Autologous free white adipose tissue (WAT) grafts are commonly used in plastic surgery for reconstruction of soft-tissue defects.1 Novel experimental therapies include softening and contour of scars and other fibrotic conditions.2–4 However, the basic biology of fat transfer surgery and the antifibrotic effects are still poorly understood. Many of the effects are thought to be mediated by the immunomodulatory properties of adipose-derived stem cells, which are present in the transferred adipose tissue.5 They have been shown to reduce proinflammatory cytokine (IL-1, IL-17) levels and increase anti-inflammatory protein (IL-10) levels in mice.6 Only a few experimental reports describe the effects of fat grafting in muscle tissue7 , 8 although fat grafting is routinely performed in large volumes for example in pectoralis and latissimus dorsi muscles in breast reconstruction and augmentation.9 , 10
The type of adipose tissue plays a great role in obesity and metabolic disorders such as type 2 diabetes. It is known that there are 2 types of adipose tissue in mammals: thermogenically active brown adipose tissue (BAT) and WAT, which is mainly involved in storing excess energy.11 Typically BAT contains adipocytes with small multilocular lipid droplets, and numerous mitochondria with uncoupling protein 1 (UCP1), the protein responsible for regulation of thermogenesis and characteristic feature for BAT.12 , 13 Previously, BAT was thought to exist only in small mammals and infants, but during the past decade, several studies have demonstrated significant amounts of functional BAT also in adults.14 Interestingly, also adult humans have inducible brown-like adipocytes called beige (or brite) adipose tissue, which consists of brown-like adipocytes among WAT bringing about similar characteristics in energy metabolism as BAT.15–17 Recent evidence suggest that beige adipocytes derive from the same mesenchymal stem cells but different precursor cells as WAT18 and begin to differentiate due to environmental stimuli, such as changes in temperature and nutrition.19
We have previously used an experimental model to evaluate the vascularization, survival, and metabolic changes after free fat transfer using18F-fluorodeoxyglucose (FDG) PET/CT imaging described by Tervala et al.20 Results showed that transfer of the metabolically inactive (epididymal) fat into a new environment modulates the metabolic activity of the fat grafts to resemble the situation in the recipient site. We also noted metabolic differences in subcutaneous, visceral, and epididymal WAT depots. In this study, we wanted to investigate whether the metabolic increase of the transplant can be a result of “browning” of the transplanted WAT and whether it was donor-site specific. In this study, we examined the metabolic and histological characteristics and BAT marker Ucp1 expression in different types of WAT grafts placed either subcutaneously or intramuscularly. The purpose of the study was to investigate whether fat grafting can induce browning of WAT, especially when fat is transferred to a metabolically more active area such as muscle tissue, and determine the potential benefits of fat grafting in the surgical treatment of metabolic disorders.
MATERIALS AND METHODS
Permissions for the animal studies were obtained from the National Animal Experiment Board of Finland. We used 3 different types of WAT as free fat grafts transferred into subcutaneous and muscle tissue, as described in Figure 1. According to the WAT type, 3 groups of graft mice were formed and maintained during the experiment: epididymal (n = 6), visceral (n = 5), and subcutaneous group (n = 5). A control group (n = 5) was used as a reference to determine the basic muscle, BAT, and WAT metabolism. C57BL/6 mice (C57BL/6NCrl, Harlan, The Netherlands) were killed and used as donor mice for harvesting WAT. Athymic nude mice (Hsd:Athymic Nude-Foxn1nu, Harlan, The Netherlands) with inhibited immune system were used as recipients to avoid graft rejection response. All mice were anesthetized with xylazine hydrochloride 5 mg/kg (Rompun 20 mg/mL, Bayer Animal Health, Leverkusen, Germany) and ketamine 70 mg/kg (Ketalar, Hameln Pharmaceuticals, Hameln, Germany). For analgesia, buprenorphine hydrochloride 0.075 mg/kg (Temgesic, RB Pharmaceuticals Limited, Slough, Berkshire, Great Britain) was administered to recipient mice during 2 days postoperatively.
After harvesting, the excised grafts were weighted and subsequently placed under the skin of forehead to subcutaneous space and among muscle tissue in hind leg of the recipient mice (n = 16). Into forehead, the fat was placed as a whole piece, and into leg with a 14 G cannula after mechanical disruption combined with a small amount of saline. The average volume of grafted fat to forehead was 0.25 ± 0.09 mL and 0.13 ± 0.05 mL to leg, respectively. Wounds were closed using 5-0 Polysorb sutures (Covidien, Dublin, Ireland).
In Vivo FDG-PET/CT Imaging and Data Analysis
Metabolic activity of fat grafts was examined in mice at 4- and 12-week time points by PET/CT (Siemens Medical Solutions USA, Knoxville, Tenn.) using 18F-FDG, a glucose analogue known to accumulate to metabolically active tissues. Metabolism of BAT increases in cold temperature, and to stimulate the activity of BAT and induce thermogenesis, PET/CT scan was performed on 2 consecutive days: first with cold exposure (26–28°C) and the second with warm exposure (36–38°C).21 FDG was given approximately 5 MBq into the tail vein and the 2-step whole-body PET/CT scan was performed at fasting state (4 h) during isoflurane gas anesthesia on. The cold exposure was accomplished by exposing mice to cold air by circulating propylene glycol through an animal holder using a heating circulator (S-12 Julabo, Seelbach, Germany) with simultaneous temperature monitoring. The warm exposure was carried out in normal room temperature by using a heating pad. At 4-week time point, PET/CT was performed only with warm exposure, and at 12-weeks with both cold and warm scans.
FDG images were reconstructed as described by Tervala et al.20 Volumes of interest (VOIs) were drawn on the forehead subcutaneous fat graft and leg region intramuscular fat graft by using either radioactivity uptake or the computed tomography template as an anatomical reference. In the leg where grafts were smaller and less visible than that of the forehead, VOIs were drawn to cover the whole leg. VOIs were drawn also on a part of liver, interscapular BAT, and contralateral hind leg (control muscle) as a reference. Standardized uptake value (SUV) was defined as the ratio of FDG radioactivity (Bq) per milliliter (mL) of tissue to the radioactivity in the injected dose corrected by decay and by animal body weight. The SUV of fat grafts were compared with that of liver, and results expressed as fat-to-liver uptake ratios (SUV of fat/SUV of liver).
Ex Vivo Radioactivity Analysis of Tissue Samples
Immediately after the last follow-up scan at 12-week time point, the graft mice were killed for ex vivo tissue sample analysis. Samples were measured for 18F-radioactivity in a gamma counter (Wizard2 3”, PerkinElmer, Turku, Finland) and expressed as the percentage of injected dose per gram tissue (% ID/g). Thereafter, samples were fixed in formalin for histologic examination. A separate tissue sample for RNA analysis was snap frozen in liquid nitrogen and stored in −70°C.
Gene Expression Analysis
Tissue samples were homogenized in QIAzol Lysis Reagent (Qiagen), and total RNA was extracted according to the manufacturer’s recommendations. RNA was cleaned up from the aqueous phase of the extracts using the RNeasy Micro Kit (Qiagen) according to the manufacturer’s recommendations and included an on-column DNase I treatment step. cDNA synthesis was performed using the first Strand cDNA Synthesis Kit for real-time polymerase chain reaction (PCR) (AMV) (Roche). Quantitative real-time PCR was performed on a ViiA7 instrument (Applied Biosystems) using Power SYBR Green PCR Master Mix (Applied Biosystems). The standard curve method with Rplp0 as the normalizing gene was used to determine relative Ucp1 gene expression levels. Primer sequences are supplied in Supplementary Table 1 (see table, Supplementary Digital Content 1, which displays primer sequences, http://links.lww.com/PRSGO/A793).
Histology and Immunohistochemistry
Tissue samples fixed overnight in formalin were embedded in paraffin. Sections of 4 μm thickness were cut and stained with hematoxylin and eosin for the evaluation and grading (0–5) or percentage (0–100%) of adipose tissue survival, fibrosis, cystic degeneration, presence of multilocular BAT morphology, and degree of inflammation. To confirm the Ucp1 gene expression results at protein level, UCP1 immunohistochemical staining was performed. Samples were stained with rabbit antimouse anti-UCP1 (dilution 1:1,000, Abcam, United Kingdom). An horseradish peroxidase (HRP)-conjugated goat-antirabbit immunoglobulin G (IgG) polyclonal (ImmunoLogic, Netherlands) was used as the secondary antibody.
All results were analyzed using SPSS (IBM SPSS Statistics IBM SPSS Statistics for Macintosh, version 23.0. Armok, NY: IBM Corp.). Groups were compared using 1-way ANOVA, and pairwise comparisons were made using the Kruskall-Wallis test or Student’s t test. Statistical significance was set at P ≤ 0.05.
In Visual Analysis of FDG-PET/CT Imaging, Most Grafts Were Visible and Showed FDG Uptake
At 4-week time point, all subcutaneous grafts at the forehead region were visible and showed FDG uptake, and most grafts (63%) showed increase in volume (mean graft volume 0.32 ± 0.19 mL, median 0.30 mL). Visual FDG uptake in the intramuscular grafts at leg region was seen in 11 of 16 mice (68.8%) evenly distributed in all groups. Intramuscular graft volume was not measurable with PET/CT due to small amount of transferred fat.
At 12-week time point, the FDG uptake remained visible in 10 of 16 subcutaneous grafts during follow-up. Poorest survival of grafts was in the visceral group, where only 1 graft was visible. Graft volumes showed great variability (mean 0.53 ± 0.61 mL, median 0.24 mL). Eight intramuscular grafts showed visual FDG uptake with even distribution between groups (Fig. 2).
FDG-PET/CT Quantitative Analysis Showed High Uptake in Both Graft Regions and BAT But Low Uptake in Control WATs
The FDG uptake in VOI was expressed as fat-to-liver ratio shown in Table 1. PET/CT scans at both 4-week and 12-week time points showed increased FDG uptake subcutaneous grafts without significant difference between the groups (P = 0.069). In intramuscular grafts, FDG uptake was greater in all graft groups compared with control leg (P = 0.019). There was no statistical difference between fat-to-liver ratios in different time points at 4 and 12 weeks (P = 0.421). Overall, FDG uptake in all regions was higher in warm temperature scans (P < 0.001; Fig. 3A). Ex vivo tissue samples were also measured for 18F-FDG uptake (Fig. 3B) showing high uptake in both graft regions and BAT but low uptake in control WATs.
Gene Expression Analysis Showed Elevated Ucp1 Expression in 4 out of 15 Intramuscular Grafts
Fifteen intramuscular and 13 subcutaneous graft samples were analyzed in addition to control BAT and WAT samples (Fig. 4). The highest Ucp1/Rplp0 ratio of all fat tissue samples was used as a reference, and results are expressed as percentage (%) of the reference value. The Ucp1 expression was relatively low in all subcutaneous grafts (mean 0.0015% ± 0.0022%), whereas the Ucp1 level in intramuscular grafts was elevated in all groups (mean 2.51% ± 7.65%), but the comparison did not reach statistical significance (P = 0.25). The mean Ucp1 expression in the control samples was 0.0015% ± 0.00077% in epididymal fat, 0.017% ± 0.025% in visceral fat, and 0.12% ± 0.10% in subcutaneous fat. Average BAT Ucp1 level was 63% ± 12%. In individual analysis, 4 intramuscular fat grafts showed significantly elevated expression of Ucp1 (mean, 9.4% ± 14%) compared with control WAT and presented later in this section.
Histological Analysis Revealed Brown Adipose Tissue-like, UCP1-positive Multilocular Fat in 4 of 15 Intramuscular Grafts
The histological analysis did not make any difference between the types of WAT used in grafting, and therefore comparison was performed mainly between the graft locations. The average percentage of survived fat was 23% ± 28%, and there seemed to be a trend toward a greater adipose tissue percentage in the intramuscular grafts (P = 0.12). No significant statistical difference was observed with respect to the degree of inflammation and fibrosis between graft locations (P = 0.92 and P = 0.28, respectively), but cystic degeneration was significantly lower in intramuscular grafts (P < 0.001). BAT-like multilocular fat was seen only in intramuscular grafts (mean, 11% ± 21%), but also in control WAT in some degree (7% ± 14%; Fig. 5). However, UCP1 staining was negative and Ucp1 gene expression low regarding these control samples. When examining samples individually, multilocular fat was found in 4 intramuscular grafts (2 in the epididymal group, 1 in the visceral group, and 1 in the subcutaneous group), and these samples showed also positive UCP1 staining and elevated Ucp1 gene and protein expression as a difference to control samples. Detailed results of histological analysis by groups are shown in Table 2.
In Individual Sample Analysis, 4 of 15 Intramuscular Fat Grafts Showed Browning Toward Beige Fat
According to the results, the same 4 intramuscular fat grafts were characterized by multilocular fat droplets and increased Ucp1 gene and protein expression, both typical for BAT, indicating metabolic and morphologic change toward beige adipose tissue. As seen in Figure 5, with hematoxylin and eosin (HE) staining beige fat was characterized by both multilocular BAT-like fat and large lipid droplets similar WAT, but other grafts showed mostly original WAT-like fat with large oil cysts. To confirm the presence of beige fat, we examined the samples also with UCP1 staining, which showed positively stained tissue samples. Also some WAT samples showed multilocular fat but UCP1 staining remained negative. The amount of the multilocular fat varied in the beige samples (60%, 50%, 10%, and 40%). The beige fat was well preserved in the target region (fat cells 76 ± 8 % versus 15 ± 18 % in other fat grafts, P < 0.001), and there was less inflammation (grading 0.25 ± 0.5 versus 2.96 ± 1.30, P < 0.001), fibrosis (0.5 ± 0.6 versus 3.1 ± 1.2, P < 0.001) and cystic degeneration (0 in beige grafts, others 2.0 ± 1.7, P = 0.03). Glucose uptake in beige grafts was low: the mean ex vivo FDG uptake was 3.5 ± 1.5 %ID/g in the beige grafts and 11.1 ± 6.7 %ID/g in others (P = 0.140; Fig. 3B). Mean Ucp1 expression was 9.4% ± 14 % for beige grafts and 0.002% ± 0.004% for other grafts, respectively, which makes the Ucp1 expression in beige grafts 4,700-fold greater compared with the other grafts (P < 0.001) and from 430 to 1,383-fold greater than in the respective control fat (Fig. 4).
In this article, we aimed to investigate the metabolic adaptation of WAT grafts in subcutaneous and muscle tissue, as the metabolic and morphologic changes in fat grafts are still poorly understood despite the fact that lipotransfer is commonly used in plastic surgery. Our results show a novel phenomenon: browning of intramuscularly placed fat grafts. This was demonstrated with HE and UCP1-stained histology samples showing multilocular lipid droplets and an increase in Ucp1 gene and protein expression. These grafts were also of good quality with limited inflammation, fibrosis, and cystic degeneration.
This finding presents an interesting outcome of a common procedure in the field of plastic surgery. Obesity and the large amount of inactive fat are major factors in metabolic disorders. By changing the metabolism of fat into a more active form, fat grafting could have beneficial effects also on glucose tolerance and body weight. To this point, many experimental studies have shown that fat has effects on glucose balance, but different depots of WAT seem to have different metabolic characteristics. Previous reports have shown that the fatty acid metabolism is lower in epididymal than in subcutaneous WAT.22 It has also been reported that transfer of epididymal or inguinal WAT into the visceral cavity improves glucose tolerance,23 decreases the total amount of fat and weight and insulin resistance in mice.24 Especially the transfer of subcutaneous fat into the visceral cavity reduces the amount of several proinflammatory cytokines (TNF alfa, IL-17, IL-12) in plasma.25
A limited degree of inflammation may be 1 factor allowing browning to initiate. A recent study has shown that inflammation of WAT is associated with diminished generation of Ucp1 expressing beige adipocytes26 giving a possible explanation why browning occurred only in limited amount of samples in our study. Unlike previous studies have shown,27 , 28 also interscapular BAT showed increased glucose (FDG) uptake during warm exposure than in cold exposure. However, recent reports demonstrate that FDG uptake can be increased in BAT of UCP1-deficient mice suggesting also UCP1 independent mechanisms for the increased uptake.29
The amount of BAT precursors in fat grafts also varies, and therefore some grafts may be more prone to browning than others. According to previous studies, the metabolic adaptation of white adipose cells to beige cells through direct transformation is regulated by many factors.16 , 30 , 31 Beige adipocytes, at least in the subcutaneous depot, have been shown to arise from a precursor population rather than from preexisting adipocytes.32 The beige adipocyte precursor population is different from the white precursor population,18 and beige adipocytes have been shown to be most abundant in the inguinal WAT, which is a major subcutaneous depot in rodents.33
An autocrine/paracrine mechanism between muscle and adipose cells may also be the cause behind browning. Myocytes secrete irisin, and its concentrations increase in mice and humans by exercise training and it has been shown to stimulate the browning of WAT through specific actions on the beige preadipocyte population.34 An advantage in muscle tissue is also the abundant vascular bed improving the survival of fat graft.
As a conclusion, this study suggests a novel finding: fat grafting can induce browning of the graft. Subcutaneous and intramuscular grafting might have different outcomes as browning of fat was only seen in grafts placed in muscle tissue. It is known that many factors can initiate browning, and external manipulation (liposuction and lipotransfer) may be 1 of these factors. Also transferring WAT to metabolically more active muscle tissue may bring about metabolic adaptation in the fat. Therefore, fat transfer may have beneficial effects on body metabolism by increasing the amount of metabolically active tissue. Clinical research is still needed to investigate the metabolic changes in autologous fat grafting in humans, but the potential browning and metabolic activation of WAT opens up new research areas in exploiting fat grafting in metabolic diseases.
The authors thank Marko Vehmanen, Elisa Riuttala, Aake Honkaniemi, and Erica Nyman for excellent technical assistance.
1. Khouri RK Jr, Khouri RK. Current clinical applications of fat grafting. Plast Reconstr Surg. 2017;140:466e–486e.
2. Condé-Green A, Marano AA, Lee ES, et al. Fat grafting and adipose-derived regenerative cells in burn wound healing and scarring: a systematic review of the literature. Plast Reconstr Surg. 2016;137:302–312.
3. Lolli P, Malleo G, Rigotti G. Treatment of chronic anal fissures and associated stenosis by autologous adipose tissue transplant: a pilot study. Dis Colon Rectum. 2010;53:460–466.
4. Ulrich D, Ulrich F, van Doorn L, et al. Lipofilling of perineal and vaginal scars: a new method for improvement of pain after episiotomy and perineal laceration. Plast Reconstr Surg. 2012;129:593e–594e.
5. Mizuno H. The potential for treatment of skeletal muscle disorders with adipose-derived stem cells. Curr Stem Cell Res Ther. 2010;5:133–136.
6. Siniscalco D, Giordano C, Galderisi U, et al. Long-lasting effects of human mesenchymal stem cell systemic administration on pain-like behaviors, cellular, and biomolecular modifications in neuropathic mice. Front Integr Neurosci. 2011;5:79.
7. Karacaoglu E, Kizilkaya E, Cermik H, et al. The role of recipient sites in fat-graft survival: experimental study. Ann Plast Surg. 2005;55:63–68; discussion 68.
8. Shi Y, Yuan Y, Dong Z, et al. The fate of fat grafts in different recipient areas: subcutaneous plane, fat pad, and muscle. Dermatol Surg. 2016;42:535–542.
9. Rosing JH, Wong G, Wong MS, et al. Autologous fat grafting for primary breast augmentation: a systematic review. Aesthetic Plast Surg. 2011;35:882–890.
10. Sinna R, Delay E, Garson S, et al. Breast fat grafting (lipomodelling) after extended latissimus dorsi flap breast reconstruction: a preliminary report of 200 consecutive cases. J Plast Reconstr Aesthet Surg. 2010;63:1769–1777.
11. Cypess AM, Lehman S, Williams G, et al. Identification and importance of brown adipose tissue in adult humans. N Engl J Med. 2009;360:1509–1517.
12. Vijgen G, van Marken Lichtenbelt W. Brown adipose tissue: clinical impact of a re-discovered thermogenic organ. Front Biosci (Elite Ed). 2013;5:823–833.
13. Lidell ME, Betz MJ, Dahlqvist Leinhard O, et al. Evidence for two types of brown adipose tissue in humans. Nat Med. 2013;19:631–634.
14. Virtanen KA, Lidell ME, Orava J, et al. Functional brown adipose tissue in healthy adults. N Engl J Med. 2009;360:1518–1525.
15. Wu J, Boström P, Sparks LM, et al. Beige adipocytes are a distinct type of thermogenic fat cell in mouse and human. Cell. 2012;150:366–376.
16. Harms M, Seale P. Brown and beige fat: development, function and therapeutic potential. Nat Med. 2013;19:1252–1263.
17. Sidossis L, Kajimura S. Brown and beige fat in humans: thermogenic adipocytes that control energy and glucose homeostasis. J Clin Invest. 2015;125:478–486.
18. Rosen ED, Spiegelman BM. What we talk about when we talk about fat. Cell. 2014;156:20–44.
19. Park A, Kim WK, Bae KH. Distinction of white, beige and brown adipocytes derived from mesenchymal stem cells. World J Stem Cells. 2014;6:33–42.
20. Tervala TV, Grönroos TJ, Hartiala P, et al. Analysis of fat graft metabolic adaptation and vascularization using positron emission tomography-computed tomographic imaging. Plast Reconstr Surg. 2014;133:291–299.
21. Cannon B, Nedergaard J. Nonshivering thermogenesis and its adequate measurement in metabolic studies. J Exp Biol. 2011;214:242–253.
22. del Mar Romero M, Fernández-López JA, Esteve M, et al. Site-related white adipose tissue lipid-handling response to oleoyl-estrone treatment in overweight male rats. Eur J Nutr. 2009;48:291–299.
23. Foster MT, Shi H, Softic S, et al. Transplantation of non-visceral fat to the visceral cavity improves glucose tolerance in mice: investigation of hepatic lipids and insulin sensitivity. Diabetologia. 2011;54:2890–2899.
24. Tran TT, Yamamoto Y, Gesta S, et al. Beneficial effects of subcutaneous fat transplantation on metabolism. Cell Metab. 2008;7:410–420.
25. Hocking SL, Stewart RL, Brandon AE, et al. Subcutaneous fat transplantation alleviates diet-induced glucose intolerance and inflammation in mice. Diabetologia. 2015;58:1587–1600.
26. Chung KJ, Chatzigeorgiou A, Economopoulou M, et al. A self-sustained loop of inflammation-driven inhibition of beige adipogenesis in obesity. Nat Immunol. 2017;18:654–664.
27. Park JW, Jung KH, Lee JH, et al. 18F-FDG PET/CT monitoring of β3 agonist-stimulated brown adipocyte recruitment in white adipose tissue. J Nucl Med. 2015;56:153–158.
28. David JM, Chatziioannou AF, Taschereau R, et al. The hidden cost of housing practices: using noninvasive imaging to quantify the metabolic demands of chronic cold stress of laboratory mice. Comp Med. 2013;63:386–391.
29. Hankir MK, Kranz M, Keipert S, et al. Dissociation between brown adipose tissue 18F-FDG uptake and thermogenesis in uncoupling protein 1-deficient mice. J Nucl Med. 2017;58:1100–1103.
30. Fu J, Li Z, Zhang H, et al. Molecular pathways regulating the formation of brown-like adipocytes in white adipose tissue. Diabetes Metab Res Rev. 2015;31:433–452.
31. Jeremic N, Chaturvedi P, Tyagi SC. Browning of white fat: novel insight into factors, mechanisms, and therapeutics. J Cell Physiol. 2017;232:61–68.
32. Wang QA, Tao C, Gupta RK, et al. Tracking adipogenesis during white adipose tissue development, expansion and regeneration. Nat Med. 2013;19:1338–1344.
33. Vitali A, Murano I, Zingaretti MC, et al. The adipose organ of obesity-prone C57BL/6J mice is composed of mixed white and brown adipocytes. J Lipid Res. 2012;53:619–629.
34. Boström P, Wu J, Jedrychowski MP, et al. A PGC1-α-dependent myokine that drives brown-fat-like development of white fat and thermogenesis. Nature. 2012;481:463–468.
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Copyright © 2018 The Authors. Published by Wolters Kluwer Health, Inc. on behalf of the American Society of Plastic Surgeons. All rights reserved.