Access the full text.
Sign up today, get DeepDyve free for 14 days.
Loading next page...
/lp/ou_press/utility-of-adipocyte-fractions-in-fat-grafting-in-an-athymic-rat-model-sTaPIr70GU
- Publisher
- Oxford University Press
- Copyright
- © 2018 The American Society for Aesthetic Plastic Surgery, Inc. Reprints and permission: journals.permissions@oup.com
- ISSN
- 1090-820X
- eISSN
- 1527-330X
- DOI
- 10.1093/asj/sjy111
- Publisher site
- See Article on Publisher Site
Abstract
Abstract Background Multiple processing and handling methods of autologous fat yield to variations in graft retention and viability, which results in unpredictable clinical outcomes. Objectives This study aims to understand the skin effects of fat graft preparations that contain a varying ratio of free-lipid and stem-cell-bearing stromal vascular fractions (SVF). Methods Lipoaspirates from consenting patients were processed into emulsified fat and then SVF and adipocyte fractions (free-lipid). SVF enriched with 0%, 5%, and 15% free-lipid were grafted along the dorsum of athymic rats. The xenografts were collected 45 days after grafting and then prepped for immunostaining. Results Xenografts resulted in viable tissue mass under the panniculus carnosus of rats as confirmed with human specific markers. A low percentage of human cells was also detected in the lower reticular dermis. Although grafts with SVF formed adipocytes of normal architecture, grafts formed with free-lipid alone resulted in large lipid vacuoles in varying sizes. Among graft preparations, SVF with 10% free-lipid resulted in much-developed adipocyte architecture with collagen and elastin. Compared with SVF alone grafts, SVF with free-lipid had higher CD44 expression, suggesting a localized immune response of adipocytes. Conclusions Current studies suggest that SVF enriched with approximately 10% free-lipid provides the best conditions for fat graft differentiation into viable fat tissue formation as well as collagen and elastin production to provide mechanical support for overlaying skin in an athymic rat model. Additionally, application of this therapeutic modality in a simple clinical setting may offer a practical way to concentrate SVF with free-lipid in a small volume for the improvement of clinical defects. Level of Evidence: 5 Autologous fat grafting is increasingly utilized across a range of aesthetic and reconstructive procedures in plastic surgery, including the face, breast, and buttocks.1 In 2015, autologous fat grafting to the face alone was the ninth most popular surgical cosmetic procedure in the United States.1 Previous studies with autologous fat grafting demonstrated its potential and usefulness as a means of skin rejuvenation from improving dermal elasticity for facial scars to rejuvenating skin from radiotherapy scars and ulcers.2–4 Advantages of autologous fat grafting include: (1) there is no risk of immunological rejection; (2) it is long-lasting without the risk of infection; (3) it has the ability to rejuvenate and take on the properties of the injected area; (4) it is natural in appearance and texture; and (5) it is technically easy to obtain and inexpensive.5 Despite its benefits and popularity in recent years, autologous fat has multiple processing and handling methods that yield to variations in fat graft retention and viability, which results in unpredictable clinical outcomes. Currently, there is no compelling objective evidence to advocate for a singular technique of lipoaspirate, and the underlying molecular mechanisms belying the varying fat grafting processes are not well understood.6–10 Among the variety of methods currently utilized for fat harvest processing, one that maintains the highest concentration of the adipose-derived mesenchymal stem cells (AdMSCs) in the fat graft is correlated with the highest fat graft retention and viability.10–12 A recent clinical study by Tonnard and colleagues13 reported significant improvements in skin quality 6 months after autografting of special fat, termed “nanofat,” which incorporates smaller injection cannulae up to 27 gauge. This method is very practical in a clinical setting, because it does not require complicated and lengthy processing of lipoaspirate to harvest. In this method, mechanical emulsification of lipoaspirate fractures adipocytes, leaving a mixture of cells, including AdMSCs, endothelial cells, macrophages, monocytes, granulocytes, and lymphocytes, termed nanofat. Although long-term viability and effectiveness of the nanofat grafting is reported, it is not clear whether these mesenchymal stem cells (MSCs) are solely responsible for the skin rejuvenation. Hence, adipocyte fractures and other cells such us macrophages may play a critical role in facilitating AdMSC differentiation into adipocytes and the production of extracellular matrix (ECM) molecules. It is also possible that the inflammatory response to fat grafting through monocytes and macrophages plays a critical role in fat tissue rejuvenation by improving overlying skin quality and elasticity, as suggested previously.14 Here we show the utility of free-lipid in fat graft generated by a modification of the nanofat procedure. This study utilized an athymic rat subdermal grafting model with various human lipoaspirate processing methods: emulsified fat, stromal vascular fraction (SVF) enriched with free-lipid (0%, 5%, 10%, and 15% adipocyte fractions), free-lipid alone, and regular fat (lipoaspirate or macrofat). The different grafts were analyzed to study fat tissue formation/rejuvenation by evaluating histomorphology, adipocytes, macrophages, adipocyte immune response, and the ECM molecules collagen and elastin. METHODS Lipoaspirate Processing Lipoaspirate was obtained from consented patients undergoing body-contouring procedures in the Department of Plastic Surgery at UT Southwestern Medical Center in accordance with Institutional Review Board-approved protocol. Adipose tissue utilized for this study came from 6 female patients between the ages of 29 and 61 years (mean, 43 years; SD, ± 11.9) during a 2-year period (June 2013 to May 2015). Patients were excluded from having adipose tissue harvested if they were pregnant, planning on becoming pregnant, had cancer, had emotional or cognitive impairment, were HIV positive, were hepatitis positive, or had chronic diseases that would affect the integrity of the waste remnant. The lipoaspirate collection and fat processing were done as described previously with modifications.13 Briefly, the patient’s abdomen was infiltrated with a wetting solution consisting of 1 L of Ringer’s lactate and one ampule of epinephrine (1:1000), so that a 1 mL infiltration to aspiration was obtained to achieve a 1:1 ratio of aspiration to filtration. Utilizing the standard Coleman technique, approximately 50 mL of lipoaspirate was collected. The rinsed lipoaspirate was then centrifuged at 300 x g for 3 minutes to separate the lipoaspirate (fat tissue) from the serum and saline (Figure 1A). The lower portion of the sample containing fat tissue was then emulsified between two 10-cc syringes connected through a Luer Lock connector and passed between the syringes 30 times to generate emulsified fat (Figure 1B). The emulsified fat was then centrifuged at 300 x g for 3 minutes for 5 times allowing a separation of broken-down adipocytes (free-lipid) and SVF containing a mixture of cells from mesodermal or mesenchymal origin including pre-adipocytes, fibroblasts, endothelial cells, vascular smooth muscle cells, immune cells, and adipose derived stem cells (ADSCs).15 During the lipoaspirate processing, 8 treatment groups of graft preparations were collected that are lipoaspirate (fat tissue), serum/saline, emulsified fat, free-lipid alone (adipocyte fractions), SVF alone (0% free-lipid), and SVF enriched with free-lipid (5%, 10%, and 15% adipocyte fractions) as listed in Table 1. Table 1. Fat Graft Preparations Graft type Procedure Lipoaspirate Serum/saline Centrifugation of lipoaspirate (Figure 1A) Emulsified fat (nanofat) Emulsification of fat tissue (Figure 1B) Free-lipid Centrifugation of emulsified fat (Figure 1C) SVF Centrifugation of emulsified fat (Figure 1C) SVF + 5% FL SVF containing 5% free-lipid SVF + 10% FL SVF containing 10% free-lipid SVF + 15% FL SVF containing 15% free-lipid Graft type Procedure Lipoaspirate Serum/saline Centrifugation of lipoaspirate (Figure 1A) Emulsified fat (nanofat) Emulsification of fat tissue (Figure 1B) Free-lipid Centrifugation of emulsified fat (Figure 1C) SVF Centrifugation of emulsified fat (Figure 1C) SVF + 5% FL SVF containing 5% free-lipid SVF + 10% FL SVF containing 10% free-lipid SVF + 15% FL SVF containing 15% free-lipid FL, free-lipid; SVF, stromal vascular fraction. View Large Table 1. Fat Graft Preparations Graft type Procedure Lipoaspirate Serum/saline Centrifugation of lipoaspirate (Figure 1A) Emulsified fat (nanofat) Emulsification of fat tissue (Figure 1B) Free-lipid Centrifugation of emulsified fat (Figure 1C) SVF Centrifugation of emulsified fat (Figure 1C) SVF + 5% FL SVF containing 5% free-lipid SVF + 10% FL SVF containing 10% free-lipid SVF + 15% FL SVF containing 15% free-lipid Graft type Procedure Lipoaspirate Serum/saline Centrifugation of lipoaspirate (Figure 1A) Emulsified fat (nanofat) Emulsification of fat tissue (Figure 1B) Free-lipid Centrifugation of emulsified fat (Figure 1C) SVF Centrifugation of emulsified fat (Figure 1C) SVF + 5% FL SVF containing 5% free-lipid SVF + 10% FL SVF containing 10% free-lipid SVF + 15% FL SVF containing 15% free-lipid FL, free-lipid; SVF, stromal vascular fraction. View Large Figure 1. View largeDownload slide Illustration of lipoaspirate processing and subdermal injection. Figure 1. View largeDownload slide Illustration of lipoaspirate processing and subdermal injection. Animals and Fat Grafting All animals were handled and euthanized in accordance with the standards of humane animal care described by the National Institutes of Health Guide for the Care and Use of Laboratory Animals, utilizing protocols approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Texas Southwestern Medical Center at Dallas. A total of 14 male athymic rats (RNU; Foxn1rnu) were purchased from Charles River Laboratories (Wilmington, MA). The rats were housed in a temperature-controlled sterile environment with a 12-hour light cycle. Animals were fed standard chow (#2916; Harlan-Teklad, Houston, TX) and water was available ad libitum. At approximately 10 weeks of age, the athymic rats were anesthetized with 2% isoflurane and their back hair was shaved. The hairless skin was then tattooed in 3 to 4 columns of 5 dots with black ink to serve as markers for fat grafting. The rats were anesthetized again 3 days later, and the animals were prepared for grafting. The graft preparations (saline-serum mix, free-lipid, emulsified fat, or SVF enriched with 5%, 10%, or 15% free-lipid as presented in Table 1) were administered under the panniculus carnosus along the rats’ dorsum in a linear dot pattern with a 22-gauge needle as shown in Figure 1D. The so-called “dermal wheel” was raised in each dot area by applying a 50 μl to 100 μl sample volume. To manage postoperative pain, rats were administered 0.01 mg/kg buprenorphine and given 4.4 mg/kg carprofen wafers. Buprenorphine was further administered every 8 to 12 hours for the next 48 hours. Tissue Harvesting Tissues were harvested at 45 days from the graft placement. The rats were deeply anesthetized and then euthanized with 0.5 mL of 120 mg/kg pentobarbital barbiturate. Incisions were made along the tailbase of the dorsum from left to right, and the dorsal skin layer was elevated to visualize the tattoo markings. Grafts were identified by gross analysis and the proximity of the black tattoo markings. The grafts along with the surrounding tissues were then excised. Histomorphology and Immunostaining Rat skin with grafts were cut into cranial sections and fixed in 10% neutral buffered formalin and gently shaken for 48 hours. Each graft sample was then embedded in paraffin wax, sectioned, and stained with hematoxylin and eosin (H&E) and Masson’s trichrome. Sections (5 μm) were deparaffinized and preincubated with 10% normal donkey serum (Life Technologies, Carlsbad, CA) for 20 minutes at room temperature. Subsequently, tissue sections were incubated with 10% donkey serum containing the primary antibodies elastin (ab21610, Abcam, Cambridge, United Kingdom), adiponectin (MAB1273B, Sigma-Aldrich, St. Louis, MO), mitochondria (HPA051767, Sigma-Aldrich), CD68 (ab955, Abcam), and CD44 (PA5-21419, Thermo Fisher Scientific, Waltham, MA) in 10% normal donkey serum for approximately 16 hours at 4°C. The slides were washed three times with phosphate buffered saline (PBS), and then incubated with a solution containing Alexa Fluor 488 or 555 conjugated secondary antibodies (1:1000 dilution; Life Technologies) in 10% normal donkey serum. This secondary incubation was done for 1 hour at 25°C. The slides were washed 3 times in PBS, and coverslips were mounted to the slides with ProLong Gold containing DAPI (4’,6-diamidino-2-phenylindole, Life Technologies). Immunohistochemistry and fluorescein images were obtained utilizing an Olympus LX51 microscope (Olympus Corporation, Tokyo, Japan) equipped with DP72 camera and Leica TCS-SP laser scanning confocal microscope (Leica Camera, Wetzlar, Germany), respectively. ImageJ software was utilized to generate individual images. The percent area collagen and relative fluorescent signal intensity ± SEM, was quantified from immunohistochemistry and confocal images, respectively, utilizing ImageJ software according to its user guide. For each calculation, 3 to 4 areas from a section of 3 separate rats were utilized. Statistical Analysis for Image Evaluation Intensity data acquired from histological images were analyzed utilizing one-way analysis of variance (ANOVA) with pairwise multiple comparisons performed with the Tukey test for data normally distributed. Data are displayed as mean ± SEM. RESULTS According to histomorphological evaluation of the sections with Masson’s trichrome staining, H&E showed that grafted tissue developed below the muscle layer of the rat skin with no notable changes in the rat epidermis or dermis (muscle layer thickness varies among rats, explaining the thickness, and it is not caused by experimental procedures or tissue handling). The subdermal location of the graft allowed a clean assessment of graft viability and the degree of fat tissue formation. Human-specific adiponectin and mitochondria staining confirmed the origin and viability of the human fat graft forming the subdermal layer and the smaller percentage in the reticular dermis above the rat panniculus carnosus after 45 days post-inoculation (Figure 2). Serum grafts did not yield to any viable human cells and served as a negative control to demonstrate the specificity of the human antibodies (Figure 2, bottom row). Figure 2. View largeDownload slide Confirmation of human xenograft viability 45 days after subdermal fat grafting in athymic rat. Human specific antibodies against adiponectin expressed on adipocytes (red) and mitochondria of all human cells (green) counter-stained against DAPI (4’,6-diamidino-2-phenylindole, blue), which marks nuclei in both human and rat cells. The dotted orange line represents muscle and graft junction. E, epidermis; D, dermis; M, muscle (panniculus carnosus); and G, graft. The representative images are of at least 3 grafts/group. The scale bar represents 500 µm. Figure 2. View largeDownload slide Confirmation of human xenograft viability 45 days after subdermal fat grafting in athymic rat. Human specific antibodies against adiponectin expressed on adipocytes (red) and mitochondria of all human cells (green) counter-stained against DAPI (4’,6-diamidino-2-phenylindole, blue), which marks nuclei in both human and rat cells. The dotted orange line represents muscle and graft junction. E, epidermis; D, dermis; M, muscle (panniculus carnosus); and G, graft. The representative images are of at least 3 grafts/group. The scale bar represents 500 µm. Masson’s trichrome staining showed that the emulsified fat injections led to viable adipocyte architecture, with some collagen accumulation between and around typical fat vacuoles and confined in a fibrous capsule (Figure 3). In contrast, the free-lipid grafts formed large vacuoles without typical fat ultrastructure and were characterized by high collagen accumulation (Figure 3). The serum-saline mix injection did not yield to graft tissue formation as expected (Figure 3). Overall, the expression of collagen is correlated with the amount of free-lipid present in grafts, because the pure free-lipid resulted in high collagen accumulation, while the fat grafts expressed collagen only in the periphery. Figure 3. View largeDownload slide Masson’s trichrome staining of full-thickness tissue sections 45 days post-grafting. Collagen is stained blue, whereas cell cytoplasm and muscle are stained pink or red. E, epidermis; D, dermis; M, muscle (panniculus carnosus); and G, graft. The scale bar on the bottom right of the images represents 500 µm. The representative images are of at least 3 grafts/group. Figure 3. View largeDownload slide Masson’s trichrome staining of full-thickness tissue sections 45 days post-grafting. Collagen is stained blue, whereas cell cytoplasm and muscle are stained pink or red. E, epidermis; D, dermis; M, muscle (panniculus carnosus); and G, graft. The scale bar on the bottom right of the images represents 500 µm. The representative images are of at least 3 grafts/group. Inflammatory cell expression at 45 days was evaluated in the grafts stained with H&E, CD44, and CD68 (Figure 4). The inflammatory response consisted of a mononuclear infiltrate, predominantly macrophages, typically on the periphery of the grafts where blood vessels were most commonly visualized (Figure 4). Figure 4. View largeDownload slide Immune cell expression of full-thickness tissue sections 45 days post-grafting utilizing hematoxylin and eosin (H&E, top row). Immunofluorescence staining of CD44 (green), CD68 (red), and counter-stained against nuclei marker DAPI (4’, 6-diamidino-2-phenylindole, blue). M, muscle (panniculus carnosus); and G, graft. The scale bar represents 100 µm. The representative images are of at least the grafts/group. Figure 4. View largeDownload slide Immune cell expression of full-thickness tissue sections 45 days post-grafting utilizing hematoxylin and eosin (H&E, top row). Immunofluorescence staining of CD44 (green), CD68 (red), and counter-stained against nuclei marker DAPI (4’, 6-diamidino-2-phenylindole, blue). M, muscle (panniculus carnosus); and G, graft. The scale bar represents 100 µm. The representative images are of at least the grafts/group. Further studies were conducted to determine the optimal free-lipid to SVF ratio in the emulsified fat that yields viable graft formation. Compared with regular lipoaspirate or the SVF without the free-lipid group, adipose tissue with elastin accumulated more in and around the SVF graft enriched with free-lipid (5%-15%), resulting in arching of the above muscle layer (Figure 5). Although there is variation in different animals of the same group, the 10% free-lipid group resulted in the most consistent fat tissue formation (Supplementary Figure 1). Figure 5. View largeDownload slide (A) Masson’s trichrome and elastin immunofluorescence staining of stromal vascular fraction (SVF) with 0% to 15% free-lipid and whole lipoaspirate grafts at 45 days post-grafting. The trichrome stains the collagen blue, cell cytoplasm pink, and muscle red. Elastin (green) was counter-stained against nuclei marker DAPI (4’,6-diamidino-2-phenylindole, blue). E, epidermis; D, dermis; M, muscle (panniculus carnosus); and G, graft. The scale bar on the bottom right of the images represents 500 µm. (B) Percent area of collagen (trichrome staining, blue) and fluorescence intensity of elastin relative to nuclei in grafts. N = 3 grafts/group with no statistical significance (one-way ANOVA, P > 0.05). Figure 5. View largeDownload slide (A) Masson’s trichrome and elastin immunofluorescence staining of stromal vascular fraction (SVF) with 0% to 15% free-lipid and whole lipoaspirate grafts at 45 days post-grafting. The trichrome stains the collagen blue, cell cytoplasm pink, and muscle red. Elastin (green) was counter-stained against nuclei marker DAPI (4’,6-diamidino-2-phenylindole, blue). E, epidermis; D, dermis; M, muscle (panniculus carnosus); and G, graft. The scale bar on the bottom right of the images represents 500 µm. (B) Percent area of collagen (trichrome staining, blue) and fluorescence intensity of elastin relative to nuclei in grafts. N = 3 grafts/group with no statistical significance (one-way ANOVA, P > 0.05). To evaluate adipocyte expression compared with non-adipocyte cells in the SVF and free-lipid grafts, the sections were stained with human-specific adiponectin (adipocyte), mitochondria (all human cells), and nuclear marker DAPI (both rat and human cells, Figure 6). Human cell expression was significantly higher in the SVF with 10% free-lipid grafts (Figure 6B, mitochondria) compared with other groups. In contrast, adipocyte expression in the 10% free-lipid group was significantly higher than the SVF alone (no free-lipid) and lipoaspirate grafts but not in the 5% or 10% free-lipid groups (Figure 6B, adiponectin). Non-adipose cells were also higher in the 10% group compared with the lipoaspirate grafts (Figure 6, non-adipocytes). Figure 6. View largeDownload slide (A) Human adipocyte and non-adipocyte cell expression and localization in stromal vascular fraction (SVF) and free-lipid grafts. Adiponectin (red) and mitochondria (green) are utilized to evaluate adipocytes and all human cells (mitochondria specific staining, green), respectively. The scale bar represents 500 µm. (B) Fluorescence intensity of adipocytes and mitochondria relative to nuclei in grafts. Non-adipocytes were calculated by extracting adiponectin intensity from corresponding mitochondria relative to nuclei. N = 3 rats/group. Brackets represent statistical differences between groups (one-way ANOVA, P < 0.05). Figure 6. View largeDownload slide (A) Human adipocyte and non-adipocyte cell expression and localization in stromal vascular fraction (SVF) and free-lipid grafts. Adiponectin (red) and mitochondria (green) are utilized to evaluate adipocytes and all human cells (mitochondria specific staining, green), respectively. The scale bar represents 500 µm. (B) Fluorescence intensity of adipocytes and mitochondria relative to nuclei in grafts. Non-adipocytes were calculated by extracting adiponectin intensity from corresponding mitochondria relative to nuclei. N = 3 rats/group. Brackets represent statistical differences between groups (one-way ANOVA, P < 0.05). Finally, we evaluated local adipose tissue inflammation by CD44 staining16 and macrophage expression by CD68 staining.17 Although CD68 positive macrophage expression was similar between groups, CD44 expression was 2- to 5-fold greater in the free-lipid containing grafts (5%-15% and lipo) compared with SVF without free-lipid (Figure 7, bar graph). Whereas the fluorescence intensity of CD44 was similar in the 10% and 15% free-lipid groups, inflammatory response in the fat tissue was more visible in the 10% group (Figure 7). Figure 7. View largeDownload slide (A) Immune response evaluated by immunofluorescence staining of nanofat with 0% to 15% free-lipid and whole lipoaspirate grafts at 45 days post-grafting. Macrophage marker (CD68 [red]), immune activity in adipocytes (CD44 [green]), counter-stained against nuclei marker DAPI (4’,6-diamidino-2-phenylindole [blue]). The scale bar on the bottom right of the images represents 200 µm. SVF = stromal vascular fraction. (B) Relative fluorescence intensity of CD68 and CD44 calculated from 4 separate regions of an image. *, P < 0.05 (one-way ANOVA). N = 3/group. Figure 7. View largeDownload slide (A) Immune response evaluated by immunofluorescence staining of nanofat with 0% to 15% free-lipid and whole lipoaspirate grafts at 45 days post-grafting. Macrophage marker (CD68 [red]), immune activity in adipocytes (CD44 [green]), counter-stained against nuclei marker DAPI (4’,6-diamidino-2-phenylindole [blue]). The scale bar on the bottom right of the images represents 200 µm. SVF = stromal vascular fraction. (B) Relative fluorescence intensity of CD68 and CD44 calculated from 4 separate regions of an image. *, P < 0.05 (one-way ANOVA). N = 3/group. DISCUSSION Autologous fat grafting for small volume defects has long been a common tool for correcting these deficiencies; however, there remains a gap in understanding the fundamental mechanisms of fat grafting, the clinical application of these grafts, and the effects of the grafts on skin rejuvenation. In addition, multiple processing and handling methods of autologous fat yield to variations in fat graft retention and viability, which results in unpredictable clinical outcomes. The current study evaluated the skin effects of various human fat xenograft preparations in an athymic rat model to prevent host inflammatory response that compromises the survival of fat grafts, because the immune cells of the normal rat would kill living human graft cells. Our present findings in this athymic rat model suggest that certain components of the graft may be important to maximize the inflammatory process. The current study indicates that approximately 10% free-lipid within a fat graft that is primarily fractured adipocytes provides improved conditions for SVF differentiation into viable fat tissue as well as collagen and elastin production to provide mechanical support for the overlying skin. Among the proposed mechanisms for this skin rejuvenation process, the presence of adipose-derived stem cells appears important for successful lipofilling.18 Recent studies have shown that stem-cell-rich nanofat grafts yielded 2.5 times more weight than stem-cell-free grafts and were significantly less fibrous compared with the grafts without stem cells.18 Tonnard and colleagues13 suggest that stem cells are solely responsible for skin rejuvenation. Our study suggests that free-lipids or adipocyte fragments provide a milieu-enhancing cell viability and stem cell differentiation into adipocytes and other cells. In addition, this study implies that the free-lipid stimulates inflammation in the fat graft and increases collagen and elastin production; this is similar to the current study, in which the SVF with free-lipid yielded more CD44 and elastin expression (Figures 5 and 7), and the free-lipid alone grafts expressed more collagen accumulation and macrophages (Figures 3-4). This controlled pro-inflammatory response likely results in the desired response during skin rejuvenation. It is suggested that the inflammatory response to fat grafting plays a critical role in fat tissue rejuvenation by improving the overlying skin quality and elasticity.14 The presence of inflammation in fat tissue can activate macrophages, which is critical for tissue remodeling as they secrete the growth factor TGF-β, which promotes adipogenesis from MSCs.19 Furthermore, macrophages have been shown to secrete growth factors such as VEGF to stimulate angiogenesis, vascular remodeling, the secretion of ECM molecules, the structural repair of fat tissues, graft viability, and injury repair.14,20–22 Whereas the study utilized grafts from only 6 patients resulting in variations, subdermal grafting in the athymic rat model allowed the clean evaluation of graft formation; this is in contrast to intradermal injection, in which it would have been hard to rule out contributions of host cells, especially with ECM production and local immune response. The adipocyte expression is correlated with the presence of free-lipid, because the 10% free-lipid grafts resulted in significantly higher graft transformation to non-adipose cells and even more so to adipocytes, compared with SVF alone (Figure 6). This suggests that the free-lipid portion of lipoaspirate drives stem cells to differentiate into adipocytes and non-adipose cells (lesser extent) when adequate SVF is present. It is also interesting that the human graft did not yield to fat accumulation in the rat dermis, unlike subdermal fat tissue formation in the current study (Figure 2). This may be because of the fast clearance or low presence of the free-lipid in the dermis, thus providing inadequate nourishment to stem cells and leading to low cell viability. In contrast to the athymic rat model, the degree of immune response, ECM production, and cell differentiation may be different in the intradermal injection of human skin. It is possible that the free-lipid and SVF components can be cleared from the dermis by macrophages or other mechanisms leading to decreased cell viability and ECM secretion. Fat grafting has clinical potential when it is utilized to correct small volume defects such as soft tissue deficiencies and atrophic scars.23 Our study showed that the free-lipid portion of the fat graft helps promote even further tissue formation and ECM production, which may improve graft viability as well as skin changes in the context of atrophic acne scars or possibly aging or sun-damaged skin. The current study demonstrates that the SVF can be concentrated in a smaller volume (50 µL-100 µL), which may be useful for the robust correction of small volume defects. CONCLUSION This study attempted to better understand the beneficial effects of controlling the free-lipid-to-SVF ratio in fat grafting utilizing simple fat processing in a clinical setup. Through the efficient, volumetric, and pro-inflammatory response it elicits, fat grafting containing approximately 10% free-lipid may enhance and optimize micronized fat grafts, possibly enhancing viability and possible skin changes. This effect is modulated through establishing the proper milieu of tissue rejuvenation. Future studies should evaluate the viability of this approach in the clinical setting. Disclosures The authors declared no potential conflicts of interest with respect to the research, authorship, and publication of this article. Funding The authors received no financial support for the research, authorship, and publication of this article. REFERENCES 1. Cosmetic surgery national data bank statistics . Aesthet Surg J . 2016 ; 36 ( Suppl 1 ): 1 - 29 . 2. Sardesai MG , Moore CC . Quantitative and qualitative dermal change with microfat grafting of facial scars . Otolaryngol Head Neck Surg . 2007 ; 137 ( 6 ): 868 - 872 . Google Scholar Crossref Search ADS PubMed 3. Rigotti G , Marchi A , Galiè 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 ( 5 ): 1409 - 1422 ; discussion 1423. Google Scholar Crossref Search ADS PubMed 4. Akita S , Yoshimoto H , Ohtsuru A , Hirano A , Yamashita S . Autologous adipose-derived regenerative cells are effective for chronic intractable radiation injuries . Radiat Prot Dosimetry . 2012 ; 151 ( 4 ): 656 - 660 . Google Scholar Crossref Search ADS PubMed 5. James IB , Coleman SR , Rubin JP . Fat, stem cells, and platelet-rich plasma . Clin Plast Surg . 2016 ; 43 ( 3 ): 473 - 488 . Google Scholar Crossref Search ADS PubMed 6. Jones CM , Morrow BT , Albright WB , Long RE , Samson TD , Mackay DR . Structural fat grafting to improve reconstructive outcomes in secondary cleft lip deformity . Cleft Palate Craniofac J . 2017 ; 54 ( 1 ): 70 - 74 . Google Scholar Crossref Search ADS PubMed 7. Mojallal A , Shipkov C , Braye F , Breton P , Foyatier JL . Influence of the recipient site on the outcomes of fat grafting in facial reconstructive surgery . Plast Reconstr Surg . 2009 ; 124 ( 2 ): 471 - 483 . Google Scholar Crossref Search ADS PubMed 8. Gir P , Brown SA , Oni G , Kashefi N , Mojallal A , Rohrich RJ . Fat grafting: evidence-based review on autologous fat harvesting, processing, reinjection, and storage . Plast Reconstr Surg . 2012 ; 130 ( 1 ): 249 - 258 . Google Scholar Crossref Search ADS PubMed 9. Constantine RS , Harrison B , Davis KE , Rohrich RJ . Fat graft viability in the subcutaneous plane versus the local fat pad . Plast Reconstr Surg Glob Open . 2014 ; 2 ( 12 ): e260 . Google Scholar Crossref Search ADS PubMed 10. Cleveland EC , Albano NJ , Hazen A . Roll, spin, wash, or filter? Processing of lipoaspirate for autologous fat grafting: an updated, evidence-based review of the literature . Plast Reconstr Surg . 2015 ; 136 ( 4 ): 706 - 713 . Google Scholar Crossref Search ADS PubMed 11. Zuk PA , Zhu M , Ashjian P , et al. Human adipose tissue is a source of multipotent stem cells . Mol Biol Cell . 2002 ; 13 ( 12 ): 4279 - 4295 . Google Scholar Crossref Search ADS PubMed 12. Kølle SF , Fischer-Nielsen A , Mathiasen AB , et al. Enrichment of autologous fat grafts with ex-vivo expanded adipose tissue-derived stem cells for graft survival: a randomised placebo-controlled trial . Lancet . 2013 ; 382 ( 9898 ): 1113 - 1120 . Google Scholar Crossref Search ADS PubMed 13. Tonnard P , Verpaele A , Peeters G , Hamdi M , Cornelissen M , Declercq H . Nanofat grafting: basic research and clinical applications . Plast Reconstr Surg . 2013 ; 132 ( 4 ): 1017 - 1026 . Google Scholar Crossref Search ADS PubMed 14. Ogle ME , Segar CE , Sridhar S , Botchwey EA . Monocytes and macrophages in tissue repair: implications for immunoregenerative biomaterial design . Exp Biol Med (Maywood) . 2016 ; 241 ( 10 ): 1084 - 1097 . Google Scholar Crossref Search ADS PubMed 15. Shukla L , Morrison WA , Shayan R . Adipose-derived stem cells in radiotherapy injury: a new frontier . Front Surg . 2015 ; 2 : 1 . Google Scholar Crossref Search ADS PubMed 16. Kodama K , Horikoshi M , Toda K , et al. Expression-based genome-wide association study links the receptor CD44 in adipose tissue with type 2 diabetes . Proc Natl Acad Sci U S A . 2012 ; 109 ( 18 ): 7049 - 7054 . Google Scholar Crossref Search ADS PubMed 17. Holness CL , Simmons DL . Molecular cloning of CD68, a human macrophage marker related to lysosomal glycoproteins . Blood . 1993 ; 81 ( 6 ): 1607 - 1613 . Google Scholar PubMed 18. Moseley TA , Zhu M , Hedrick MH . Adipose-derived stem and progenitor cells as fillers in plastic and reconstructive surgery . Plast Reconstr Surg . 2006 ; 118 ( 3 Suppl ): 121S - 128S . Google Scholar Crossref Search ADS PubMed 19. Margoni A , Fotis L , Papavassiliou AG . The transforming growth factor-beta/bone morphogenetic protein signalling pathway in adipogenesis . Int J Biochem Cell Biol . 2012 ; 44 ( 3 ): 475 - 479 . Google Scholar Crossref Search ADS PubMed 20. Godwin JW , Pinto AR , Rosenthal NA . Chasing the recipe for a pro-regenerative immune system . Semin Cell Dev Biol . 2017 ; 61 : 71 - 79 . Google Scholar Crossref Search ADS PubMed 21. Yi CG , Xia W , Zhang LX , et al. VEGF gene therapy for the survival of transplanted fat tissue in nude mice . J Plast Reconstr Aesthet Surg . 2007 ; 60 ( 3 ): 272 - 278 . Google Scholar Crossref Search ADS PubMed 22. Nishimura T , Hashimoto H , Nakanishi I , Furukawa M . Microvascular angiogenesis and apoptosis in the survival of free fat grafts . Laryngoscope . 2000 ; 110 ( 8 ): 1333 - 1338 . Google Scholar Crossref Search ADS PubMed 23. Coleman SR , Katzel EB . Fat grafting for facial filling and regeneration . Clin Plast Surg . 2015 ; 42 ( 3 ): 289 - 300 , vii. Google Scholar Crossref Search ADS PubMed © 2018 The American Society for Aesthetic Plastic Surgery, Inc. Reprints and permission: journals.permissions@oup.com This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices)
Journal
Aesthetic Surgery Journal – Oxford University Press
Published: Nov 12, 2018