinical areas where SVF and ADSCs are used do overlap to a substantial extent. Nevertheless, there are understandable differences between the two, but the few comparative pre-clinical and clinical studies available do not reach a unanimous conclusion. However, to summarise where the field stands as of now, a comparative overview of both modes with a few examples favouring either option is provided in Table 4[9, 11, 12, 77].
Major applications of SVF- and ADSC-based therapeutics with corresponding clinical trials and/or case study references
A superficial glance at the treatments highlights the two most preferred pathways, that is employing the vasculogenic and the immunomodulatory properties. We are yet to fully explore the multipotent properties of SVF cells which will only increase the breadth of their application. One recent example of enhanced osteoinduction by using SVF for dental implant surgery in human subjects provides encouraging results, wherein researchers found bone formation on implanting artificial graft material with SVF supplement compared to the graft alone [66]. The use of matrices/scaffolds and populating those with SVF and/or ADSCs is a promising area of application, though still in experimental phases [13, 78, 79]. Here, we will not go into much detail regarding the applications as that has been well accomplished in a recent two-part review [3, 26].
“Fat stem cell” therapies and regulatory scenario
Clinics all across the globe began providing “fat stem cell”-based therapy shortly after its discovery, promising miraculous results and more, but often running into controversies [80–86]. Such therapies in the US are known to charge anywhere from USD5000 to USD100,000, and, although mostly harmless and sometimes beneficial, there have been reports of vision loss, tumours, and even deaths [80–86]. Being a major issue in the USA, the FDA had to step in with a draft guideline late in 2014 [87]. These guidelines can be considered in future development of technologies and procedures related to SVF and therapies. Although the “stem cell therapy” genre includes many types of stem cells, ADSCs remain the most marketed variety in the US [88].
The common practices of enzymatic and mechanical disruption of adipose tissue for isolating SVF are explicitly mentioned in the FDA document as “more than minimal manipulation” [87]. As and when the guidelines are implemented, SVF isolated by current protocols (enzymatic digestion) can be treated as a Category 351 product, that is a “drug/biologic” and in need of complete FDA regulation [68]. This calls for exploration of alternate methods, keeping in mind that regulations in the US often trickle down to other geographies, especially in matters of food and drugs.
Introduction of the Reliable and Effective Growth for Regenerative Health Options that Improve Wellness (REGROW) Act [89] in the US Senate last year led to scientific and policy debate, with prominent organisations such as the ISCT, the International Society for Stem Cell Research (ISSCR), and many patient and advocacy groups refusing to support it, at least in its current form [88, 90–94]. The REGROW Act aims to hasten the “conditional approval” of certain cell and tissue therapeutic products which demonstrate “reasonable expectation of effectiveness” along with a few other criteria [89]. However, the use of open-ended terms such as “reasonable expectation of effectiveness” amounts to a lack of clear scientific definition, thus leaving scope for interpretation of the law, consequently leading to potential abuse; such concerns are possibly behind this strong opposition towards the act.
Nevertheless, an urgent consensus is required among all stakeholders with regards to realising the translational potential of stem cells and other cell-based therapeutics, especially when it comes to serious unmet medical needs.
Conclusions
MSCs have been long known for their remarkable properties when it comes to regeneration and therapeutic potential. ADSCs are possibly the easiest to isolate among all the different types of MSCs in an adult human and in relative abundance too; up to 500 times more stem/stromal cells per gram as compared to a bone marrow source [95]. Simply put, ADSCs are potentially the most abundant regenerative cells in the human body and SVF is a step in the protocol to isolate ADSCs. As has been repeatedly mentioned in this review, the potential for use of both SVF and ADSCs in regenerative medicine are immense. However, care must be taken to go about it without harming the intended beneficiary, that is the patients and public in general. Guidelines, such as the ones from US FDA and their counterparts elsewhere will be important parameters in judging new therapies and technologies being developed, and we ought to keep abreast of such issues. Technology development is the single most important factor to realise the full potential of any new therapy, and SVF-based therapy is no exception. At the same time, it is evident that we need a better understanding of SVF and ADSC biology. This is a continuous endeavour and will only help to better establish the core principles and mechanisms of SVF- and ADSC-based therapies. In the process, we are likely to discover newer applications apart from the plethora already identified. Combining these therapies with other technologies such as decellularised or three-dimensional printed scaffolds with the aim of transplantation will jump-start other areas of clinical and commercial developments.
Acknowledgements
We thank Dr. Swathi SundarRaj and Mr. Murali Cherat for critical reading of the manuscript and feedback, and members of Stempeutics, especially Mr. Vasanth Kumar, Ms. Pradnya Shahani, and Ms. Ankita Walvekar along with the rest of the Research & Development division for their help and support during writing of this review.
Funding
Not applicable.
Availability of data and materials
Not applicable.
Authors’ contributions
PB: Conceptualisation, data mining, and writing of manuscript. ASM: Conceptualisation, manuscript review, editing suggestions, and final approval. Both authors read and approved the final manuscript.
Competing interests
PB and ASM are or have been part of Stempeutics Research and were involved in the development of Stempeutron™ as salaried employees of Stempeutics Research Pvt. Ltd.
Consent for publication
The Tables and the Figure are original for this article and the sources used have been cited both within the article and within the Tables and Figure.
Ethics approval and consent to participate
Not applicable.
Publisher’s Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Abbreviations
ADSC | Adipose-derived stem/stromal cell |
BADSC | Brown adipose-derived stem cell |
BAT | Brown adipose tissue |
BMMSC | Bone marrow mesenchymal stromal/stem cell |
CAL | Cell-assisted lipotransfer |
CD | Cluster of differentiation |
EC | Endothelial cell |
EPC | Endothelial precursor cell |
FDA | Food and Drug Administration |
HIF1 | Hypoxia inducible factor 1 |
HSC | Haematopoietic stem cell |
IFATS | International Federation for Adipose Therapeutics and Science |
IL | Interleukin |
ISCT | International Society of Cellular Therapy |
ISSCR | International Society for Stem Cell Research |
MSC | Mesenchymal stem/stromal cell |
PDGF | Platelet-derived growth factor |
REGROW | Reliable and Effective Growth for Regenerative Health Options that Improve Wellness |
SVF | Stromal vascular fraction |
VEGF | Vascular endothelial growth factor |
VLS | Vulvar lichen sclerosus |
WAT | White adipose tissue |
Contributor Information
Pablo Bora, Email: moc.liamtoh@arob.olbap.
Anish S. Majumdar, Phone: +91-80-39992456, Email: moc.scituepmets@radmujam.hsina.
References
1. Bourin P, Bunnell BA, Casteilla L, 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 and Science (IFATS) and the International Society for Cellular Therapy (ISCT) Cytotherapy. 2013;15:641–8. doi: 10.1016/j.jcyt.2013.02.006. [PMC free article][PubMed] [Cross Ref]
2. Gimble JM, Bunnell BA, Frazier T, et al. Adipose-derived stromal/stem cells. Organogenesis. 2013;9:3–10. doi: 10.4161/org.24279. [PMC free article] [PubMed] [Cross Ref]
3. Nguyen A, Guo J, Banyard DA, et al. Stromal vascular fraction: a regenerative reality? Part 1: current concepts and review of the literature. J Plast Reconstr Aesthetic Surg. 2016;69:170–9. doi: 10.1016/j.bjps.2015.10.015. [PubMed] [Cross Ref]
4. Bunnell B, Flaat M, Gagliardi C, et al. Adipose-derived stem cells: isolation, expansion and differentiation. Methods. 2008;45:115–20. doi: 10.1016/j.ymeth.2008.03.006. [PMC free article] [PubMed][Cross Ref]
5. Casteilla L. Adipose-derived stromal cells: their identity and uses in clinical trials, an update. World J Stem Cells. 2011;3:25. doi: 10.4252/wjsc.v3.i4.25. [PMC free article] [PubMed] [Cross Ref]
6. Suzuki E, Fujita D, Takahashi M, et al. Adipose tissue-derived stem cells as a therapeutic tool for cardiovascular disease. World J Cardiol. 2015;7:454–65. doi: 10.4330/wjc.v7.i11.707. [PMC free article][PubMed] [Cross Ref]
7. Bony C, Cren M, Domergue S, et al. Adipose mesenchymal stem cells isolated after manual or water-jet-assisted liposuction display similar properties. Front Immunol. 2016;6:1–8. doi: 10.3389/fimmu.2015.00655. [PMC free article] [PubMed] [Cross Ref]
8. Mi HM, Sun YK, Yeon JK, et al. Human adipose tissue-derived mesenchymal stem cells improve postnatal neovascularization in a mouse model of hindlimb ischemia. Cell Physiol Biochem. 2006;17:279–90. doi: 10.1159/000094140. [PubMed] [Cross Ref]
9. van Dijk A, Naaijkens BA, Jurgens WJFM, et al. Reduction of infarct size by intravenous injection of uncultured adipose derived stromal cells in a rat model is dependent on the time point of application. Stem Cell Res. 2011;7:219–29. doi: 10.1016/j.scr.2011.06.003. [PubMed] [Cross Ref]
10. Charles-de-Sá L, Gontijo-de-Amorim NF, Maeda Takiya C, et al. Antiaging treatment of the facial skin by fat graft and adipose-derived stem cells. Plast Reconstr Surg. 2015;135:999–1009. doi: 10.1097/PRS.0000000000001123. [PubMed] [Cross Ref]
11. Semon JA, Zhang X, Pandey AC, et al. Administration of murine stromal vascular fraction ameliorates chronic experimental autoimmune encephalomyelitis. Stem Cells Transl Med. 2013;2:789–96. doi: 10.5966/sctm.2013-0032. [PMC free article] [PubMed] [Cross Ref]
12. You D, Jang MJ, Kim BH, et al. Comparative study of autologous stromal vascular fraction and adipose-derived stem cells for erectile function recovery in a rat model of cavernous nerve injury. Stem Cells Transl Med. 2015;4:351–8. doi: 10.5966/sctm.2014-0161. [PMC free article] [PubMed] [Cross Ref]
13. Mohammadi R, Sanaei N, Ahsan S, et al. Stromal vascular fraction combined with silicone rubber chamber improves sciatic nerve regeneration in diabetes. Chinese J Traumatol. 2015;18:212–8. doi: 10.1016/j.cjtee.2014.10.005. [PubMed] [Cross Ref]
14. Yoshimura K, Sato K, Aoi N, et al. Cell-assisted lipotransfer for cosmetic breast augmentation: supportive use of adipose-derived stem/stromal cells. Aesthetic Plast Surg. 2008;32:48–55. doi: 10.1007/s00266-007-9019-4. [PMC free article] [PubMed] [Cross Ref]
15. Pak J, Chang J-J, Lee JH, et al. Safety reporting on implantation of autologous adipose tissue-derived stem cells with platelet-rich plasma into human articular joints. BMC Musculoskelet Disord. 2013;14:337. doi: 10.1186/1471-2474-14-337. [PMC free article] [PubMed] [Cross Ref]
16. Yoshimura K, Suga H, Eto H. Adipose-derived stem/progenitor cells: roles in adipose tissue remodeling and potential use for soft tissue augmentation. Regen Med. 2009;4:265–73. doi: 10.2217/17460751.4.2.265. [PubMed] [Cross Ref]
17. Matsumoto D, Sato K, Gonda K, et al. Cell-assisted lipotransfer: supportive use of human adipose-derived cells for soft tissue augmentation with lipoinjection. Tissue Eng. 2006;12:3375–82. doi: 10.1089/ten.2006.12.3375. [PubMed] [Cross Ref]
18. Silva FJ, Holt DJ, Vargas V, et al. Metabolically active human brown adipose tissue derived stem cells. Stem Cells. 2014;32:572–81. doi: 10.1002/stem.1595. [PubMed] [Cross Ref]
19. Gimble JM, Katz AJ, Bunnell BA. Adipose-derived stem cells for regenerative medicine. Circ Res. 2007;100:1249–60. doi: 10.1161/01.RES.0000265074.83288.09. [PMC free article] [PubMed] [Cross Ref]
20. Colen BD. A pill to shed fat? Harvard stem cell researchers say they finally can turn “bad” fat into “good.” Harvard Gaz 2014. http://news.harvard.edu/gazette/story/2014/12/a-pill-to-shed-fat/. Accessed 8 Sept 2016.
21. Moisan A, Lee Y-K, Zhang JD, et al. White-to-brown metabolic conversion of human adipocytes by JAK inhibition. Nat Cell Biol. 2014;17:57–67. doi: 10.1038/ncb3075. [PMC free article] [PubMed][Cross Ref]
22. Oberbauer E, Steffenhagen C, Wurzer C, et al. Enzymatic and non-enzymatic isolation systems for adipose tissue-derived cells: current state of the art. Cell Regen. 2015;4:7. doi: 10.1186/s13619-015-0020-0. [PMC free article] [PubMed] [Cross Ref]
23. Tonnard P, Verpaele A, Peeters G, et al. Nanofat grafting: basic research and clinical applications. Plast Reconstr Surg. 2013;132:1017–26. doi: 10.1097/PRS.0b013e31829fe1b0. [PubMed] [Cross Ref]
24. Aronowitz JA, Lockhart RA, Hakakian CS. Mechanical versus enzymatic isolation of stromal vascular fraction cells from adipose tissue. Springerplus. 2015;4:713. doi: 10.1186/s40064-015-1509-2.[PMC free article] [PubMed] [Cross Ref]
25. Shah FS, Wu X, Dietrich M, et al. A non-enzymatic method for isolating human adipose tissue-derived stromal stem cells. Cytotherapy. 2013;15:979–85. doi: 10.1016/j.jcyt.2013.04.001. [PubMed] [Cross Ref]
26. Guo J, Nguyen A, Banyard DA, et al. Stromal vascular fraction: a regenerative reality? Part 2: mechanisms of regenerative action. J Plast Reconstr Aesthetic Surg. 2016;69:180–8. doi: 10.1016/j.bjps.2015.10.014. [PubMed] [Cross Ref]
27. Dominici M, Le Blanc K, Mueller I, et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy. 2006;8:315–7. doi: 10.1080/14653240600855905. [PubMed] [Cross Ref]
28. Zuk PA, Zhu M, Mizuno H, et al. Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng. 2001;7:211–28. doi: 10.1089/107632701300062859. [PubMed] [Cross Ref]
29. SundarRaj S, Deshmukh A, Priya N, et al. Development of a system and method for automated isolation of stromal vascular fraction from adipose tissue lipoaspirate. Stem Cells Int. 2015;2015:1–11. doi: 10.1155/2015/109353. [PMC free article] [PubMed] [Cross Ref]
30. Riis S, Zachar V, Boucher S, et al. Critical steps in the isolation and expansion of adipose-derived stem cells for translational therapy. Expert Rev Mol Med. 2015;17:e11. doi: 10.1017/erm.2015.10. [PubMed][Cross Ref]
31. Tamburino S, Lombardo GA, Tarico MS, et al. The role of nanofat grafting in vulvar lichen sclerosus: a preliminary report. Arch Plast Surg. 2016;43:93. doi: 10.5999/aps.2016.43.1.93. [PMC free article][PubMed] [Cross Ref]
32. KemaloÄŸlu CA. Nanofat grafting under a split-thickness skin graft for problematic wound management. Springerplus. 2016;5:138. doi: 10.1186/s40064-016-1808-2. [PMC free article] [PubMed] [Cross Ref]
33. Fraser JK, Hicok KC, Shanahan R, et al. The Celution(®) system: automated processing of adipose-derived regenerative cells in a functionally closed system. Adv Wound Care. 2014;3:38–45. doi: 10.1089/wound.2012.0408. [PMC free article] [PubMed] [Cross Ref]
34. Invitrogen™ Countess™. Invitrogen cell data sheet-ADSC. http://www.thermofisher.com/content/dam/LifeTech/migration/en/filelibrary/cell-tissue-analysis/pdfs.par.71179.file.dat/co13964-stem-cell-data-sheet-adsc.pdf. Accessed 7 Sept 2016.
35. Sponsored paper. Rapid analysis of human adipose-derived stem cells and 3 T3-L1 differentiation toward adipocytes using the Scepter™ 2.0 cell counter. Biotechniques. 2012;53:109–11.
36. Ryu YJ, Cho TJ, Lee DS, et al. Phenotypic characterization and in vivo localization of human adipose-derived mesenchymal stem cells. Mol Cells. 2013;35:557–64. doi: 10.1007/s10059-013-0112-z.[PMC free article] [PubMed] [Cross Ref]
37. Asahara T, Kawamoto A, Masuda H. Concise review: circulating endothelial progenitor cells for vascular medicine. Stem Cells. 2011;29:1650–5. doi: 10.1002/stem.745. [PubMed] [Cross Ref]
38. Garipcan B, Maenz S, Pham T, et al. Image analysis of endothelial microstructure and endothelial cell dimensions of human arteries—a preliminary study. Adv Eng Mater. 2011;13:B54–7. doi: 10.1002/adem.201080076. [Cross Ref]
39. Bergman RA, Afifi AK, Heidger PM. Anatomy atlases: atlas of microscopic anatomy: section 4—blood. Plate 4.53: lymphocytes. http://www.anatomyatlases.org/MicroscopicAnatomy/Section04/Plate0453.shtml. Accessed 7 Sept 2016.
40. Rosenbluth MJ, Lam WA, Fletcher DA. Force microscopy of nonadherent cells: a comparison of leukemia cell deformability. Biophys J. 2006;90:2994–3003. doi: 10.1529/biophysj.105.067496.[PMC free article] [PubMed] [Cross Ref]
41. Krombach F, Münzing S, Allmeling AM, et al. Cell size of alveolar macrophages: an interspecies comparison. Environ Health Perspect. 1997;105:1261–3. doi: 10.1289/ehp.97105s51261.[PMC free article] [PubMed] [Cross Ref]
42. Duke University Medical School. Histology learning resources. https://web.duke.edu/histology/MoleculesCells/Muscle/Muscle.html#webslide96. Accessed 7 Sept 2016.
43. Duke University Medical School. Histology learning resources. https://web.duke.edu/histology/MoleculesCells/Muscle/muscle.jpg. Accessed 7 Sept 2016.
44. Proebstl D, Voisin M-B, Woodfin A, et al. Pericytes support neutrophil subendothelial cell crawling and breaching of venular walls in vivo. J Exp Med. 2012;209:1219–34. doi: 10.1084/jem.20111622.[PMC free article] [PubMed] [Cross Ref]
45. Lai N, Sims JK, Jeon NL, et al. Adipocyte induction of preadipocyte differentiation in a gradient chamber. Tissue Eng Part C Methods. 2012;18:958–67. doi: 10.1089/ten.tec.2012.0168. [PMC free article][PubMed] [Cross Ref]
46. Traktuev DO, Merfeld-Clauss S, Li J, et al. A population of multipotent CD34-positive adipose stromal cells share pericyte and mesenchymal surface markers, reside in a periendothelial location, and stabilize endothelial networks. Circ Res. 2008;102:77–85. doi: 10.1161/CIRCRESAHA.107.159475. [PubMed][Cross Ref]
47. Traktuev DO, Prater DN, Merfeld-Clauss S, et al. Robust functional vascular network formation in vivo by cooperation of adipose progenitor and endothelial cells. Circ Res. 2009;104:1410–20. doi: 10.1161/CIRCRESAHA.108.190926. [PubMed] [Cross Ref]
48. Pallua N, Serin M, Wolter TP. Characterisation of angiogenetic growth factor production in adipose tissue-derived mesenchymal cells. J Plast Surg Hand Surg. 2014;48:412–6. doi: 10.3109/2000656X.2014.903196. [PubMed] [Cross Ref]
49. Grasys J, Kim B, Pallua N. Content of soluble factors and characteristics of stromal vascular fraction cells in lipoaspirates from different subcutaneous adipose tissue depots. Aesthetic Surg J. 2016;36:831–41. doi: 10.1093/asj/sjw022. [PubMed] [Cross Ref]
50. Szöke K, Brinchmann JE. Concise review: therapeutic potential of adipose tissue-derived angiogenic cells. Stem Cells Transl Med. 2012;1:658–67. doi: 10.5966/sctm.2012-0069. [PMC free article] [PubMed][Cross Ref]
51. Crisan M, Yap S, Casteilla L, et al. A perivascular origin for mesenchymal stem cells in multiple human organs. Cell Stem Cell. 2008;3:301–13. doi: 10.1016/j.stem.2008.07.003. [PubMed] [Cross Ref]
52. Maumus M, Peyrafitte J-A, D’Angelo R, et al. Native human adipose stromal cells: localization, morphology and phenotype. Int J Obes. 2011;35:1141–53. doi: 10.1038/ijo.2010.269. [PMC free article][PubMed] [Cross Ref]
53. Lin C-S, Ning H, Lin G, et al. Is CD34 truly a negative marker for mesenchymal stromal cells? Cytotherapy. 2012;14:1159–63. doi: 10.3109/14653249.2012.729817. [PMC free article] [PubMed][Cross Ref]
54. Suga H, Matsumoto D, Eto H, et al. Functional implications of CD34 expression in human adipose–derived stem/progenitor cells. Stem Cells Dev. 2009;18:1201–10. doi: 10.1089/scd.2009.0003. [PubMed][Cross Ref]
55. Sidney LE, Branch MJ, Dunphy SE, et al. Concise review: evidence for CD34 as a common marker for diverse progenitors. Stem Cells. 2014;32:1380–9. doi: 10.1002/stem.1661. [PMC free article] [PubMed][Cross Ref]
56. 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;5:1–8. doi: 10.4252/wjsc.v5.i1.1.[PMC free article] [PubMed] [Cross Ref]
57. Keith B, Simon MC. Hypoxia-inducible factors, stem cells, and cancer. Cell. 2007;129:465–72. doi: 10.1016/j.cell.2007.04.019. [PMC free article] [PubMed] [Cross Ref]
58. Das R, Jahr H, van Osch GJVM, et al. The role of hypoxia in bone marrow-derived mesenchymal stem cells: considerations for regenerative medicine approaches. Tissue Eng Part B Rev. 2010;16:159–68. doi: 10.1089/ten.teb.2009.0296. [PubMed] [Cross Ref]
59. Ejtehadifar M, Shamsasenjan K, Movassaghpour A, et al. The effect of hypoxia on mesenchymal stem cell biology. Adv Pharm Bull. 2015;5:141–9. doi: 10.15171/apb.2015.021. [PMC free article] [PubMed][Cross Ref]
60. Beegle J, Lakatos K, Kalomoiris S, et al. Hypoxic preconditioning of mesenchymal stromal cells induces metabolic changes, enhances survival, and promotes cell retention in vivo. Stem Cells. 2015;33:1818–28. doi: 10.1002/stem.1976. [PubMed] [Cross Ref]
61. Haque N, Rahman MT, Abu Kasim NH, et al. Hypoxic culture conditions as a solution for mesenchymal stem cell based regenerative therapy. Sci World J. 2013;2013:1–12. doi: 10.1155/2013/632972. [PMC free article] [PubMed] [Cross Ref]
62. Grayson WL, Zhao F, Bunnell B, et al. Hypoxia enhances proliferation and tissue formation of human mesenchymal stem cells. Biochem Biophys Res Commun. 2007;358:948–53. doi: 10.1016/j.bbrc.2007.05.054. [PubMed] [Cross Ref]
63. Feng Y, Zhu M, Dangelmajer S, et al. Hypoxia-cultured human adipose-derived mesenchymal stem cells are non-oncogenic and have enhanced viability, motility, and tropism to brain cancer. Cell Death Dis. 2014;5:e1567. doi: 10.1038/cddis.2014.521. [PMC free article] [PubMed] [Cross Ref]
64. Brunet De La Grange P, Barthe C, Lippert E, et al. Oxygen concentration influences mRNA processing and expression of the CD34 gene. J Cell Biochem. 2006;97:135–44. doi: 10.1002/jcb.20597. [PubMed][Cross Ref]
65. Rigotti G, Marchi A, Galie M, et al. Clinical treatment of radiotherapy tissue damage by lipoaspirate transplant: a healing process mediated by adipose-derived adult stem cells. Plast Reconstr Surg. 2007;119:1409–22. doi: 10.1097/01.prs.0000256047.47909.71. [PubMed] [Cross Ref]
66. Prins H-J, Schulten EAJM, ten Bruggenkate CM, et al. Bone regeneration using the freshly isolated autologous stromal vascular fraction of adipose tissue in combination with calcium phosphate ceramics. Stem Cells Transl Med. 2016;5:1362–74. doi: 10.5966/sctm.2015-0369. [PMC free article] [PubMed][Cross Ref]
67. Haahr MK, Jensen CH, Toyserkani NM, et al. Safety and potential effect of a single intracavernous injection of autologous adipose-derived regenerative cells in patients with erectile dysfunction following radical prostatectomy: an open-label phase I clinical trial. EBioMedicine. 2016;5:204–10. doi: 10.1016/j.ebiom.2016.01.024. [PMC free article] [PubMed] [Cross Ref]
68. Tocco I, Widgerow AD, Lalezari S, et al. Lipotransfer: the potential from bench to bedside. Ann Plast Surg. 2014;72:599–609. doi: 10.1097/SAP.0000000000000154. [PubMed] [Cross Ref]
69. Doi K, Tanaka S, Iida H, et al. Stromal vascular fraction isolated from lipo-aspirates using an automated processing system: bench and bed analysis. J Tissue Eng Regen Med. 2013;7:864–70. doi: 10.1002/term.1478. [PubMed] [Cross Ref]
70. Rigotti G, Charles-de-Sá L, Gontijo-de-Amorim NF, et al. Expanded stem cells, stromal-vascular fraction, and platelet-rich plasma enriched fat: comparing results of different facial rejuvenation approaches in a clinical trial. Aesthet Surg J. 2016;36:261–70. doi: 10.1093/asj/sjv231. [PMC free article][PubMed] [Cross Ref]
71. Jackson WM, Nesti LJ, Tuan RS. Concise review: clinical translation of wound healing therapies based on mesenchymal stem cells. Stem Cells Transl Med. 2012;1:44–50. doi: 10.5966/sctm.2011-0024.[PMC free article] [PubMed] [Cross Ref]
72. Borowski DW, Gill TS, Agarwal AK, et al. Autologous adipose-tissue derived regenerative cells for the treatment of complex cryptoglandular fistula-in-ano: a report of three cases. BMJ Case Rep. 2012;2012:4–7. doi: 10.1136/bcr-2012-006988. [PMC free article] [PubMed] [Cross Ref]
73. Riordan NH, Ichim TE, Min W-P, et al. Non-expanded adipose stromal vascular fraction cell therapy for multiple sclerosis. J Transl Med. 2009;7:29. doi: 10.1186/1479-5876-7-29. [PMC free article][PubMed] [Cross Ref]
74. Lee HC, An SG, Lee HW, et al. Safety and effect of adipose tissue-derived stem cell implantation in patients with critical limb ischemia. Circ J. 2012;76:1750–60. doi: 10.1253/circj.CJ-11-1135. [PubMed][Cross Ref]
75. Parcero JJ, Perez JA, Patel AN, et al. Autologous adipose-derived stromal stem cell implantation to resolve critical limb ischemia: case report. Cureus. 2014;6(5):e182. doi:10.7759/cureus.182. http://www.cureus.com/articles/2376-autologous-adipose-derived-stromal-stem-cell-implantation-to-resolve-critical-limb-ischemia-case-report.
76. Pers Y-M, Rackwitz L, Ferreira R, et al. Adipose mesenchymal stromal cell-based therapy for severe osteoarthritis of the knee: a phase I dose-escalation trial. Stem Cells Transl Med. 2016;5:847–56. doi: 10.5966/sctm.2015-0245. [PMC free article] [PubMed] [Cross Ref]
77. Domergue S, Bony C, Maumus M, et al. Comparison between stromal vascular fraction and adipose mesenchymal stem cells in remodeling hypertrophic scars. PLoS One. 2016;11:e0156161. doi: 10.1371/journal.pone.0156161. [PMC free article] [PubMed] [Cross Ref]
78. Frazier TP, Bowles A, Lee S, et al. Serially transplanted nonpericytic CD146– adipose stromal/stem cells in silk bioscaffolds regenerate adipose tissue in vivo. Stem Cells. 2016;34:1097–111. doi: 10.1002/stem.2325. [PubMed] [Cross Ref]
79. Lin S-D, Huang S-H, Lin Y-N, et al. Injected implant of uncultured stromal vascular fraction loaded onto a collagen gel. Ann Plast Surg. 2016;76:S108–16. doi: 10.1097/SAP.0000000000000687. [PubMed][Cross Ref]
80. NEWSmax. Unregulated stem cell industry is “wild west.” NEWSmax; 2015. http://www.newsmax.com/Health/Health-News/stem-cells-treatments-regulation/2015/05/18/id/645186/. Accessed 7 Sept 2016.
81. McFarling UL. FDA moves to crack down on unproven stem cell therapies. STAT News. 2016. https://www.statnews.com/2016/02/08/fda-crackdown-stem-cell-clinics/. Accessed 7 Sept 2016.
82. Jabr F. In the flesh: the embedded dangers of untested stem cell cosmetics. Sci Am. 2012. Available at http://www.scientificamerican.com/article/stem-cell-cosmetics/. Accessed 7 Sept 2016.
83. Wilson C. Stem cell treatment causes nasal growth in woman’s back. New Sci. 2014. https://www.newscientist.com/article/dn25859-stem-cell-treatment-causes-nasal-growth-in-womans-back/. Accessed 7 Sept 2016.
84. Cyranoski D. Korean deaths spark inquiry. Nature. 2010;468:485. doi: 10.1038/468485a. [PubMed][Cross Ref]
85. McLean AK, Stewart C, Kerridge I. Untested, unproven, and unethical: the promotion and provision of autologous stem cell therapies in Australia. Stem Cell Res Ther. 2015;6:12. doi: 10.1186/s13287-015-0047-8. [PMC free article] [PubMed] [Cross Ref]
86. Ledford H. Boom in unproven cell therapies intensifies regulatory debate. Nature. 2016;537:148. doi: 10.1038/537148a. [PubMed] [Cross Ref]
87. US Department of Health and Human Services (Food and Drug Administration). Human cells, tissues, and cellular- and tissue-based products (HCT/Ps) from adipose tissue : regulatory considerations; draft guidance. http://www.fda.gov/BiologicsBloodVaccines/GuidanceComplianceRegulatoryInformation/Guidances/Tissue/ucm427795.htm#HCT_QUESTION. Accessed 7 Sept 2016.
88. Turner L, Knoepfler P. Selling stem cells in the USA: assessing the direct-to-consumer industry. Cell Stem Cell. 2016;19:154–7. doi: 10.1016/j.stem.2016.06.007. [PubMed] [Cross Ref]
89. Kirk MS, Manchin J, Collins SM. REGROW Act. Congress.gov 2016. https://www.congress.gov/bill/114th-congress/senate-bill/2689/cosponsors. Accessed 7 Sept 2016.
90. Research MJFF for P. MJFF signs letter opposing the REGROW Act 2016. https://www.michaeljfox.org/foundation/news-detail.php?mjff-signs-letter-opposing-the-regrow-act. Accessed 7 Sept 2016.
91. International Society for Cellular Therapy. ISCT calls for changes to proposed US REGROW Act on cell therapies 2016. http://www.celltherapysociety.org/news/news.asp?id=304128&hhSearchTerms=%22regrow%22. Accessed 7 Sept 2016.
92. The Alliance for Regenerative Medicine. ARM statement in response to U.S. Senator Kirk’s REGROW Act. http://alliancerm.org/sites/default/files/ARMSenatorKirk_REGROWActletter_March2016_.pdf. Accessed 7 Sept 2016.
93. Knoepfler P. California stem-cell institute’s political gamble. San Fr Chron 2016. http://www.sfchronicle.com/opinion/article/California-stem-cell-institute-s-political-8250137.php?t=4c01ff1973cefdcb88&cmpid=twitter-premium. Accessed 7 Sept 2016.
94. Joseph A. Drive to get more patients experimental stem cell treatments stirs concern. STAT News 2016. https://www.statnews.com/2016/06/30/stem-cell-political-fight. Accessed 7 Sept 2016.
95. Hass R, Kasper C, Böhm S, et al. Different populations and sources of human mesenchymal stem cells (MSC): a comparison of adult and neonatal tissue-derived MSC. Cell Commun Signal. 2011;9:12. doi: 10.1186/1478-811X-9-12. [PMC free article] [PubMed] [Cross Ref]
Springerplus. 2015; 4: 713.
Published online 2015 Nov 23. doi: 10.1186/s40064-015-1509-2
PMCID: PMC4656256
Mechanical versus enzymatic isolation of stromal vascular fraction cells from adipose tissue
Copyright © Aronowitz et al. 2015
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.
This article has been cited by other articles in PMC.
Background
The clinical use of autologous adipose-derived stem cells (ASCs) is rapidly expanding because of promising results across a wide range of conditions. While progress in the use of cultured, modified and induced pluripotential cells has been measured in laboratory milestones, the use of autologous adipose-derived pluripotent cells is burgeoning at the clinical level. Clinical and pre-clinical studies show that autogenous ASCs demonstrably survive after transplantation, show pluripotential differentiation (Zuk et al. 2001; Planat-Benard et al. 2004; Naderi et al. 2014; Ude et al. 2014) and exhibit anti-apoptotic, anti-inflammatory, and angiogenic effects (Rehmam et al. 2004; Kapur and Katz 2013; Suga et al. 2010; Eto et al. 2012; Kato et al. 2014).
Applications as diverse as myocardial infarction, cosmetic surgery, osteoarthritis and bone regeneration, inflammatory bowel disease and chronic wounds are reported among a myriad of others (Savi et al. 2015; Matsumoto et al. 2006; Di Rocco et al. 2010; Asatrian et al. 2015; Nagaishi et al. 2015). There is some variation in the number of stem cells present in various donor sites and with donor age (Jurgens et al. 2008; Vilaboa et al. 2014; Buschmann et al. 2013). In general, the most efficient methods 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. In general, because no additional adverse effects are reported with the use of autologous ASCs in fat grafting, the largest number of cells isolated at the point of care without expansion in culture is typically used. Despite a lack of reported clinical risk, in vitro studies have demonstrated potential oncological risks which clinicians should be cautious of when using SVF based therapies (Bertolini et al. 2012; Bielli et al. 2014).
The surge in clinical applications for ASCs increases the need for clear and reliable information about the efficiency, cost and safety of automated equipment and manual techniques which facilitate separation of the stromal vascular fraction (SVF) from adipose tissue. In clinical practice, adipose-derived stem cells are often not administered as a pure isolate but rather as one constituent of stromal vascular fraction, a heterogeneous mixture of cells resulting from the mechanical or enzymatic processing of aspirated adipose tissue. SVF contains a variety of cells including macrophages, various blood cells, pericytes, fibroblasts, smooth muscle cells, vascular endothelial progenitors and adipose-derived stem cells (Yoshimura et al. 2006; Bourin et al. 2013; Han et al. 2010; McIntosh et al. 2006; Bonab et al. 2006; Yoshimura et al. 2009). Stromal vascular fraction is one component of the heterogeneous mixture of adipose tissue fragments, stromal tissue, blood and tumescent fluid which constitutes lipoaspirate. The ASC content of SVF varies substantially depending on the method employed, with reports from less than 1 % of cells to over 15 % (Table 1). SVF cells can be safely isolated, quantified and characterized at the point of care in approximately 90 min. This is a timeframe which permits isolation and treatment to occur in the same surgical procedure, that is, at the point of care (POC).
Enzymatic methods
Enzymatic methods of isolating SVF cells from adipose tissue at the POC are based on a commonly used laboratory method of obtaining stem cells. The methods used to manually process adipose tissue using collagenase follow the same basic steps, but vary slightly in technique and reagents used. Lipoaspirate is washed 2–3 times using an aqueous salt solution such as PBS, Lactated Ringer’s solution, or Hank’s Balanced Salt Solution (HBSS). The washed lipoaspirate is then incubated with a collagenase solution of variable concentration and composition, depending on the method and tissue dissociation enzyme product used. Enzymatic digestion is typically carried out in a heated shaker to provide constant agitation at 37 °C for 30 min to 2 h. The digested adipose tissue is then centrifuged (speed/duration vary. See Table 1) which separates the processed lipoaspirate into three main layers, the oil/adipose tissue layer, the aqueous layer, and the pellet. The SVF is contained within the pellet, so the other layers are discarded, although SVF cells can be recovered from the aqueous layer (Yoshimura et al. 2006). The pellet is washed to remove any residual enzyme and filtered to remove tissue fragments and detritus. Collagenase-based enzymatic methods can be up to 1000 times more effective in SVF cell recovery than mechanical methods. Enzymatic methods are more efficient in isolating SVF cells because disruption of the collagen-based extracellular matrix (ECM) which binds together adipocytes and other cells of adipose tissue.
Tissue dissociation enzyme mixtures used for the separation process are usually a mixture of type I and type II collagenases isolated from Clostridium histolyticum, and various other proteolytic enzymes such as neutral protease (Dispase) (Fogarty and Griffin 1973; Griffin and Fogarty 1973) isolated from P. polymyxaor thermolysin (Ke et al. 2013) isolated from G. stearothermophilus or B. thermoproteolyticus, depending on the product used. Commonly enzymatic methods are carried out using tissue dissociation enzyme mixtures such as CIzyme™ AS (Vitacyte LLC, Indianapolis, Indiana) or Liberase™ Research Grade (Roche Diagnostics, Basel, Switzerland). CIzyme™ AS is a mixture of type I and type II clostridial collagenase and dispase. The Liberase™ Research Grade enzyme mixture recommended for adipose-tissue digestion is mixture of type I and type II clostridial collagenase and thermolysin. Mixtures of enzymes have been shown to yield more nucleated cells than using only one enzyme, a quality attributed to the synergistic effect of the proteolytic enzymes in the breakdown of the ECM (McCarthy et al. 2010, 2011; Breite et al. 2010); however collagenase is still frequently used as the sole proteolytic enzyme in methods using products such as Collagenase NB6 (SERVA Electrophoresis GmbH, Heidelberg, Germany) or Collagenase type I CLS 270 (Worthington Biochemical Corporation, Lakewood, NJ).
Published yields of viable, nucleated SVF cells achieved using manual, collagenase-based digestions range from 100,000 nucleated cells/cc to 1,300,000 nucleated cells/cc of lipoaspirate processed (Table 1). Equipment like the PNC Multi-Station (PNC International, Gyeonggi-do, Republic of Korea) is commercially available for use in the manual preparation of SVF. The PNC Multi-Station contains a centrifuge and heated shaker inside of a sterile biohood which allows the entire processing to be conducted in sterile conditions.
Mechanical isolation methods
Mechanical methods for SVF isolation report 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 (Table 1). Mechanical methods seek alternative non-enzymatic means of removing SVF cells from the adipose tissue and tend to be focused around washing and shaking/vibrating lipoaspirate followed by centrifugation in order to concentrate the SVF cells. All of the mechanical methods mentioned in this article contain a centrifugation step in order to concentrate the SVF cells. The composition of the cell populations recovered through simple centrifugation and other non-enzymatic methods have been shown to contain a greater frequency of peripheral blood mononuclear cells and a substantially lower number of progenitor cells (Conde-Green et al. 2014; Raposio et al. 2014; Shah et al. 2013). This is because ASCs are concentrated in the small and medium sized vascular structures of adipose tissue, and without enzymatic lysis of the collagen-based extracellular matrix many progenitor cells remain trapped within the vascular endothelium layers and connective tissue fragments in the lipoaspirate.
While enzymatic methods consistently yield higher cell counts with a higher frequency of progenitor cells, mechanical methods do offer some distinct advantages. The digestion of adipose tissue to disperse the cellular constituents prolongs the isolation time and can be fairly expensive, with costs of $2–$5 per gram of tissue processed using GMP grade enzymes (Aronowitz and Ellenhorn 2013). In settings where maximum numbers of progenitor cells are not critical, a non-enzymatic separation method like that of Raposio et al. can provide a cost-effective alternative (Raposio et al. 2014). Additionally, mechanical methods tend to offer a faster processing time, some less than 15 min, because they do not require the extra 30–120 min allotted for enzymatic digestion to occur.
Mechanical vs enzymatic methods
In 2014, Raposio et al. reported a non-enzymatic method for SVF isolation (Raposio et al. 2014). This method involves shaking lipoaspirate in a vibrating shaker for 6 min at 600 vibrations per minute and then centrifuging at 1600 rpm for 6 min to isolate the SVF cells. Raposio et al. reported that they were able to isolate around 125,000 nucleated cells per cc of lipoaspirate processed, however only about 5 % of these cells were progenitor cells, with the other 95 % being predominantly blood cells and endothelial cells. In comparison, enzymatic methods have reported SVF yields with significantly higher numbers of progenitor cells, for example one automated collagenase-based isolation system which was shown to yield over 15 % progenitor cells in the SVF (Aronowitz and Ellenhorn 2013). The discrepancy in SVF composition was supported by the paper by Conde-Green et al. (2014). Conde-Green et al. compared a standard collagenase-based method to two different mechanical methods. They reported that both mechanical methods yielded SVF populations with lower nucleated cell counts and lower frequencies of progenitor cells than the manual, enzymatic approach examined.
In 2014, Markarian et al. compared a variety of processing methods for SVF isolation side by side, both enzymatic and mechanical. Collagenase-based digestion was shown to be the most effective in terms of cell recovery Markarian et al. (2014). They reported about 350,000 nucleated cells/cc of lipoaspirate processed using a collagenase-based method. Another method examined was a non-enzymatic method involving centrifugation of lipoaspirate at either 800 g or 1280 g. At both speeds tested, far fewer nucleated cells were isolated, with only about 10,000 nucleated cells recovered per cc of lipoaspirate. They report no significant difference in viability between the various methods they examined.
In 2009, Baptista et al. reported another manual, mechanical method (Baptista et al. 2009). In this method, lipoaspirate is incubated with red blood cell (RBC) lysis buffer (150 mM NH4Cl, 10 mM KHCO3, 1 mM EDTA) at 37 °C for 15 min and then centrifuged for 15 min at 900g. They reported an average yield of about 240,000 nucleated cells per cc of lipoaspirate processed, but only about 12,000 of these (5 %) were adipose-derived stem cells. This was supported by Shah et al. (2013). They compared a similar method using PBS instead of RBC lysis buffer with the common collagenase-based method. Shah et al. cultured samples from each method to determine ASC content. They reported that once samples reached 80–90 % confluence that an average of 25,000 adipose-derived stem cells per cc of lipoaspirate processed were found in the sample acquired using this mechanical method, but 480,000 adipose-derived stem cells per cc of lipoaspirate we found in the sample acquired using the enzymatic method. Additionally, Shah et al. observed that the cells acquired using collagenase proliferated much more quickly when cultured, requiring less than half the time to reach 80–90 % confluence (6 days vs 13 days). This method using RBC lysis buffer was also tested by Markarian et al. (2014). They however reported a much lower yield, only about 25,000 viable cells/cc lipoaspirate processed.
The differences resulting in the yields observed using mechanical and enzymatic methods can be partially attributed to the physical location of SVF cells in adipose tissue. The SVF cells, particularly the mesenchymal stem cells and pericytes, tend to be localized in the perivascular space (Baer and Geiger 2012). As demonstrated by Zimmerlin et al. in 2010, immunohistochemical and immunofluorescent analysis reveal a localization of ASC and pericytes in these perivascular niches (Zimmerlin et al. 2010). Mechanical methods of isolation do not afford the same release of cells from the perivascular spaces because the disruption of the extracellular matrix is significantly reduced compared to enzymatic methods, leaving many of the desired cells trapped in larger tissue fragments which are subsequently discarded. As a result, the composition of the SVF resulting from mechanical isolations tends to be deficient in CD34 expression. This relative CD34+ progenitor deficiency has been suggested as a contributing factor to longer culture times required to reach 80–90 % confluence, as demonstrated by Shah et al. (2013).
Automated/semi-automated devices for SVF isolation
Due to increasing interest of SVF cells in the clinical setting, various fully automated and semi-automated devices for SVF cell isolation, both enzymatically and mechanically based, have been developed by companies hoping to capitalize on this relatively new cellular technology. These devices employ similar methods to manual enzymatic and mechanical methods, but under more controlled conditions. In efforts to improve the yield of SVF isolation, many companies have developed processing systems which seek to optimize the isolation process by reducing the human element and limiting loss of viability due to processing, while still adhering to the current Good Manufacturing Practices (cGMP) (FDA 2014a). Some of these devices have been able to isolate large numbers of cells, while other devices have been shown to be less impressive. These companies continue to improve the devices and technology so as to optimize the cellular recovery and viability. While many of the automated systems are currently too expensive for use in the lab setting, it is very possible that these automated systems could become a common item used to provide safe and effective cellular therapies to patients in the clinical setting. Many of these companies are actively pursuing clinical trials in order to clinically validate their devices and technologies while also providing cellular therapy to patients in need, like the upcoming STAR trial (Cytori Therapeutics 2015) for treatment of scleroderma by Cytori Therapeutics, Inc. which has received an IDE from the FDA and is currently enrolling patients as of August 2015.
These automated and semi-automated systems tend to be small self-contained systems which are able to carry out each step of the process with little or no interference from a technician. One of the main benefits offered by many of these systems is increased sterility through the use of a closed system. Once the lipoaspirate is added to the device, it remains in a sterile environment, unlike many manual methods. In some devices, such as the GID SVF platform (GID Europe, London, UK), the lipoaspirate is harvested directly into the system (Vilaboa et al. 2014). These devices are all slightly different, but ultimately seek to achieve the same goal.
The Cytori Celution system (Cytori Therapeutics, Inc., San Diego, CA) has been reported in multiple studies. The Celution system is a closed, fully automated system which employs Cytori’s proprietary enzyme blend, Celase. The Celution system is capable of processing up to 360 cc of lipoaspirate at one time. The Celution system has been consistently reported to yield between 240,000–360,000 nucleated cells/cc of lipoaspirate processed and 84–93 % viability, while also yielding a large population of progenitors (Table 1) (Aronowitz and Ellenhorn 2013; Lin et al. 2008; Fraser et al. 2013). The Celution system has been reported for use in a variety of clinical applications including treatment of lower extremity ulcers, treatment of cryptoglandular fistulae, and breast augmentation (Marino et al. 2013; Borowski et al. 2015; Kakamura and Ito 2011). The Celution system possesses the CE mark, but is not commercially available in the United States; however, Cytori does have a number of Investigational Drug Exemptions (IDE) for trials using its ADRC technology though. Cytori currently has five clinical trials underway for indications including scleroderma, knee osteoarthritis, urinary incontinence and cutaneous thermal injury.
Another device which has been described in literature is the GID SVF platform mentioned above. The GID SVF platform offers a completely disposable, single use, closed system process using its proprietary enzyme mixture, GIDzyme-2 (GID Europe 2015). The device can process up to 350 cc of dry adipose at one time. Vilaboa et al. (2014) reported that using the GID SVF platform they were able to isolate 719,000 nucleated cells/cc of lipoaspirate with 83 % viability. No information is provided pertaining to progenitor content or clinical applications. The GID SVF platform has received the CE mark for distribution in the European Economic Area (EEA).
A device also reporting high cellular yields is the Tissue Genesis Icellator Cell Isolation system (Tissue Genesis, Honolulu, HI). The Icellator system is an automated, closed system which uses the Tissue Genesis proprietary enzyme blend, Adipase (Tissue Genesis 2015). In 2013, Williams et al. reported a staggering 7.1 million viable SVF cells/mL of canine adipose tissue with 78 % viability processed using the Icellator system (Williams et al. 2013). Another study conducted by Doi et al. (2013) reported a lower, but still impressive yield of 702,000 nucleated cells/cc of lipoaspirate with 80.7 % viability. Doi et al. compared the Icellator system to a manual collagenase-based method using 0.075 % collagenase to digest adipose tissue. They reported that using this manual method they were able to isolate 701,000 nucleated cells/cc of lipoaspirate with 82.4 % viability. No information is provided pertaining to progenitor content. The Icellator system has not been evaluated by the FDA for use in humans.
The Sepax Technology from BioSafe America (Biosafe Group, Lake Geneva, Switzerland) is an enzymatic, fully-automated, closed system. While marketed primarily for cord blood, bone marrow, and peripheral blood processing (Biosafe America 2015), it has been reported for use with adipose tissue as well. Guven et al. (2012) reported a yield of 260,000 nucleated cells/cc of lipoaspirate processed with around 14 % CFU-F, which they compared to a manual, enzymatic method which was able to isolate 160,000 nucleated cells/cc of lipoaspirate with around 11 % CFU-F. Over 90 % viability was reported in both groups. The Sepax-2 system has received a CE mark, 510(k) approval from the FDA and approval from the SFDA in China for processing of cord blood, bone marrow, and peripheral blood, not adipose tissue.
The Lipokit (Medi-Kan Int., West Hollywood, CA) is another semi-automated, enzymatic system. The Lipokit is an all in one system for the harvest, processing and transplant of SVF which can be used with or without enzyme (LipoKit II infomation 2015). The Lipokit uses custom disposable centrifuge syringes for the processing and handling of lipoaspirate, primarily for fat grafting, but can be used for isolation of SVF cells as well. There are very few articles published using the Lipokit, and in these reports, results vary widely. A study by Wang et al. (2012), reported on the effects of using the Lipokit for cell-assisted lipotransfer procedures in 18 patients. They reported 41.67 % ASCs in the SVF, but no data on cell count or viability was able to be acquired from the article. This report was contradicted by Aronowitz et al. (2013), who reported a much lower ASC frequency (1.7 %) with a fairly low nucleated cell yield, only about 35,000 cells/cc of lipoaspirate processed. The Lipokit platform has a CE mark as well as 510 (k) approval from the FDA in the United States as a graft preparation system, but not as an isolation system for SVF cells.
There are fewer mechanical, automated and semi-automated devices available for SVF cell isolation because most mechanical isolations can be conducted using standard laboratory equipment, so there is less of a need for an all in one device. Multiple companies advertise automated and semi-automated, mechanical systems, but many do not have published articles to attest to the yields of these devices. In addition, many of those which have been developed have been deemed to be ineffective in the clinical setting, such as the Fastem/Corios system recently described by Domenis et al. (2015). Domenis compared three methods of SVF isolation and cell-enhanced fat graft preparation. Overall, they concluded that the two enzymatic methods examined, the Lipokit and the Celution system, resulted in significantly more nucleated cells and clonogenic and multipotent progenitor cells for fat graft enhancement, while the Fastem/Corios system was unable to isolate adequate cells to significantly enhance a fat graft. No numbers for nucleated cell count, viability, or progenitor cell content are clearly reported.
One mechanical, semi-automated device which has reported adequate yields is the StromaCell system (Microaire Aesthetics, Charlottesville, VA). The StromaCell system is a patented centrifuge canister which allows for lipoaspirate to be harvested directly into the canister and easy recovery of the SVF cells from the canister after centrifugation at 1000g for 10 min (MicroAire Aesthetics 2013). In a 2014 study by Millan et al. (2014), collagenase based digestion was compared to mechanical isolation using the StromaCell device for SVF isolation. While isolating fewer cells than the standard collagenase-based method (368,000 cells/cc of lipoaspirate vs 140,000 cells/cc of lipoaspirate), they did report similar compositions in terms of progenitor content when analyzed by flow cytometry.
The main drawback of many of these devices is the cost of operation. The closed, enzymatic systems can be very expensive, with some costing over $50,000 for the system. In addition to purchasing the device, many require single-use disposable kits which can cost hundreds or thousands of dollars for a single disposable kit in some cases. A mechanical system like the StromaCell offers the benefit of a closed sterile system and tends to be more affordable, but does not provide the superior yield afforded by the enzymatic systems such as the Cytori Celution system or the Tissue Genesis Icellator system. All of the systems mentioned here can be operated by a single trained technician at the point of care. The processing times vary between systems, with mechanical systems being in the 15–30 min range and the enzymatic systems ranging from about 60–90 min depending on the amount of tissue processed.
Regulatory concerns
Many of the mechanical methods were initially developed in an attempt to isolate a population of cells which could be considered “minimally manipulated,” which many believed would allow them to circumvent a large amount of regulatory oversight by the United States Food and Drug Administration (FDA) and other regulatory agencies around the world. Enzymatic methods produce cell populations which the FDA considers to be “more than minimally manipulated,” causing them to be more heavily regulated as a drug, while the non-enzymatic methods were thought to be considered “minimally manipulated” due to the ambiguity of certain areas of previous regulatory documents. Recent non-binding draft guidances for industry from the FDA (2014b, c) which clarify the FDA’s stance on minimal manipulation and adipose tissue derived HCT/P’s seek to classify all methods of SVF isolation, both enzymatic and mechanical, as yielding “more than minimally manipulated” cells, and thereby classifying SVF as a drug.
Conclusion
Methods used to isolate of pluripotential mesenchymal cells from adipose tissue at the point of care are of increasing importance in medicine as a large body of clinical research shows promise for a burgeoning number of conditions. Mechanical techniques, such as simple washing or centrifugation of lipoaspirate are effective in isolating ASCs. Mechanical methods are appealing because they are simple, quick and generally not associated with expensive equipment or disposables. While more expensive than mechanical options, enzymatic methods for the isolation of stromal vascular fraction cells from adipose tissue yield more nucleated cells with a higher number of progenitor cells per volume of lipoaspirate processed, but overall viability tends to be unaffected by processing method. While mechanical methods may be cost-effective in the laboratory setting, enzymatic methods provide a superior SVF output for use in the clinical setting. The method that a certain lab or facility uses ultimately depends upon their needs and financial capabilities. Labs and clinics with insufficient funding to use enzymatic methods or automated/semi-automated devices still have the option of pursuing mechanical methods. There are differences in the number of adipose stem cells present in the various adipose tissue deposits of an individual and significant variation between individuals but adipose tissue in general is a rich source of pluripotential mesenchymal cells.
Authors’ contributions
All authors contributed equally to the drafting, analysis and critical revisions of this manuscript. All authors read and approved the final manuscript.
Acknowledgements
No other parties assisted in the intellectual development, preparation, funding, or submission of this article other than the listed authors.
Competing interests
The authors declare that they have no competing interests.
Contributor Information
Joel A. Aronowitz, Phone: 310- 659-0705, Email: moc.dmztiwonora@ard.
Ryan A. Lockhart, Email: moc.dmztiwonora@nayr.
Cloe S. Hakakian, Email: moc.dmztiwonora@eolc.
References
- Aronowitz JA, Ellenhorn JD. Adipose stromal vascular fraction isolation: a head-to-head comparison of four commercial cell separation systems. Plast Reconstr Surg. 2013;132(6):932e–939e. doi: 10.1097/PRS.0b013e3182a80652. [PubMed] [Cross Ref]
- Asatrian G, Pham D, Hardy WR, et al. Stem cell technology for bone regeneration: current status and potential applications. Stem Cells Cloning. 2015;8:39–48. [PMC free article] [PubMed]
- Aust L, Devlin B, Foster SJ, et al. Yield of human adipose-derived adult stem cells from liposuction aspirates. Cytotherapy. 2004;6:7–14. doi: 10.1080/14653240310004539. [PubMed] [Cross Ref]
- Baer PC, Geiger H. Adipose-derived mesenchymal stromal/stem cells: tissue localization, characterization, and heterogeneity. Stem Cell Int. 2012;2012:812693. [PMC free article] [PubMed]
- Baptista LS, do Amaral RJ, Carias RB, Aniceto M, Claudio-da-Silva C, Borojevic R. An alternative method for the isolation of mesenchymal stromal cells derived from lipoaspirate samples. Cytotherapy. 2009;11(6):706–715. doi: 10.3109/14653240902981144. [PubMed] [Cross Ref]
- Bertolini F, Lohsiriwat V, Petit JY, et al. Adipose tissue cells, lipotransfer and cancer: a challenge for scientists, oncologists and surgeons. Biochim Biophys Acta. 2012;1862:209–214. [PubMed]
- Bielli A, Scioli MG, Gentile P, et al. Adult adipose-derived stem cells and breast cancer: a controversial relationship. Springerplus. 2014;3:345. doi: 10.1186/2193-1801-3-345. [PMC free article] [PubMed][Cross Ref]
- Biosafe America (2015) Sepax 2. Biosafe Group SA website. Available at: http://www.biosafe.ch/?portfolio=sepax2. Accessed 10 Mar 2015
- Bonab MM, Alimoghaddam K, Talebian F, et al. Aging of mesenchymal stem cell in vitro. BMC Cell Biol. 2006;7:14. doi: 10.1186/1471-2121-7-14. [PMC free article] [PubMed] [Cross Ref]
- Borowski DW, Gill TS, Agarwal AK, et al. Adipose tissue-derived regenerative cell-enhanced lipofilling for treatment of crytpoglandular fistulae-in-ano: the ALFA technique. Surg Innov. 2015;22(6):593–600. doi: 10.1177/1553350615572656. [PubMed] [Cross Ref]
- Bourin P, Bunnell BA, Casteilla L, 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 and Science (IFATS) and the International Society for Cellular Therapy (ISCT) Cytotherapy. 2013;15:641–648. doi: 10.1016/j.jcyt.2013.02.006.[PMC free article] [PubMed] [Cross Ref]
- Breite AG, Dwulet FE, McCarthy RC. Tissue dissociation enzyme neutral protease assessment. Transplant Proc. 2010;42:2052–2054. doi: 10.1016/j.transproceed.2010.05.118. [PMC free article] [PubMed][Cross Ref]
- Buschmann J, Gao S, Harter L, et al. Yield and proliferation rate of adipose-derived stromal cells as a function of age, body mass index and harvest site-increasing the yield by use of adherent and supernatant fractions. Cytotherapy. 2013;15(9):1098–1105. doi: 10.1016/j.jcyt.2013.04.009. [PubMed][Cross Ref]
- Conde-Green A, Rodriguez RL, Slezak S, et al. Enzymatic digestion and mechanical processing of aspirated adipose tissue. Plast Recons Surg. 2014;134:54. doi: 10.1097/01.prs.0000455394.06800.62.[Cross Ref]
- Cytori Therapeutics (2015) Clinical trials page. Available at http://www.cytori.com/en/Technology/ClinicalTrials.aspx. Accessed 10 Mar 2015
- Di Rocco G, Gentile A, Antonini A, et al. Enhanced healing of diabetic wounds by topical administration of adipose tissue-derived stromal cells overexpressing stromal-derived factor-1: biodistribution and engraftment analysis of bioluminescent imaging. Stem Cells Int. 2010;2011:1–11. doi: 10.4061/2011/304562. [PMC free article] [PubMed] [Cross Ref]
- Doi K, Tanaka S, Iida H, et al. Stromal vascular fraction isolated from lipo-aspirates using an automated processing system: bench and bed analysis. J Tiss Eng Regen Med. 2013;7:864–870. doi: 10.1002/term.1478. [PubMed] [Cross Ref]
- Domenis R, Lazzaro L, Calabrese S, et al. Adipose tissue derived stem cells: in vitro and in vivo analysis of a standard and three commercially available cell-assisted lipotransfer techniques. Stem Cell Res Ther. 2015;6(1):2. doi: 10.1186/scrt536. [PMC free article] [PubMed] [Cross Ref]
- Eto H, Kato H, Suga H, et al. The fate of adipocytes after nonvascularized fat grafting: evidence of early death and replacement of adipocytes. Plast Reconstr Surg. 2012;129:1081–1092. doi: 10.1097/PRS.0b013e31824a2b19. [PubMed] [Cross Ref]
- FDA (2014a) Medical devices; current good manufacturing practice (CGMP) final rule; quality system regulation. FDA website. Available at http://www.fda.gov/MedicalDevices/DeviceRegulationandGuidance/PostmarketRequirements/QualitySystemsRegulations/ucm230127.htm. Accessed 10 Mar 2015
- FDA (2014b) Minimal manipulation of human cells, tissues, and cellular and tissue-based products: draft guidance for industry and food and drug administration staff. FDA website http://www.fda.gov/BiologicsBloodVaccines/GuidanceComplianceRegulatoryInformation/Guidances/CellularandGeneTherapy/ucm427692.htm. Accessed 10 Mar 2015
- FDA (2014c) Human cells, tissues, and cellular and tissue-based products (HCT/Ps) from adipose tissue: regulatory considerations; draft guidance for industry. FDA Website. http://www.fda.gov/BiologicsBloodVaccines/GuidanceComplianceRegulatoryInformation/Guidances/Tissue/ucm427795.htm. Accessed 10 Mar 2015
- Fogarty WM, Griffin PJ. Production and purification of metalloprotease of Baccilus polymyxa. Appl Microbiol. 1973;26(2):185–190. [PMC free article] [PubMed]
- Fraser JK, Hicok KC, Shanahan R, et al. The Celution system: automated processing of adipose-derived regenerative cells in a functionally closed system. Adv Wound Care. 2013;3(1):38–45. doi: 10.1089/wound.2012.0408. [PMC free article] [PubMed] [Cross Ref]
- GID Europe (2015) Complete tissue processing in a single disposable device. Available at. http://www.gideurope.com/gid-svf-1/. Accessed 10 Mar 2015
- Griffin PJ, Fogarty WM. Physiochemical properties of the native, zinc- and manganese-prepared metalloprotease of Bacillus polymyxa. Appl Microbiol. 1973;26(2):191–195. [PMC free article][PubMed]
- Guven S, Karagianni M, Schwalbe M, et al. Validation of an automated procedure to isolate human adipose tissue-derived cells by using the Sepax technology. Tissue Eng Methods. 2012;18(8):575–582. doi: 10.1089/ten.tec.2011.0617. [PMC free article] [PubMed] [Cross Ref]
- Han J, Koh YJ, Moon HR, Ryoo HG, Cho CH, Kim I, et al. Adipose tissue is an extramedullary reservoir for functional hematopoietic stem and progenitor cells. Blood. 2010;115:957–964. doi: 10.1182/blood-2009-05-219923. [PubMed] [Cross Ref]
- Jurgens WJ, Oedayrajsingh-Varma MJ, Helder MN, et al. Effect of tissue-harvest site on yield of stem cells derived from adipose tissue: implications for cell-based therapies. Cell Tissue Res. 2008;332:415–426. doi: 10.1007/s00441-007-0555-7. [PMC free article] [PubMed] [Cross Ref]
- Kakamura T, Ito K. Autologous cell-enriched fat grafting for breast augmentation. Aesthetic Plast Surg. 2011;35(6):1120–1130. [PubMed]
- Kapur SK, Katz AJ. Review of the adipose derived stem cell secretome. Biochimie. 2013;95:2222–2228. doi: 10.1016/j.biochi.2013.06.001. [PubMed] [Cross Ref]
- Kato H, Mineda K, Eto H, et al. Degeneration, regeneration and cicatrization after fat grafting: dynamic total tissue remodeling during the first 3 months. Plast Reconstr Surg. 2014;133:303e–313e. [PubMed]
- Ke Q, Chen A, Minoda M, et al. Safety evaluation of a thermolysin enzyme from Geobacillus stearothermophilus. Food Chem Toxicol. 2013;59:541–548. doi: 10.1016/j.fct.2013.06.046. [PubMed][Cross Ref]
- Lin K, Matsubara Y, Masuda Y, et al. Characterization of adipose tissue-derived cells isolated with the Celution system. Cytotherapy. 2008;10(4):417–426. doi: 10.1080/14653240801982979. [PubMed][Cross Ref]
- LipoKit II infomation (2015) Medi-Kan Int. Website. Available at http://www.medikanint.com/lipokit.html. Accessed 16 Mar 2015
- Marino G, Moraci M, Armenia E, et al. Therapy with autologous adipose-derived regenerative cells for the care of chronic ulcers of lower limbs in patients with peripheral arterial disease. J Surg Res. 2013;185(1):36–44. doi: 10.1016/j.jss.2013.05.024. [PubMed] [Cross Ref]
- Markarian FM, Frey GZ, Silveira MD, et al. Isolation of adipose-derived stem cells: a comparison among different methods. Biotechnol Lett. 2014;36:693–702. doi: 10.1007/s10529-013-1425-x. [PubMed][Cross Ref]
- Matsumoto D, Sato K, Gonda K, et al. Cell-assisted lipotansfer: supportive use of human adipose-derived stem cells for soft tissue augmentation with lipoinjection. Tissue Eng. 2006;12(12):3375–3383. doi: 10.1089/ten.2006.12.3375. [PubMed] [Cross Ref]
- McCarthy RC, Breite AG, Dwulet FE (2010) Biochemical analysis of crude collagenase products used in adipose derived stromal cell isolation procedures and development of a purified tissue dissociation enzyme mixture. Available at. http://www.vitacyte.com/wp-content/uploads/2009/01/ifats-vitacyte.pdf. Accessed 3 Nov 2014
- McCarthy RC, Breite AG, Green ML, Dwulet FE. Tissue dissociation enzymes for isolating human islets for transplantation: factors to consider in setting enzyme acceptance criteria. Transplantation. 2011;91:137–145. doi: 10.1097/TP.0b013e3181ffff7d. [PMC free article] [PubMed] [Cross Ref]
- McIntosh K, Zvonic S, Garrett S, et al. The immunogenicity of human adipose derived cells: temporal changes in vitro. Stem Cells. 2006;24:1245–1253. doi: 10.1634/stemcells.2005-0235. [PubMed][Cross Ref]
- MicroAire Aesthetics (2013) Stromacell MicroAire Aesthetics Website. http://old.microaire.com/products/microaire-aesthetics/stromacell/. Accessed 10 Mar 2015
- Millan A, Landerholm T, Chapman JR. Comparison between collagenase adipose digestion and Stromacell mechanical dissociation for mesenchymal stem cell separation. McNair Scholars J CSUS. 2014;15:86–101.
- Mitchell JB, McIntosh K, Zvonic S, et al. Immunophenotype of human adipose-derived cells: temporal changes in stromal-associated and stem cell-associated markers. Stem Cells. 2006;24:376–385. doi: 10.1634/stemcells.2005-0234. [PubMed] [Cross Ref]
- Naderi N, Wilde C, Haque T, et al. Adipogenic differentiation of adipose-derived stem cells in a 3-dimensional spheroid culture (microtissue): implications for the reconstructive surgeon. J Plast Reconstr Aesthet Surg. 2014;67(12):1726–1734. doi: 10.1016/j.bjps.2014.08.013. [PubMed][Cross Ref]
- Nagaishi K, Arimura Y, Fujimiya M. Stem cell therapy for inflammatory bowel disease. J Gastroenterol. 2015;50(3):280–286. doi: 10.1007/s00535-015-1040-9. [PubMed] [Cross Ref]
- Planat-Benard V, Silvestre JS, Cousin B, et al. Plasticity of human adipose lineage cells towards endothelial cells: physiological and therapeutic perspectives. Circulation. 2004;109:656–663. doi: 10.1161/01.CIR.0000114522.38265.61. [PubMed] [Cross Ref]
- Raposio E, Caruana G, Bronomini S, Libondi G. 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. 2014;133(6):1406–1409. doi: 10.1097/PRS.0000000000000170. [PubMed] [Cross Ref]
- Rehmam J, Traktuev D, Li J, et al. Secretion of angiogenic and antiapoptotic factors by human adipose stromal cells. Circulation. 2004;109:1292–1298. doi: 10.1161/01.CIR.0000121425.42966.F1.[PubMed] [Cross Ref]
- Savi M, Bocchi L, Fiumana E, et al. Enhanced engraftment and repairing ability of human adipose-derived stem cells, conveyed by pharmacologically active microcarriers continuously releasing HGF and IGF-1, in healing myocardial infarction. J Biomed Mater Res A. 2015 [PubMed]
- Shah FS, Wu X, Dietrich M, Rood J, Gimble J. A non-enzymatic method for isolating human adipose-derived stromal stem cells. Cytotherapy. 2013;15:979–985. doi: 10.1016/j.jcyt.2013.04.001. [PubMed][Cross Ref]
- Suga H, Eto H, Aoi N, et al. Adipose tissue remodeling under ischemia: death of adipocytes and activation of stem/progenitor cells. Plast Reconstr Surg. 2010;126:911–923. doi: 10.1097/PRS.0b013e3181f4468b. [PubMed] [Cross Ref]
- Tissue Genesis (2015) Tissue Genesis Icellator cell isolation system. Tissue Genesis Website. Available at: http://www.tissuegenesis.com/icellator.html. Accessed 10 Mar 2015
- Ude CC, Sulaiman SB, Min-Hwei N, et al. Cartilage regeneration by chondrogenic induced adult stem cells in osteoarthritic sheep model. PLoS One. 2014;9(6):e98770. doi: 10.1371/journal.pone.0098770.[PMC free article] [PubMed] [Cross Ref]
- Vilaboa SD, Navarro-Palou M, Llull R. Age influence on stromal vascular fraction cell yield obtained from human lipoaspirates. Cytotherapy. 2014;12:1092–1097. doi: 10.1016/j.jcyt.2014.02.007. [PubMed][Cross Ref]
- Wang L, Lu Y, Luo X, et al. Cell-assisted lipotransfer for breast augmentation: a report of 18 patients. Zhonghua Zheng Xing Wai Ke Za Zhi. 2012;28(1):1–6. [PubMed]
- Williams SK, Kosnik PE, Kleinert LB, et al. Adipose stromal vascular fraction cells isolated using an automated point of care system improve the patency of expanded polytetrafluoroethylene vascular grafts. Tissue Eng. 2013;19(11, 12):1295–1302. doi: 10.1089/ten.tea.2012.0318. [PubMed] [Cross Ref]
- Yoshimura K, Shiguera T, Matsumoto D, et al. Characterization of freshly isolated and cultured cells derived from the fatty and fluid portions of liposuction aspirates. J Cell Physiol. 2006;208:64–76. doi: 10.1002/jcp.20636. [PubMed] [Cross Ref]
- Yoshimura K, Suga H, Eto H, et al. Adipose-derived stem/progenitor cells: roles in adipose tissue remodeling and potential use for soft tissue augmentation. Regen Med. 2009;4(2):265–273. doi: 10.2217/17460751.4.2.265. [PubMed] [Cross Ref]
- Zimmerlin L, Donnenberg VS, Pfeifer ME, et al. Stromal vascular progenitors in adult human adipose tissue. Cytometry . 2010;77(1):22–30. [PMC free article] [PubMed]
- Zuk PA, Zhu M, Mizuno H, et al. Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng. 2001;7:211–229. doi: 10.1089/107632701300062859. [PubMed] [Cross Ref]
No comments:
Post a Comment
Note: Only a member of this blog may post a comment.