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

Here is another automatic unit recommended in 2 papers below: waiting to hear from rep on cost.
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

Pablo Bora1,2 and Anish S. Majumdar1

Author information ► Copyright and License information ▼

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.

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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.

Centrifugation of the aqueous fraction yields a reddish pellet which contains SVF cells. Erythrocytes, a major contaminant present in the SVF pellet, can be lysed to isolate a purer population of ADSCs and/or SVF cells if intended for in vitro expansion [7, 30].

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.

Table 1

Important components of SVF, respective sizes, and surface markers

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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].

Fig. 1

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].

Table 2

Overview of characteristics of native and culture expanded ADSCs

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.

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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].

Table 3

Major applications of SVF- and ADSC-based therapeutics with corresponding clinical trials and/or case study references

Table 4

Comparative overview of SVF and ADSCs

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].

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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

Joel A. Aronowitz, Ryan A. Lockhart, and Cloe S. Hakakian

Cedars-Sinai Medical Center, Los Angeles, USA

University Stem Cell Center, 8635 W 3rd St. Suite 1090W, Los Angeles, CA 90048 USA

USC, Keck School of Medicine, Los Angeles, USA

Joel A. Aronowitz, Phone: 310- 659-0705, Email: moc.dmztiwonora@ard.

Contributor Information.

Corresponding author.

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Copyright © Aronowitz et al. 2015

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Abstract

Clinical use of adipose-derived stem cells (ASCs) for a variety of indications is rapidly expanding in medicine. Most commonly, ASCs are isolated at the point of care from lipoaspirate tissue as the stromal vascular fraction (SVF). The cells are immediately administered to the patient as an injection or used to enrich fat grafts. Isolation of ASCs from adipose tissue is a relatively simple process performed routinely in cell biology laboratories, but isolation at the point of care for immediate clinical administration requires special methodology to prevent contamination, ensure integrity of clinical research and comply with regulatory requirements. A lack of practical laboratory experience, regulatory uncertainty and a relative paucity of objective published data can make selection of the optimum separation method for specific indications a difficult task for the clinician and can discourage clinical adoption. In this paper, we discuss the processes which can be used to separate SVF cells from fat tissue. We compare the various mechanical and enzymatic methods. We discuss the practical considerations involved in selecting an appropriate method from a clinical perspective. Studies consistently show that breakdown of the extracellular matrix achieved with proteolytic enzymes affords significantly greater efficiency to the separation process. SVF isolated through mechanical methods is equally safe, less costly and less time consuming but the product contains a higher frequency of blood mononuclear cells and fewer progenitor cells. Mechanical methods can provide a low cost, rapid and simple alternative to enzymatic isolation methods, and are attractive when smaller quantities of ASCs are sufficient.

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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).

Table 1

Summary of reported SVF isolation methods

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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.

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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.

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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).

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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.

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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.

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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.

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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.

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Table 1

Summary of reported SVF isolation methods

References	Method summary	Mechanical or enzymatic	Automated, semi-automated, or manual	Total nucleated cells/cc lipoaspirate	ASC content	Viability
Baptista et al. (2009)	Lipoaspirate incubated with RBC lysis buffer for 15 min, then centrifuged 15 min at 900g	Mechanical	Manual	240,000	12,000/cc of lipoaspirate (5 %)	n/a
Shah et al. (2013)	Lipoaspirate vigorously shaken for 1–2 min with PBS.  Infranatant saved.  Repeated 2 times.  Infranatant centrifuged 1200 rpm for 5 min	Mechanical	Manual	n/a	25,000/cc of lipoaspirate after culture	n/a
	Incubate adipose with 0.1 % collagenase at 37 °C for 1 h.  Centrifuge 1200 rpm 10 min	Enzymatic	Manual	n/a	480,000/cc of lipoaspirate after culture	n/a
Markarian et al. (2014)	Lipoaspirate incubated with RBC lysis buffer for 15 min, then centrifuged 10 min at 600g	Mechanical	Manual	25,000	n/a	65 %
	Centrifuged lipoaspirate at 800g or 1280g for 15 min	Mechanical	Manual	10,000	n/a	70 %
	Lipoaspirate incubated with collagenase solution at 37C for 30 min.  Centrifuge for 10 min at 600g	Enzymatic	Manual	350,000	n/a	65 %
Raposio et al. (2014)	Shake lipoaspirate in vibrating shaker for 6 min at 600 vpm.  Centrifuge 6 min at 1600 rpm.  Considered ASC to be any cell CD31−/CD34+/CD45−	Mechanical	Manual	125,000	6250/cc of lipoaspirate (5 %)	n/a
Mitchell et al. (2006)	Incubate lipoaspirate in 0.1 % collagenase for 60 min at 37 °C	Enzymatic	Manual	308,000	n/a	n/a
Aust et al. (2004)	Incubate lipoaspirate in 0.1 % collagenase for 45 min at 37 °C	Enzymatic	Manual	400,000	n/a	93.9 %
Yoshimura et al. (2006)	Incubate with 0.075 % Collagenase at 37 °C for 30 min with constant agitation	Enzymatic	Manual	1,310,000	n/a	n/a
Suga et al. (2010)	Incubate with 0.075 % Collagenase at 37 °C for 30 min with constant agitation	Enzymatic	Manual	100,000	n/a	n/a
Conde-Green et al. (2014)	High speed centrifugation or vortexing and centrifuging	Mechanical	Manual	11,500 –23,000	MSC frequency: 6–13 %	80–90 %
	Collagenase-based digestion	Enzymatic	Manual	230,000	MSC frequency: 60 %	80–90 %
Fraser et al. (2013)	Cytori Celution System	Enzymatic	Automated	360,000	1900 CFU-F/g (<1 %)	84.7 %
Lin et al. (2008)	Cytori Celution System	Enzymatic	Automated	295,000	CFU-F/g =1.6 %	86.6 %
Aronowitz et al. (2013)	Cytori Celution System	Enzymatic	Automated	240,000	39,000 CFU-F/g (16 %)	93 %
	PNC Multi-Station:  35U collagenase/50 mL lipoaspirate.  Incubate 30 min at 37 °C with constant agitation.  Centrifuge at 2000 rpm for 10 min	Enzymatic	Manual	107,000	6,000 CFU/g (5.6 %)	57 %
	CHA Biotech Cha-Station	Enzymatic	Semi-automated	5000	390 CFU-F/g (7.8 %)	87 %
	Medi-Kan Lipokit with MaxStem	Enzymatic	Semi-automated	35,000	615 CFU-F/g (1.7 %)	72 %
Doi et al. (2013)	Tissue Genesis Cell Isolation system	Enzymatic	Automated	702,000	n/a	80.7 %
	Lipoaspirate incubated with 0.075 % collagenase for 30 min at 37 °C with constant agitation, then centrifuged at 800g for 10 min	Enzymatic	Manual	701,000	n/a	82.4 %
Williams et al. (2013)	Tissue Genesis Cell Isolation System	Enzymatic	Automated	7,100,000	n/a	78 %
Güven et al. (2012)	Sepax Technology	Enzymatic	Automated	260,000	CFU-F frequency 14 %	>90 %
	Lipoaspirate incubated with 0.15 % (w/v) collagenase for 60 min at 37 °C with agitation	Enzymatic	Manual	160,000	CFU-F frequency 11 %	>90 %
Vilaboa et al. (2014)	GID SVF Platform	Enzymatic	Semi-automated	719,000	n/a	83 %
Millan et al. (2014)	StromaCell by Microaire	Mechanical	Semi-automated	140,000	n/a	87.3 %
	Lipoaspirate incubated in 0.2 % (w/v) collagenase for 90 min at 37 °C	Enzymatic	Manual	368,000	n/a	74.5 %
Wang et al. (2012)	Medi-Kan Lipokit	Enzymatic	Semi-automated	n/a	41.67 %	n/a

———–

Ref:

Springerplus. 2014; 3: 345.

Published online 2014 Jul 8. doi:  10.1186/2193-1801-3-345

PMCID: PMC4117859

Adult adipose-derived stem cells and breast cancer: a controversial relationship

Alessandra Bielli, Maria Giovanna Scioli, Pietro Gentile, Sara Agostinelli, Chiara Tarquini, Valerio Cervelli, and Augusto Orlandi

Anatomic Pathology, Tor Vergata University of Rome, Via Montpellier, 00133 Rome, Italy

Plastic Surgery, Department of Biomedicine and Prevention, Tor Vergata University, Rome, Italy

Alessandra Bielli, Email: ti.liamtoh@illeibardnassela.

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Copyright © Bielli et al.; licensee Springer. 2014

This article is published under license to BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly credited.

This article has been cited by other articles in PMC.

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Abstract

Breast cancer is the most common cancer in women and autologous fat grafting is an important clinical application in treatment of post-surgical deformities. The simplicity of fat grafting procedures and the absence of subsequent visible scar prompted an increasing interest for this technique. The plasticity of adipose-derived stem cells (ASCs) obtained from stromal vascular fraction (SVF) of adult adipose tissue provided exciting perspectives for regenerative medicine and surgery. The recent discovery that SVF/ASC enrichment further ameliorates clinical efficacy of grafting ASCs suggest as ASC-mediated new adipogenesis and vasculogenesis. ASC adipogenic differentiation involves Akt activity and EGFRs, FGFRs, ERbB2 receptor-mediated pathways that also play a pivotal role in the regulation of breast cancer growth. Moreover, the finding that platelet-derived growth factors and hormones improved long-term maintenance of fat grafting raises new concerns for their use during breast reconstruction after cancer surgery. However, it remains unclear whether grafted or resident ASCs may increase the risk of de novocancer development or recurrence. Preliminary follow-up studies seem to support the efficacy and safety of SVF/ASCs enrichment and the additional benefit from the combined use of autologous platelet-derived growth factors and hormones during breast reconstruction procedures. In the present review we highlighted the complex interplay between resident or grafted ASCs, mature adipocytes, dormant or active breast cancer cells and tumor microenvironment. Actually, data concerning the permissive role of ASCs on breast cancer progression are contrasting, although no clear evidence speaking against their use exists.

Keywords: Adipose-derived stem cell, Stromal vascular fraction, Fat grafting, Breast cancer, Carcinogenesis, Breast reconstruction, Akt

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Introduction

Adult adipose tissue is a multifunctional organ that contains various cellular types, including mature adipocytes, macrophages and stromal cells, supported by connective tissue surrounding fine capillaries (Zuk et al. 2001; Gentile et al. 2012a; Tran et al. 2012). When isolated in vitro, stromal cells have the potential to form bone, cartilage, muscle and fat tissues and have been variously termed, including preadipocytes and multipotent adipose-derived stem cells. However, the term “adipose-derived stem cells” (ASCs) has been successively recommended for the consistency between research groups (Zhao et al. 2010). In adult adipose tissue (Figure 1A), ASCs are considered to reside between mature adipocytes and extracellular matrix around small vessels (Tran et al. 2012). In fact, transmission electron microscopy of human breast adipose tissue showed the presence of cells featuring ASCs arranged around the endothelial cells of small vessels (Figure 1B), strongly supporting their perivascular origin (Traktuev et al. 2008; Crisan et al. 2008). ASCs likely contribute to adipose tissue cell turn-over (Strawford et al. 2004; Wang et al. 2012a). ASCs can be isolated from subcutaneous adult adipose tissue after liposuction by enzymatic digestion (Gimble et al. 2007; Cervelli et al. 2009). After centrifugation, the obtained heterogeneous mixture of endothelial cells, smooth muscle cells, fibroblast, pericytes, mast cells and preadipocytes is named stromal-vascular fraction (SVF). ASCs can be separated from SVF by adhesion on plastic dishes (Gimble et al. 2007; Cervelli et al. 2009). Before discovering of the plasticity of ASCs, bone marrow was clinically considered the major tissue source of human adult stem cells, the so-called mesenchymal stem cells (MSCs) (Izadpanah et al. 2006). ASCs and MSCs share the ability to differentiate along multiple lineage pathways, including vasculogenetic properties. Cells with stem phenotype and vasculogenetic capacities have been also identified in the circulatory system, in the vessel wall and in various extravascular sites (Grenier et al. 2007; Orlandi & Bennett 2010). Furthermore, ASCs can easily differentiate in mature adipocytes, and adipogenic differentiation is greatly increased by combined treatment with insulin and platelet-derived growth factors, with an increased long-term maintenance of fat grafts (Cervelli et al. 2012). Autologous fat grafting with SVF enrichment for regenerative surgical purposes, in particular in the therapy of post-traumatic lower extremity ulcers, give promising results (Cervelli et al. 2011). Similarly, SVF enrichment allows fat graft maintenance, likely favoring vascularization and collagen synthesis activity (Gentile et al. 2012b). These findings suggest the innovative use of the SVF/ASC enrichment and growth factors also in the breast reconstruction to avoid the frequent complications of fat grafting, including fat necrosis, cyst formation and calcification (Gutowski 2009). Adipose tissue is extremely metabolically active, as documented by its capacity to secrete hormones, growth factors and cytokines by both mature adipocytes and ASCs (Fruhbeck et al. 2001; Kilroy et al. 2007). Similarly, the discovery of ASCs as main actors in the regulatory scenario of adipose tissue cell turn-over requires further attention for the potential interplay between resident, grafted ASCs and residual breast cancer cells or adjacent in situ lesions. This finding induced caution and suggested some concerns about the use of fat grafting with SVF/ASC enrichment in breast reconstruction following cancer surgery. In the present review, we tried to describe the biomolecular pathways regulating proliferation and differentiation of ASCs, in order to define potential implications of breast cancer cell biology and risks for their use in post-surgery breast cancer reconstruction.

Figure 1

Microscopic characterization of human breast adipose tissue. A, Normal mammary adipose tissue after Haematoxylin and Eosin staining. Scale bar, 100 μm. B, transmission electron microscopy image of human breast adipose tissue showing perivascular …

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Phenotypic characterization of adipose-derived stem cells

ASCs share with MSCs the differentiation potential along several mesenchymal lineages (Gimble et al. 2007) (Peng et al. 2008). Nevertheless, some characteristics of ASCs, in particular the maintenance of proliferating ability in culture, are even greater than those of MSCs (Xu et al. 2005). The surface antigen profile of ASCs isolated from human adipose tissue, changes in vitro as a function of time and/or passage in culture (Mitchell et al. 2006). Table 1 summarizes the antigenic profile of ASCs. After two or more passages in vitro, ASC surface immunophenotype resembles that of MSCs, with a similarity greater than 90% (Gimble et al. 2007). Nevertheless, some differences in surface protein expression have been described. The presence of the glycoprotein CD34 on the surface of human ASCs is not reported in MSCs (Pittenger et al. 1999). Since CD34 is abundantly expressed in human ASCs, immunofluorescence makes possible to identify ASCs as CD34+ cells and to differentiate them from circulating precursors (Pittenger et al. 1999), confirming in human adipose tissue the presence of CD34+ cells and the perivascular origin of ASCs (Figure 1C-E). Cytofluorimetry and immunofluorescence represent suitable methods to investigate ASCs phenotype in vitro (Figure 2). Besides mesenchymal markers, such as CD44 and CD90, ASCs display pericytic markers, such as CD140a, CD140b and smooth muscle markers, such as α − smooth muscle actin (Traktuev et al. 2008).

Table 1

Antigen profile of adipose-derived stem cell

Figure 2

Phenotypic analysis of human adipose-derived stem cells. A and B, Flow cytometry depicting the diffuse expression of CD90 and CD44 stromal markers. C and D, Immunofluorescence staining revealing the strong expression of CD44 and CD90 in cultured ASCs. …

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Adipose-derived stem cells and angiogenesis

The fascinating differentiative pluripotency of ASCs and their ability to enhance vascularization (Bertolini et al. 2012; Merfeld-Clauss et al. 2010) progressively increased interest for their use in tissue engineering and regenerative medicine. The perivascular origin of ASCs and the expression of pericytic markers first suggested a role in vascular homeostasis of adipose tissue (Maumus et al. 2011). When transplanted, ASCs have the capacity to maintain the viability of fat transplanted through the secretion of growth factors that improve tissue survival (Kolle et al. 2013). Recent studies indicated that ASCs like MSCs are capable to promote angiogenesis through secretion of growth factors, in particular VEGF (Kinnaird et al. 2004; Salgado et al. 2010). Angiogenesis is a crucial event for cancer growth, and VEGF secretion plays a pivotal role in this process (Tarallo et al. 2010). Stem cells contribute to vascular remodelling by synthesizing collagen and secreting vascular growth factors (Orlandi and Bennett 2010). So, the expression of VEGF receptors in ASCs should be taken into account for future additional new anti-angiogenic strategies (Cassinelli et al. 2012) in breast cancer. It is worth of noting that ASCs share with resident vascular stem cells the paracrine production of VEGF (Cervelli et al. 2012; Ferlosio et al. 2012) and the expression of VEGF receptors (Kinnaird et al. 2004; Salgado et al. 2010). Furthermore, ASCs secrete hepatocyte growth factor, tumor necrosis factor-α and nerve growth factor (Salgado et al. 2010). Nerve growth factor and precursors are capable of inducing vascular remodeling by modulating vascular cells apoptotic sensitivity (Campagnolo et al. 2014).

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Cross-talk between mature adipocytes and breast cancer cells

Many studies focused the relationship between mature adipocytes and breast cancer cells. Rat mature adipocytes affect the biological behavior of epithelial cells through the production of leptin, adiponectin, tumor necrosis factor-α, heparin-binding epidermal growth factor, insulin-like growth factor-II and adipsin (Manabe et al. 2003). Furthermore, mature adipocytes metabolize androgen to estrogen trough their own aromatase activity (Miller et al. 1998). Estrogen synthesis affects breast carcinoma cell growth by a paracrine mechanism (Miller et al. 1998). Since breast adipose tissue is the major site of estrogen biosynthesis, its local delivering is likely involved in cancer progression. As matter of fact, estrogen level in breast tissues results 10 times greater than in blood as a consequence of high aromatase activity and tumor cells release stimulatory factors that amplify aromatase expression (Sasaki et al. 2010). Breast tumor cells also influence and modify the surrounding tissue microenvironment. Recent in vivo and in vitrostudies demonstrated relevant phenotypic changes in adipocytes surrounding breast cancer (Dirat et al. 2011). Murine and human mature adipocytes co-cultured with tumor cells exhibited changes of the number and size of lipid droplets, a decrease of adipose markers level and over-expression of IL6, leading to a more aggressive tumor behavior (Walter et al. 2009). Also, mature adipocytes adjacent to the tumor showed a reduction of the expression of PPARγ and lipid droplet accumulation (Chandler et al. 2012). Adipose tissue may also stimulate the growth and survival of breast tumor cells trough the secretion of adipokines, such as leptin and adiponectin (Bertolini et al. 2012). It appears evident that adipocytes surrounding breast cancer are subjected to significant transcriptional changes with a marked increased expression of endocrine-related factors, influencing the growth and survival of breast cancer cells, in a paracrine loop (Ghosh et al. 2010). These findings support the hypothesis of an intimate cross-talk between breast cancer cells and immediately adjacent adipose tissue cells.

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Adipose-derived stem cells and breast cancer

Differently from mature adipocytes, the interplay between resident mesenchymal cells, including ASCs, and breast epithelial cells is still partially unknown. In this respect, it is still unclear whether preadipocytes act differently from mature adipocytes. ASCs are located in perivascular niches and contribute to cell turn-over, vascular network for the maintenance of adipose tissue tropism (Strawford et al. 2004; Wang et al. 2012a) and to regulate stem cell homeostasis. Dynamic and reciprocal communication between epithelial and stromal compartments occurs during breast cancer progression, with the production and release of a large panel of cytokines, chemokines and growth factors which are essentials for the generation of a more favorable microenvironment for tumor growth (Dirat et al. 2011; Wiseman and Werb 2002). These signals are capable to induce the recruitment of several cells types including MSCs, so promoting cancer growth, metastasis and tumour stroma formation (Kucerova et al. 2013; Karnoub et al. 2007). Tumor microenvironment is heterogeneous and, in recent studies, the presence within tumor bulk of a cancer stem cell has been hypothesised. Cancer stem cells are defined as a subpopulation that constitute a small percentage of the tumor bulk and displayed analogies to normal stem cells, with both embryonic and adult aspects, supported by phenotypic (surface marker) and functional (metabolic enzymes and transporters) features and clonogenic potential (Donnenberg et al. 2013). In the primary tumor, cancer stem cell may arise from the transformation of resident stem cells or from dedifferentiation of differentiated tumor cells in response to specific microenvironmental signals (Park et al. 2014). The acquisition of cancer stem cell features may be a partial reminiscence of an embryonic phenotype with an increase of susceptibility to epithelial-mesenchymal transition, supporting greater tumor growth and invasiveness (Park et al. 2014). Cells shift from a epithelial-like to a spindle-like morphology, accompanied by the expression of CD44 and CD90 stem markers and the maintenance of an adult stem cell phenotype (Donnenberg et al. 2013; Park et al. 2014). Furthermore, epithelial breast cancer cells undergoing to epithelial-mesenchymal-transition showed mesenchymal features with loss of polarity and stem like spindle shape, that favor motility, invasiveness and survival (Hass & Otte 2012). In this context, ASCs may interact with breast cancer cells through the formation of gap junctions that allows intercellular communication and the exchange of low molecular weight compounds (Donahue et al. 2003). The presence of gap junctions correlate with a more malignant phenotype and greater tumor progression and they can modulate the metastatic potential of the breast cancer cells (Mandel et al. 2013). Thus, the inhibition of gap junctions could partially block the stem cell-mediated growth induction and CD90 expression in breast cells. Conflicting data are reported concerning the role of ASCs in cancer progression. As reported, ASCs express surface markers, such as CD44, able to anchor some matrix-metalloproteinases to the cell surface. This CD44-matrix-metalloproteinases association mediates the reorganization of extracellular matrix components (Hass and Otte 2012). Moreover, experiments in vivo and in vitro reported that ASCs favor tumor growth, increasing extracellular matrix deposition and vascularization, suggesting that ASCs may directly contribute to the dense network of fibroblasts and desmoplastic reaction surrounding breast cancer (Wang et al. 2012a). The desmoplastic reaction represents the stromal response to cancer cell infiltration and it is due to the disruption of the basement membrane and the inflammatory remodeling of the extracellular matrix (Pinilla et al. 2009). Desmoplastic reaction involves increased activity of tissue metalloproteinases and studies in vitro documented that co-culture of human ASCs and breast cancer cells induce high levels of metalloproteinases (Pinilla et al. 2009). Desmoplastic reaction also promotes myofibroblasts recruitment (De Wever et al. 2008; Karagiannis et al. 2012) and a large number of myofibroblasts are documented in the stromal compartment of invasive human breast cancers (Gehmert et al. 2010; Orimo et al. 2005). Myofibroblasts are stromal fibroblasts with both myocyte and fibroblast features (Orlandi et al. 1994). ASCs isolated from tumors also express high levels of alpha-smooth muscle actin, a well-known myofibroblastic marker (Tomasek et al. 2002). This suggests that ASCs could act as a potential source of tumor myofibroblasts.

To better clarify their role in cancer progression, studies in vivo and in vitro have been performed to verify ASCs influence on dormant tumor cells and on their growth and invasiveness. In literature is not yet clear whether dormant tumor cells are out of cell cycle, or persist in a dynamic state of proliferation and death. The transition between dormant and active states requires the presence of various signals, such as cytokines, hormones and growth factors (Donnenberg et al. 2010). Xenograft model experiments documented that grafted ASCs act in a different manner on active and dormant breast cancer cells. In fact, whereas the active cancer cells require growth factors and a new vascular network for the survival and invasiveness, the dormant cancer cells do not immediately require support of these factors. The latter are more autonomous and their growth is slower (Donnenberg et al. 2010; Zimmerlin et al. 2011). Consequently, these results indicate that grafted ASCs favour the growth of active, but not dormant, breast cancer cells. Moreover, ASCs transplantation or co-injected into mouse breast cancer model did not promote tumor growth or metastasis, and this inhibitory effect has been identified as the cause for ASC-dependent Poly ADP ribose polymerase cleavage, so inducing tumor cell apoptosis (Sun et al. 2009).

Altogether, these studies highlight the concept that resident ASCs and cancer cells may interact in a complex and dynamic fashion influencing the tumor behavior. Further studies are needed to better clarify this in vivo interaction and define how selectively stimulate ASCs regenerative function without influencing tumorigenesis.

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Similarities of growth and differentiative pathways of adipose-derived stem cells and breast cancer cells

The proliferative arrest and/or cell loss are potential limitations in regenerative surgery strategies. So, exogenous growth factors should provide the necessary microenvironmental signals to accelerate cell proliferation, extracellular matrix synthesis and tissue deposition (Chen et al. 2010). Various receptor pathways regulated ASC proliferation and differentiation. Epidermal growth factor receptors (EGFRs), fibroblast growth factor receptors (FGFRs) and ErbB tyrosine kinase receptor (ErbB) families are involved in growth control and differentiation of cancer stem cells (Flageng et al. 2013; Nguyen et al. 2013; Reed et al. 2012; Liu et al. 2009). Recent studies documented the presence of EGFR and ErbB2 transcripts and proteins in ASCs (Cervelli et al. 2012). Platelet-derived growth factors stimulated ASCs proliferation and improved the maintenance of breast fat grafting in patients affected by soft tissue defects (Cervelli et al. 2012), but did not affect adipogenic differentiation of ASCs in vitro (Cervelli et al. 2009). This suggests that the pathways regulating proliferation and differentiation of ASCs are partially distinct. Platelet-derived growth factors clinically ameliorated efficacy of fat grafting, likely favoring ASC proliferation (Cervelli et al. 2009). Insulin further increase long-term fat graft maintenance and greatly promoted adipogenic differentiation by increasing Akt activity and down-regulating the expression and activity of EGFR and ErbB2 receptors, without significant proliferative arrest of ASCs (Cervelli et al. 2012). Adipogenic differentiation also associated to the increased FGFR-2 and FGFR-1 transcript levels, suggests a complex receptor-mediated control of adipogenesis in ASCs (Cervelli et al. 2012). Although the clinical use of growth factors may improve long-term fat graft volume maintenance, growth factors may also influence the activity of resting cancer cells (Liu et al. 2009). As reported above, estrogen sustains growth in breast cancer through the transcriptional up-regulation of various growth factors, such as EGFR, and Akt phosphorylation (Liu et al. 2009). Moreover, a cross-talk between erbB and estrogen receptor-mediated signaling has been reported in tumor progression and resistance to endocrine therapy of breast cancer cells (Normanno et al. 2005). In addition, aberrant expression and activation of FGFR activity is involved in the progression of breast cancer (Grose et al. 2007). In particular, FGFR1 expression is associated with an early relapse and poor survival of breast cancer patients (Turner et al. 2010). In vitro data alone seem to suggest the caution in the local use of growth factors in addition to fat graft and further investigation of the interplay between ASCs and breast cancer cells should be performed also in vivo.

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Breast lipografting: clinical studies and follow-up

Autologous fat grafting is a procedure widely used in breast reconstruction after cancer surgical treatment (Gentile et al. 2012b; Coleman and Saboeiro 2007). Clinical trials documented the absence of significant differences between patients undergoing lipofilling after mastectomy compared to untreated group. To minimize adverse effects, many attempts have been made to improve long term fat graft maintenance. A recent series of cases of breast reconstruction performed using autologous fat graft documented that, after a 10 years follow-up, there is no increased risk of relapse or new cancer development (Delay et al. 2009). In another study, 321 patients after breast cancer surgery treatment were subjected to lipofilling treatment (Petit et al. 2012). After six month, patients with lipofilling showed no relapse compared to untreated group. Nevertheless, when the study focused on patients previously diagnosed with breast intraepithelial neoplasia, the lipofilling group displayed a slightly higher risk of local recurrence, although not statistically significant, compared to the untreated group (Petit et al. 2012). However, a similar study carried out on 158 patients subjected to fat grafting procedures after a history of breast cancer surgery, did not show any increase risk of relapse after 18 month of follow-up (Rietjens et al. 2011). Other works compare the local recurrence before and after lipotransfer in patients undergoing mastectomy, with no statically significant differences between groups (Rigotti et al. 2010). Preliminary data from a relatively small number of patients describe that SVF enrichment improves fat graft survival, with no evidence of breast malignant transformation (Kolle et al. 2013). A recently introduced new technique combine the use of autologous SVF enrichment with platelet-derived growth factors to improve fat grafting maintenance after breast reconstruction. A series of 23 patients with breast cancer undergoing post-surgical breast reconstruction with fat grafting-SVF and platelet-derived growth factors did not show increased risk of new cancer development after 1 year follow-up compared with the control group, but evidenced the ameliorated maintenance of breast volume (Cervelli et al. 2012). Although preliminary, these results seem to support that the addition of platelet-derived growth factors to lipografting induces a safe improvement of breast volume maintenance (Gentile et al. 2013), likely for the ability of platelet-derived growth factors to stimulate vascularization (Coleman and Saboeiro 2007; Gentile et al. 2013). Clinical studies with more patients and a longer follow-up are needed to confirm the safety of SVF/ASCs enrichment during fat grafting procedures with or without platelet-derived growth factors and hormones in breast cancer patients.

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New stem cell therapies and surgical breast cancer reconstruction

Conventional cancer therapies include surgery, chemotherapy and radiotherapy. A certain number of preclinical studies recently proposed the use of MSCs as candidates to deliver anti-cancer drugs. Chemokines secreted by breast tumor cells are capable of stimulating MSCs migration and recruitment, suggesting a potential role for MSCs as delivery agents for chemotherapeutic purposes in breast tumours in vivo. MSCs can be readily transduced via adenoviral, retroviral or lentiviral vectors without compromising the capability for differentiation or the expression of surface markers. Consequently, MSCs are potentially suitable for a gene approach in cancer therapy through the induction of a more chronic and slow release of drugs that are often limited by their toxicity or short life (Kucerova et al. 2008). Interferon-β is a powerful inhibitor of tumor cell growth, but to be efficacy it needs a dose higher than the maximally tolerated. In vitro, MSCs transduced with adenoviral vector carrying human Interferon-β and co-cultured with breast cancer cells induced the reduction of cancer cells growth (Studeny et al. 2004). The same effect occurred, in vivo, when Interferon-β − transfected MSC cells are injected intravenously in a xenograft breast cancer mouse model (Studeny et al. 2004). The most of gene approaches use cytosine deaminase/5-fluorocytosine and thymidine kinase/ganciclovir. Ganciclovir is an inhibitor of DNA polymerase and, after DNA incorporation, inhibits chain elongation (Matuskova et al. 2012). The combined use of yeast cytosine deaminase gene with 5-fluorocytosine allows the activation of 5-fluorouracile, a drug normally used in conventional chemotherapy (Matuskova et al. 2012). However, when ASCs are transduced with thymidine kinase/ganciclovir the growth of breast cancer cells was inhibited, but the latters have proved resistant to cytosine deaminase/5-fluorocytosine treatment. These opposite effects are linked to the capability of adult stem cells and tumor cells to communicate via gap junctions, determining both chemosensitivity than chemoresistance (Matuskova et al. 2012; Kucerova et al. 2012). In other works ASCs has been tested as vehicle to deliver tumor necrosis factor-α and to induce TRAIL-mediated apoptosis of cancer cells (Grisendi et al. 2010). TRAIL is a member of the tumor necrosis factor super-family that induces a selective apoptosis through the activation of death receptors in cancer target cells, with no relevant effects on healthy cells (Grisendi et al. 2010). More recently, PPARγ ligands were shown capable of stimulating the differentiation of several cancer cells types, including breast cancer cells lines, suggesting a therapeutic utility in breast cancer treatment. PPARγ ligands inhibit the expression of aromatase and hence estrogen biosynthesis in adipose tissue surrounding human breast cancer (Rubin et al. 2000). At present, most hormonal therapies for breast cancer aim to the inhibit estrogen receptor and/or aromatase activity of cancer cells (den Hollander et al. 2013). Tamoxifen is an antagonist of estrogen receptor widely used in therapy of breast cancer, but after several years of treatment clonal cell line tumors become unresponsive to the drug (Higgins and Stearns 2009). The mechanism that underly tamoxifen resistance is still unclear. It’s possible that the presence of cancer stem cell confers a drugs resistance. In vitro studies documented that cancer stem cell exert antiapoptotic effect on breast cancer cells and counteract cell-cycle changes caused by tamoxifen, so promoting tumor growth and invasiveness (Wang et al. 2012b; Lin et al. 2013).

Aromatase inhibitors are used as second-line therapy or as first-line adjuvant therapy, but they have the disadvantage to inhibit indiscriminately aromatases, including those in bone and brain tissues, with adverse effects in terms of bone mineralization and cognitive function, respectively (Rubin et al. 2000). An ideal goal is to develop a tissue-selective aromatase inhibitor. In these sense, ASCs potentially retain many of the attributes for an optimal cellular vehicle (Rubin et al. 2000). Additional studies need to document efficacy and safety of engineered ASCs before their application in clinical trials.

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Conclusions

Current evidence sustains that ASCs represent a promising tool for innovative therapies in regenerative surgery and play a significant role in lipofilling-mediated breast reconstruction after breast cancer surgery. SVF/ASCs enrichment seems to favor long-term fat graft maintenance in reconstruction of tissue defects, likely promoting vascularization and collagen synthesis. Preliminary studies in vivo seem to confirm the efficacy of SVF/ASCs enrichment and the beneficial additional use of autologous platelet-derived growth factors and hormones in breast reconstruction. The improvement in long-term maintenance strongly supports the additional combined use of fat grafts with autologous platelet-derived growth factors and hormones, such as insulin. However, additional translational research studies are needed to better clarify the possible impact of these procedures on tumor microenvironment, in particular their potential effect on cancer cells. Different studies confirmed the complex and dynamic interplay between cancer cells and resident ASCs. Latters, in the tumor microenvironment, seem to affect only active cancer cells, so promoting neoangiogenesis, matrix remodeling and intercellular communication via gap-junction. In addiction, it has been hypothesized the presence of cancer stem cells, from resident stem cell or dedifferentiated tumor cells, that may favour the epithelial-mesenchymal transition, supporting tumor growth and invasiveness. In addition, the interaction between grafted ASCs and resting cancer cells doesn’t seem to be responsible for cancer recurrence because resting cancer cells are more resistant to apoptosis and they don’t require stroma or vascular structure for their survival. Preliminary data describe that SVF/ASCs enrichment did not show increased risk of new cancer or relapse compared with control group.

Finally, ASCs characteristics appear promising for their engineered use as “carrier” of adjuvant chemotherapeutic agents against residual breast cancer cells. So, the growth of malignant cells may be counteracted by local release of drugs in tumor microenvironment while systemic plasma concentration remain low, avoiding the problems related to toxicity and short life.

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Acknowledgments

We thank Dr Sabrina Cappelli, Dr Antonio Volpe for the technical work.

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Footnotes

Competing interests

The authors declare that they have no competing interest.

Authors’ contribution

AB, MGS, AO: conception and design;PG, VC: collection and assembly of data;SA, CT: data analysis and interpretation; AO: writing and final approvation of the manuscript. All authors read and approved the manuscript.

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Contributor Information

Alessandra Bielli, Email: ti.liamtoh@illeibardnassela.

Maria Giovanna Scioli, Email: ti.2amorinu.dem@iloics.

Pietro Gentile, Email: ti.orebil@4002elitnegorteip.

Sara Agostinelli, Email: ti.liamtoh@aras.illenitsoga.

Chiara Tarquini, Email: ti.orebil@iniuqrat-araihc.

Valerio Cervelli, Email: ti.oiligriv@illevrecoirelav.

Augusto Orlandi, Email: ti.2amorinu@idnalro.

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Plastic and Reconstructive Surgery Issue: Volume 133(6), June 2014, p 1406–1409 Copyright: ©2014American Society of Plastic Surgeons Publication Type: [Experimental: Ideas and Innovations] DOI: 10.1097/PRS.0000000000000170 ISSN: 0032-1052 Accession: 00006534-201406000-00019 Hide Cover [Experimental: Ideas and Innovations]< Previous Article Table of Contents Next Article > A Novel and Effective Strategy for the Isolation of Adipose-Derived Stem Cells: Minimally Manipulated Adipose-Derived Stem Cells for More Rapid and Safe Stem Cell Therapy Raposio, Edoardo M.D., Ph.D.; Caruana, Giorgia M.D.; Bonomini, Sabrina M.D.; Libondi, Guido M.D. Author Information Parma, Italy From the Department of Surgical Sciences, Plastic Surgery Division, University of Parma; and the Department of Clinical and Experimental Medicine, Division of Hematology, Parma University Hospital. Received for publication September 12, 2013; accepted December 12, 2013. Disclosure: The authors have no financial interests to declare in relation to the content of this article. This article did not require any sources of funding. Edoardo Raposio, M.D., Ph.D., Department of Surgical SciencesUniversity of Parma, Via Gramsci 14, 43126 Parma, Italyedoardo.raposio@unipr.it Abstract Summary: Adipose-derived stem cells are an ideal mesenchymal stem cell population for regenerative medical application. The isolation procedure is performed by mechanical isolation under a laminar air flow bench without using serum or animal-derived reagents; cells were characterized by flow cytometric analysis. Cell availability is improved compared with enzymatic digestion procedures. The adipose-derived stem cell mechanical isolating procedure presented here is easier, safer, cheaper, and faster than traditional currently performed enzymatic procedures. Adipose-derived stem cells are recognized as being an effective mesenchymal stem cell population with enormous potential in different fields of regenerative medicine and stem cell therapy. Although they have been extensively characterized in the laboratory with in vitro and in vivo studies to test their multipotency and safety, to this date, there have been few existing clinical trials involving adipose-derived stem cells in cell therapy.1 Although there is unanimous agreement on the harvesting procedure for adipose tissue, there are various protocols for adipose-derived stem cell 2,3 isolation. Most of these protocols use enzymatic digestion with collagenase, serum, and animal-derived medium. Moreover, the procedure takes approximately 2 hours. This study presents a novel and effective strategy for isolating adipose-derived stem cells from a conventional liposuction procedure. The entire isolation process takes approximately 15 minutes; no collagenase, serum, or animal-derived reagents are needed; and it yields a conspicuous amount of adipose-derived stem cells, which are useful for more rapid and safe cell therapy. They started with 80ml adipose tissue: Starting from 80 ml of adipose tissue, a mean of 5 × 105 of adipose-derived stem cells (range, 4.0 to 6.0 × 105; SD, ±1 × 105), 5 percent of the total number of sample cells (1 × 107), were routinely collected, with other cells (95 percent) mostly being blood-derived and endothelial cells.