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. Author manuscript; available in PMC: 2020 Apr 1.
Published in final edited form as: Plast Reconstr Surg. 2019 Apr;143(4):1067–1075. doi: 10.1097/PRS.0000000000005483

Fat Grafting into Younger Recipients Improves Volume Retention in an Animal Model

Natalie N Chung 1,#, Ryan C Ransom 1,#, Charles P Blackshear 1, Dre M Irizarry 1, Derek Yen 1, Arash Momeni 1, Gordon K Lee 1, Dung H Nguyen 1, Michael T Longaker 1,2, Derrick C Wan 1,#
PMCID: PMC6438719  NIHMSID: NIHMS1515353  PMID: 30730498

Abstract

Background:

Soft tissue deficits associated with various craniofacial anomalies can be addressed by fat grafting, although outcomes remain unpredictable. Furthermore, consensus does not exist for timing of these procedures. While some advocate approaching soft tissue reconstruction after the underlying skeletal foundation has been corrected, other studies have suggested earlier grafting may exploit a younger recipient niche that is more conducive for fat graft survival. As there is a dearth of research investigating effects of recipient age on fat graft volume retention, this study compared the effectiveness of fat grafting in younger versus older animals through a longitudinal, in vivo analysis.

Methods:

Human lipoaspirate from three healthy female donors was grafted subcutaneously over the calvarium of immunocompromised mice. Volume retention over 8 weeks was evaluated using micro-computed tomography in three experimental ages – 3-weeks old, 6-months old, and 1-year old. Histology was performed on explanted grafts to evaluate graft health and vascularity. Recipient site vascularity was also evaluated by confocal microscopy.

Results:

Greatest retention of fat graft volume was noted in the youngest group compared to both older groups (*p < 0.05) at 6 and 8 weeks following grafting. Histological and immunohistochemical analyses revealed that improved retention in younger groups was associated with greater fat graft integrity and more robust vascularization.

Conclusion:

Our study provides evidence that grafting fat into a younger recipient site correlates with improved volume retention over time, suggesting that beginning soft tissue reconstruction with fat grafting in patients at an earlier age may be preferable to late correction.

Keywords: Fat Grafting, Recipient Age, Juvenile Mouse, Soft Tissue

Introduction

Soft tissue defects in the craniofacial region can arise from trauma, tumor removal, infection, congenital disorders, and connective tissue disease, and present a common reconstructive challenge for plastic surgeons. Autologous fat transfer has become an increasingly prevalent method to restore volume in soft tissue defects, owing to its high safety profile, low donor site morbidity, and relative abundance (1–5). However, fat grafting remains suboptimal, with variable volume retention reported (6–10). Furthermore, optimal timing of these procedures in the pediatric population has not yet been standardized.

Traditionally, reconstruction of large defects has begun with restoration of underlying bony structure, followed later by soft tissue correction, using a combination of flaps, synthetic fillers, and/or autologous fat transfer (11–14). However, some surgeons suggest that beginning autologous fat transfer earlier may take advantage of the higher reparative potential of a younger niche, which could produce more stable volume outcomes. Tanna et al. found that serial fat grafting, started early in the process of reconstruction, can lead to volumetric results comparable to those observed with use of flaps, along with the added benefit of lower complications and lower overall operating time (15). Furthermore, a study examining surgical timing for reconstruction in Poland syndrome found that starting autologous fat transfer earlier, during the period of growth, allowed for greater body image stabilization with more frequent, but less invasive procedures (16). The principle that younger recipient sites may retain volume better has also been supported by the finding that overcorrection during autologous fat transfer, a common technique to counter unpredictable volume loss in adults, can in some pediatric cases lead to the complication of “sustained overcorrection,” resulting in larger than intended volumes (17).

While the literature suggests fat grafting at earlier ages may be reasonable, there is a dearth of exploration into how age of the recipient specifically influences effectiveness of fat grafting. Fat graft retention might be improved by grafting into younger recipient sites, which would be particularly relevant for reconstructions that require multiple procedures over the course of months to years. To this end, we employed a longitudinal in vivo model to measure how fat graft volume retention differs in mice groups of various ages. Histology was also performed on explanted fat grafts to explore vascularization of the graft and compare how the architecture of the fat graft changes with age of the recipient site.

Methods

Fat Grafting

All studies were conducted in accordance with Stanford University animal use guidelines (APLAC Protocol #31212). Crl:Nu-Foxn1nu immunocompromised mice (Charles River Laboratories; Wilmington, MA) were used for in vivo analysis to minimize immunologic response to human adipose tissue (18). Three mouse age groups (n=15 per group) were chosen to reflect different clinical ages: Group A = 3-weeks old (pre-adolescent), Group B = 6-months old (middle-age adult), and Group C = 1-year old (geriatric). Human lipoaspirate from three healthy female donors with no other medical comorbidities, ages 39- to 45-years old with BMI ranging from 26 to 29, was collected under an approved IRB protocol #2188. Lipoaspirate was allowed to settle for 15 minutes, and then oil and blood layers were removed by vacuum aspiration. The remaining fat layer was centrifuged at 1300 rcf for 3 minutes at 4°C. Remaining oil and blood was again removed and the resultant fat was transferred into 1cc syringes for injection. A 14-gauge Luer-lock cannula was used to create a subcutaneous tunneled pocket over the calvarium of each mouse, and 150 microliters of fat was subsequently injected with skin sutured closed (Figure 1).

Figure 1.

Figure 1.

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Three age groups of immunocompromised mice (Group A = 3-weeks old, Group B = 6-months old, Group C = 1-year old) with 150 microliter fat grafts placed subcutaneously over the calvarium.

Fat Graft Volume Retention

Fat graft volume was measured two days after fat graft injection for baseline volume measurements and then every two weeks over 8 weeks using a MicroCAT-II in vivo X-Ray micro-CT scanner (Imtek, Inc.; Knoxville, TN). Cubic-spline interpolation was used to reconstruct three-dimensional images of grafts and measure volume changes over time (18), as previously described. All reconstructions were performed by a single investigator (N.N.C.) to avoid inter-observer variability.

Fat Graft Histological Analysis

Three fat grafts were explanted at two, four, six, and eight weeks from each age group for histological analysis. Samples were fixed, embedded in paraffin, sectioned, and stained with hematoxylin and eosin (H&E). Bright-field images were obtained with a 10x objective. Five sections per graft were analyzed at each time point by five blinded, independent reviewers, who assessed metrics of fat health/tissue integrity, as well as injury including formation of cysts/vacuoles, inflammation, and fibrosis, as previously described (19, 20). Sections were also stained using fluorescent immunohistochemical anti-CD31 antibody (Ab28364 1:100 dilution; Abcam, Cambridge, MA) to assess for vascularity. Fluorescence was quantified using ImageJ analysis.

Skin Staining and Confocal Imaging

Skin biopsies from three animals in Groups A, B, and C, were obtained prior to fat graft placement. Each sample was then stained with Isolectin endothelial cell stain (ThermoFisher Scientific; Waltham, MA) (20 μl in 2 ml PBS for a 1:100 dilution) and FITC anti-mouse CD31 antibody (BioLegend; San Diego, CA) (1:100 dilution) for one hour at room temperature. After a brief PBS wash, the samples were then incubated at room temperature in a 1:2000 dilution of Hoechst 33342 (ThermoFisher Scientific) stain in PBS for 30 minutes. The samples were finally washed twice with PBS and whole-mounted to glass microscope slides, as adapted from the protocol described by Berry et al. (21). Laser scanning confocal microscopy was performed using a Leica TCS SP8 X confocal microscope (Leica Microsystems; Wetzlar, Germany) with an objective lens (10× HC PL APO, air, N.A. 0.40). Three-dimensional volume rendering of z-stacks was performed on Imaris (Bitplane AG; Zurich, Switzerland) and quantification was performed using the Isosurface Rendering Statistics tool.

Statistical Analysis

Data are presented as medians ± range. Multiple group comparisons were performed using a nonparametric Kruskal-Wallis with post-hoc Dunn’s testing. All analyses were performed using GraphPad Prism (GraphPad Software, La Jolla, CA). A p-value <0.05 was considered statistically significant.

Results

Fat Graft Volume Retention

Three-dimensional volumetric analysis of fat grafts demonstrated that younger recipient age correlated positively with fat graft retention over 8 weeks (Figure 2). Results were statistically significant at both 6- and 8-week time points (**p < 0.01). At 6 weeks, the youngest group, Group A, demonstrated the greatest volume retention (79.7%, range 78.6 to 83.4%), followed by Group B (56.9%, range 52.7 to 66.2%), and finally Group C (45.9%, range 39.2 to 51.4%), the eldest group. By the end of the study at 8-weeks, Group A continued to retain higher fat graft volumes (73.3%, range 68.7 to 76.1%), again followed by Group B (45.3%, range 42.5 to 51.1%), and finally Group C (35.4%, range 30.7 to 39.9%), which demonstrated the poorest volume retention.

Figure 2.

Figure 2.

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Fat graft volume retention over time. (top) Representative three-dimensional reconstructed images using microcomputed tomography of fat grafts from each age group at 8 weeks post-grafting. (bottom) Median and range for fat graft volume retention (n=6 animals per group) over 8 weeks, expressed as a percentage of the baseline volume per graft, measured on post-operative day 1. (**p<0.01.)

Fat Graft Histological Analysis

Histological examination of explanted grafts demonstrated greatest overall health in the youngest group (Figure 3) (See Figure, Supplemental Digital Content 1, which shows Histological assessment of graft architecture and health at weeks 2, 4, and 6 (n=3 grafts per group per time point, five sections analyzed per graft, five blinded scorers). (A) Representative H&E stained sections of fat grafts from different age groups at weeks 2, 4, and 6 (10× magnification, scale bar = 200 μm). (B) Histology scoring for architectural characteristics of fat grafts from each group at weeks 2, 4, and 6, with fat graft median and range for health/integrity on the vertical axis, and median and range for injury score comprised of cyst/vacuole formation, inflammation, and fibrosis on the horizontal axis (**p < 0.01), INSERT HYPER LINK). Integrity scores in Group A were significantly greater than Groups B and C at all time points (**p < 0.01). Scores for tissue integrity were lowest in the oldest group, which also had the highest injury scores for inflammation, cysts or vacuoles, and fibrosis at each time point. Within a single age group, it was found that health/integrity of the tissue architecture generally decreased between weeks 2 and 8 and, as expected, injury scores increased due to formation of cyst and vacuoles, as well as fibrosis. These findings were observed in each group.

Figure 3.

Figure 3.

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Histological assessment of graft architecture and health (n=3 grafts per group per time point, five sections analyzed per graft, five blinded scorers). (top, left) Representative H&E stained sections of fat grafts from different age groups at week 8 (10× magnification, scale bar = 200 μm). (top, right) Histology scoring for architectural characteristics of fat grafts from each group at week 8, with median and range for fat graft health/integrity on the vertical axis, and median and range for injury score comprised of cyst/vacuole formation, inflammation, and fibrosis on the horizontal axis (**p < 0.01). (bottom, left) Representative CD31 stained sections of fat grafts (red) with DAPI (blue) counterstain from different age groups (A, B, and C) at week 8 (20× magnification, scale bar = 100 μm). (bottom, right) Quantification of staining with percent CD31 fluorescence per high-power field for each age group (**p < 0.01).

On immunohistochemical assessment for CD31, it was found that fat grafts in younger recipients (Group A) had significantly greater vascularity at each time point than those in Group B or Group C, measured by anti-CD31 antibody fluorescence per high-power field (**p < 0.01) (See Figure, Supplemental Digital Content 2, which shows Immunohistochemical staining and quantification of CD31 at weeks 2, 4, and 6 (n=3 grafts per group per time point. (A) Representative CD31 stained sections of fat grafts from different age groups at weeks 2, 4, and 6 (20× magnification, scale bar = 100 μm). (B) Quantification of staining with percent CD31 (red) fluorescence per high-power field (median and range) for each age group (**p < 0.01), INSERT HYPER LINK), indicating higher levels of vascularization in the youngest groups that was consistent over 8 weeks. As expected, fat grafts in Group C had the lowest amount of vascularity at each time point over 8 weeks.

Recipient Site Histological Analysis

To evaluate for differences in the recipient site soft tissue, skin samples were obtained from animals in each group and Isolectin was used as a vascular stain. Three-dimensional confocal microscopy with isosurface rendering of vasculature revealed increased staining at the recipient site samples adjacent to where fat grafts would be placed from younger animals (Group A) compared to animals from either older group (Groups B and C) (Figure 4). Quantification of three-dimensional vascular density demonstrated significantly greater amount of vascularity in Group A relative to Groups B and C (**p < 0.01).

Figure 4.

Figure 4.

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Confocal imaging of recipient site vascularity. (above) Isosurface rendering of vasculature from confocal images obtained by whole-mounted skin harvested from animals in Groups A, B, and C and stained with Isolectin (red), Hoechst 33342 (blue), and CD31 (green), scale bar = 200 μm. (Below) Quantification of vascular density demonstrated significantly greater vascularity in Group A recipient site skin compared to Groups B and C (**p < 0.01).

Discussion

Bone and soft-tissue deficiencies may contribute to facial asymmetry seen in a wide range of congenital and acquired craniofacial disorders. Multiple, staged procedures are frequently needed, and can be performed beginning as early as the neonatal period and extend into adult-hood. But despite severe asymmetry, many surgeons have advocated delaying soft tissue correction until after the facial skeletal deficiency has been addressed (22–25). By first establishing a stable skeletal foundation, soft tissue correction may then be considered as a “finishing touch.” This approach was further supported by the notion that in many instances, early soft tissue correction was not thought to alter the underlying pathophysiology and natural course of the disease (26–29).

With the popularity of autologous fat grafting increasing for correction of facial asymmetry over the last decade, however, other authors have begun to advocate for earlier intervention, especially in children with significant asymmetry who, without correction, may be subject to psychosocial issues during development. In comparing outcomes for serial fat grafting to correction of soft-tissue deficiency with microvascular free tissue transfer for craniofacial microsomia, Tanna and colleagues reported that fat grafting can be safely performed in combination with other operative interventions throughout childhood, and can result in good symmetry outcomes when done before the skeletal deficiency is addressed (15). With Parry-Romberg, serial fat grafting has also been described to be effective when begun at an early age (30). Furthermore, fat grafting performed in younger children was found to be associated with greater patient/family satisfaction scores compared to patients who had fat grafting delayed until they were older (29). Finally, in patients with Treacher Collins syndrome, Konofaos et al. reported that early fat grafting at greater than six months of age can effectively improve contour and tissue quality in the orbitozygomatic area (31). Moreover, they suggested that fat grafting prior to bone reconstruction may possibly minimize subsequent bone resorption through improved pliability and vascularization of the soft tissue envelop (32).

In considering timing for fat grafting, some studies have shown variability in fat graft quality with increasing age, particularly with regard to the stromal cell population. In older patients, unfavorable growth kinetics and differentiation/functional capacity of adipose-derived stromal cells (ASCs) have been reported (33, 34), with reduced induction of vasculogenesis in wound healing, therapeutic benefits for pulmonary fibrosis, and rescue of mouse hind-limb ischemia described for old compared to young ASCs (35–37). Age-related changes to ASCs may be particularly relevant to fat graft retention, as studies have shown stromal cells within fat to promote early re-vascularization and potentially contribute to mature adipocyte volume through direct differentiation (38–41). These findings thus argue for earlier fat grafting to enhance volume retention.

Aside from these intrinsic considerations, quality of the macroenvironmental niche into which the fat graft is placed may be of similar importance for long-term outcomes. Fat graft retention has been shown to be significantly reduced when placed into fibrotic, hypovascular recipient beds (20, 42). Relative to age, progressive changes to the vascularity and mechanical properties of soft tissue have also been well described, and these may likewise impact fat graft retention (43). Finally, it has been shown that young, but not old, extracellular matrix may preserve proliferative and self-renewal capacity of adult mesenchymal cells, possibly through increased expression of telomerase in neighboring cells or diminished expression of reactive oxygen species (44–46). These reports thus suggest that a younger macroenvironmental niche into which fat grafts are placed may be more conducive to retention of transferred fat.

In this present study, we evaluated the effects of recipient bed age on retention of human fat grafts. We observed that fat grafting into the young recipient bed improved volume retention when compared to older recipient beds. This was associated with enhanced CD31 staining and improved fat graft histology, with greater histologic quality noted in fat grafts placed into younger animals. Finally, evaluation of the recipient site with Isolectin staining and visualization by three-dimensional confocal microscopy demonstrated significantly increased vascularity in younger animals, which may promote greater early re-vascularization of ischemic fat grafts following placement. While there are undoubtedly other recipient site factors that may impact fat graft retention, these data nonetheless support beginning soft tissue correction at earlier ages, particularly in reconstructions that require multiple surgeries, such as for craniofacial microsomia, as staged procedures afford the opportunity to also perform concomitant fat grafting at younger ages.

Importantly, the findings from this investigation may have some translational limitations. Autologous fat transfer implies grafting into the same patient from which the donor fat was harvested. Therefore, it may be more appropriate to match a young recipient site with young donor fat, and investigate fat grafting outcomes within this context. However, exploring this would require sufficient samples from pediatric patients, the accessibility of which is more challenging than that of human adult fat, particularly with regard to available volume. As a result, only adult donor fat was used in this study. Nonetheless, use of only adult fat allowed for determination of recipient bed age-related effects on graft retention independent of fat graft age. Fat grafting was also done in immunocompromised mice to minimize recipient response to xenogeneic tissue. While this model has been extensively used in the literature to study human fat (18, 20, 41, 42), in the clinical setting, a normal immune/inflammatory response to autologous fat may also impact ultimate volume retention. Alternatively, other studies have frequently employed mouse inguinal fat pads transplanted into immunocompetent recipients to more accurately reflect the immune response encountered in patients (38, 40). However, dramatic species-specific differences in adipose tissue between mouse inguinal fat and human abdominal fat would potentially limit the translatability of this approach (47–50).

Our findings may also not fully extend to all causes of soft tissue deformities, as the pathophysiology of some conditions (e.g. Parry Romberg syndrome, post-radiation atrophy) may impact the recipient site quality. Slack et al. reported decreased fat graft retention in patients with Parry Romberg compared to non-syndromic/ cosmetic patients receiving fat grafts at similar locations (30). However, whether this was due to reduced blood supply in the recipient bed or something else intrinsic to the disease process remains unknown. Experiments in our study used healthy mouse recipient tissue, and it can be argued that the outcomes observed may be contingent on the presence of normal tissue in the recipient bed. Therefore, some disease states may respond differently to fat grafting, and the role recipient age may play in these settings remains less clear. Finally, recipient ages for mice in this study may not directly translate to humans, as species specific differences undoubtedly exist in the aging process. While our data did demonstrate improved fat graft retention in younger animals, recapitulation of these findings in humans will require additional clinical studies.

Conclusion

While fat grafting has become an increasing popular modality to manage soft tissue deficits, no consensus has been reached regarding optimal timing for these procedures. Our in vivo results have demonstrated that placement of fat grafts into a younger recipient bed, versus an older recipient bed, is advantageous for maintenance of graft volume. Additional histologic evidence showed improved vascularization and overall health of grafts that were placed in a younger macroenvironmental niche. A more longitudinal approach to soft tissue correction starting at a younger age may thus provide benefit to the patient with more predictable results.

Supplementary Material

SDC 1

Figure, Supplemental Digital Content 1. Histological assessment of graft architecture and health at weeks 2, 4, and 6 (n=3 grafts per group per time point, five sections analyzed per graft, five blinded scorers). (A) Representative H&E stained sections of fat grafts from different age groups at weeks 2, 4, and 6 (10× magnification, scale bar = 200 μm). (B) Histology scoring for architectural characteristics of fat grafts from each group at weeks 2, 4, and 6, with fat graft median and range for health/integrity on the vertical axis, and median and range for injury score comprised of cyst/vacuole formation, inflammation, and fibrosis on the horizontal axis (**p < 0.01), INSERT HYPER LINK.

SDC 2

Figure, Supplemental Digital Content 2. Immunohistochemical staining and quantification of CD31 at weeks 2, 4, and 6 (n=3 grafts per group per time point. (A) Representative CD31 stained sections of fat grafts from different age groups at weeks 2, 4, and 6 (20× magnification, scale bar = 100 μm). (B) Quantification of staining with percent CD31 (red) fluorescence per high-power field (median and range) for each age group (**p < 0.01), INSERT HYPER LINK.

Acknowledgements

We thank the Stanford University Plastic and Reconstructive Surgery Department for providing the biological specimens required for this investigation, and the Stanford Small Animal Imaging Facility for use of their imaging instruments and technology. We also thank Ethan Shen for assistance with quantification of confocal images. M.T.L. was supported by NIH grants U01 HL099776, R01 DE021683-03, R21 DE024230-02, the Oak Foundation, the Hagey Laboratory for Pediatric Regenerative Medicine, and the Gunn/Olivier Fund. D.C.W. was supported by NIH grant K08 DE024269, the Hagey Laboratory for Pediatric Regenerative Medicine, and the Stanford University Child Health Research Institute Faculty Scholar Award.

Footnotes

Financial Disclosure: None of the authors has a financial interest in any of the products, devices, or drugs mentioned in this manuscript.

References

  • 1.Kakagia D, Pallua N Autologous fat grafting: in search of the optimal technique. Surgical innovation 2014;21:327–336. [DOI] [PubMed] [Google Scholar]
  • 2.Al Sufyani MA, Al Hargan AH, Al Shammari NA, Al Sufyani MA Autologous Fat Transfer for Breast Augmentation: A Review. Dermatologic surgery : official publication for American Society for Dermatologic Surgery [et al. ] 2016;42:1235–1242. [DOI] [PubMed] [Google Scholar]
  • 3.Chiu CH Correction with autologous fat grafting for contour changes of the breasts after implant removal in Asian women. Journal of plastic, reconstructive & aesthetic surgery : JPRAS 2016;69:61–69. [DOI] [PubMed] [Google Scholar]
  • 4.Weichman KE, Broer PN, Tanna N, et al. The role of autologous fat grafting in secondary microsurgical breast reconstruction. Annals of plastic surgery 2013;71:24–30. [DOI] [PubMed] [Google Scholar]
  • 5.Largo RD, Tchang LA, Mele V, et al. Efficacy, safety and complications of autologous fat grafting to healthy breast tissue: a systematic review. Journal of plastic, reconstructive & aesthetic surgery : JPRAS 2014;67:437–448. [DOI] [PubMed] [Google Scholar]
  • 6.Wetterau M, Szpalski C, Hazen A, Warren SM Autologous fat grafting and facial reconstruction. The Journal of craniofacial surgery 2012;23:315–318. [DOI] [PubMed] [Google Scholar]
  • 7.Ross RJ, Shayan R, Mutimer KL, Ashton MW Autologous fat grafting: current state of the art and critical review. Annals of plastic surgery 2014;73:352–357. [DOI] [PubMed] [Google Scholar]
  • 8.Herold C, Ueberreiter K, Busche MN, Vogt PM Autologous fat transplantation: volumetric tools for estimation of volume survival. A systematic review. Aesthetic plastic surgery 2013;37:380–387. [DOI] [PubMed] [Google Scholar]
  • 9.Khouri RK, Khouri RK Jr., Rigotti G, et al. Aesthetic applications of Brava-assisted megavolume fat grafting to the breasts: a 9-year, 476-patient, multicenter experience. Plastic and reconstructive surgery 2014;133:796–807; discussion 808-799. [DOI] [PubMed] [Google Scholar]
  • 10.Khouri RK, Rigotti G, Khouri RK Jr., et al. Tissue-engineered breast reconstruction with Brava-assisted fat grafting: a 7-year, 488-patient, multicenter experience. Plastic and reconstructive surgery 2015;135:643–658. [DOI] [PubMed] [Google Scholar]
  • 11.Ring A, Kirchhoff P, Goertz O, et al. Reconstruction of Soft-Tissue Defects at the Foot and Ankle after Oncological Resection. Frontiers in surgery 2016;3:15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Gold MH Use of hyaluronic acid fillers for the treatment of the aging face. Clinical Interventions in Aging 2007;2:369–376. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Gamboa GM, Ross WA Autologous fat transfer in aesthetic facial recontouring. Annals of plastic surgery 2013;70:513–516. [DOI] [PubMed] [Google Scholar]
  • 14.Chang Q, Li J, Dong Z, Liu L, Lu F Quantitative volumetric analysis of progressive hemifacial atrophy corrected using stromal vascular fraction-supplemented autologous fat grafts. Dermatologic surgery : official publication for American Society for Dermatologic Surgery [et al. ] 2013;39:1465–1473. [DOI] [PubMed] [Google Scholar]
  • 15.Tanna N, Wan DC, Kawamoto HK, Bradley JP Craniofacial microsomia soft-tissue reconstruction comparison: inframammary extended circumflex scapular flap versus serial fat grafting. Plastic and reconstructive surgery 2011;127:802–811. [DOI] [PubMed] [Google Scholar]
  • 16.Baldelli I, Santi P, Dova L, et al. Body Image Disorders and Surgical Timing in Patients Affected by Poland Syndrome: Data Analysis of 58 Case Studies. Plastic and reconstructive surgery 2016;137:1273–1282. [DOI] [PubMed] [Google Scholar]
  • 17.Tringale KR, Lance S, Schoenbrunner A, Gosman AA Sustained Overcorrection After Autologous Facial Fat Grafting in the Pediatric Population: A Case Series. Annals of plastic surgery 2017. [DOI] [PubMed] [Google Scholar]
  • 18.Chung MT, Hyun JS, Lo DD, et al. Micro-computed tomography evaluation of human fat grafts in nude mice. Tissue engineering Part C, Methods 2013;19:227–232. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Lee JH, Kirkham JC, McCormack MC, Nicholls AM, Randolph MA, Austen WG Jr. The effect of pressure and shear on autologous fat grafting. Plastic and reconstructive surgery 2013;131:1125–1136. [DOI] [PubMed] [Google Scholar]
  • 20.Garza RM, Paik KJ, Chung MT, et al. Studies in fat grafting: Part III. Fat grafting irradiated tissue--improved skin quality and decreased fat graft retention. Plastic and reconstructive surgery 2014;134:249–257. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Berry R, Church CD, Gericke MT, Jeffery E, Colman L, Rodeheffer MS Imaging of adipose tissue. Methods Enzymol 2014;537:47–73. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Siebert JW, Anson G, Longaker MT Microsurgical correction of facial asymmetry in 60 consecutive cases. Plastic and reconstructive surgery 1996;97:354–363. [DOI] [PubMed] [Google Scholar]
  • 23.Siebert JW, Longaker MT, Angrigiani C The inframammary extended circumflex scapular flap: an aesthetic improvement of the parascapular flap. Plastic and reconstructive surgery 1997;99:70–77. [DOI] [PubMed] [Google Scholar]
  • 24.Longaker MT, Siebert JW Microsurgical correction of facial contour in congenital craniofacial malformations: the marriage of hard and soft tissue. Plastic and reconstructive surgery 1996;98:942–950. [DOI] [PubMed] [Google Scholar]
  • 25.Longaker MT, Siebert JW Microvascular free-flap correction of severe hemifacial atrophy. Plastic and reconstructive surgery 1995;96:800–809. [DOI] [PubMed] [Google Scholar]
  • 26.Saadeh P, Reavey PL, Siebert JW A soft-tissue approach to midfacial hypoplasia associated with Treacher Collins syndrome. Annals of plastic surgery 2006;56:522–525. [DOI] [PubMed] [Google Scholar]
  • 27.Grimaldi M, Gentile P, Labardi L, Silvi E, Trimarco A, Cervelli V Lipostructure technique in Romberg syndrome. The Journal of craniofacial surgery 2008;19:1089–1091. [DOI] [PubMed] [Google Scholar]
  • 28.Cervelli V, Gentile P Use of cell fat mixed with platelet gel in progressive hemifacial atrophy. Aesthetic Plast Surg 2009;33:22–27. [DOI] [PubMed] [Google Scholar]
  • 29.Slack GC, Tabit CJ, Allam KA, Kawamoto HK, Bradley JP Parry-Romberg reconstruction: optimal timing for hard and soft tissue procedures. The Journal of craniofacial surgery 2012;23:1969–1973. [DOI] [PubMed] [Google Scholar]
  • 30.Slack GC, Tabit CJ, Allam KA, Kawamoto HK, Bradley JP Parry-Romberg reconstruction: beneficial results despite poorer fat take. Annals of plastic surgery 2014;73:307–310. [DOI] [PubMed] [Google Scholar]
  • 31.Konofaos P, Arnaud E Early Fat Grafting for Augmentation of Orbitozygomatic Region in Treacher Collins Syndrome. The Journal of craniofacial surgery 2015;26:1258–1260. [DOI] [PubMed] [Google Scholar]
  • 32.Garland CB, Pomerantz JH Regenerative strategies for craniofacial disorders. Front Physiol 2012;3:453. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Zhu M, Kohan E, Bradley J, Hedrick M, Benhaim P, Zuk P The effect of age on osteogenic, adipogenic and proliferative potential of female adipose-derived stem cells. Journal of tissue engineering and regenerative medicine 2009;3:290–301. [DOI] [PubMed] [Google Scholar]
  • 34.El-Ftesi S, Chang EI, Longaker MT, Gurtner GC Aging and diabetes impair the neovascular potential of adipose-derived stromal cells. Plastic and reconstructive surgery 2009;123:475–485. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Duscher D, Rennert RC, Januszyk M, et al. Aging disrupts cell subpopulation dynamics and diminishes the function of mesenchymal stem cells. Scientific reports 2014;4:7144. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Tashiro J, Elliot SJ, Gerth DJ, et al. Therapeutic benefits of young, but not old, adipose-derived mesenchymal stem cells in a chronic mouse model of bleomycin-induced pulmonary fibrosis. Translational research : the journal of laboratory and clinical medicine 2015;166:554–567. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Abbo O, Taurand M, Monsarrat P, et al. Comparison between pediatric and adult adipose mesenchymal stromal cells. Cytotherapy 2017;19:395–407. [DOI] [PubMed] [Google Scholar]
  • 38.Eto H, Kato H, Suga H, et al. The fate of adipocytes after nonvascularized fat grafting: evidence of early death and replacement of adipocytes. Plastic and reconstructive surgery 2012;129:1081–1092. [DOI] [PubMed] [Google Scholar]
  • 39.Suga H, Eto H, Aoi N, et al. Adipose tissue remodeling under ischemia: death of adipocytes and activation of stem/progenitor cells. Plastic and reconstructive surgery 2010;126:1911–1923. [DOI] [PubMed] [Google Scholar]
  • 40.Doi K, Ogata F, Eto H, et al. Differential contributions of graft-derived and host-derived cells in tissue regeneration/remodeling after fat grafting. Plastic and reconstructive surgery 2015;135:1607–1617. [DOI] [PubMed] [Google Scholar]
  • 41.Garza RM, Rennert RC, Paik KJ, et al. Studies in fat grafting: Part IV. Adipose-derived stromal cell gene expression in cell-assisted lipotransfer. Plastic and reconstructive surgery 2015;135:1045–1055. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42.Luan A, Duscher D, Whittam AJ, et al. Cell-Assisted Lipotransfer Improves Volume Retention in Irradiated Recipient Sites and Rescues Radiation-Induced Skin Changes. Stem cells 2016;34:668–673. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43.Farage MA, M. KW; Maibach HI ed. Textbook of Aging Skin, 2010th Edition ed. New York, NY: Springer; 2010. [Google Scholar]
  • 44.Lynch K, Pei M Age associated communication between cells and matrix: a potential impact on stem cell-based tissue regeneration strategies. Organogenesis 2014;10:289–298. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45.Sun Y, Li W, Lu Z, et al. Rescuing replication and osteogenesis of aged mesenchymal stem cells by exposure to a young extracellular matrix. FASEB journal : official publication of the Federation of American Societies for Experimental Biology 2011;25:1474–1485. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Choi HR, Cho KA, Kang HT, et al. Restoration of senescent human diploid fibroblasts by modulation of the extracellular matrix. Aging cell 2011;10:148–157. [DOI] [PubMed] [Google Scholar]
  • 47.Gesta S, Bluher M, Yamamoto Y, et al. Evidence for a role of developmental genes in the origin of obesity and body fat distribution. Proceedings of the National Academy of Sciences of the United States of America 2006;103:6676–6681. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48.Rosell M, Kaforou M, Frontini A, et al. Brown and white adipose tissues: intrinsic differences in gene expression and response to cold exposure in mice. American journal of physiology Endocrinology and metabolism 2014;306:E945–964. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Linder K, Arner P, Flores-Morales A, Tollet-Egnell P, Norstedt G Differentially expressed genes in visceral or subcutaneous adipose tissue of obese men and women. J Lipid Res 2004;45:148–154. [DOI] [PubMed] [Google Scholar]
  • 50.Chusyd DE, Wang D, Huffman DM, Nagy TR Relationships between Rodent White Adipose Fat Pads and Human White Adipose Fat Depots. Front Nutr 2016;3:10. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

SDC 1

Figure, Supplemental Digital Content 1. Histological assessment of graft architecture and health at weeks 2, 4, and 6 (n=3 grafts per group per time point, five sections analyzed per graft, five blinded scorers). (A) Representative H&E stained sections of fat grafts from different age groups at weeks 2, 4, and 6 (10× magnification, scale bar = 200 μm). (B) Histology scoring for architectural characteristics of fat grafts from each group at weeks 2, 4, and 6, with fat graft median and range for health/integrity on the vertical axis, and median and range for injury score comprised of cyst/vacuole formation, inflammation, and fibrosis on the horizontal axis (**p < 0.01), INSERT HYPER LINK.

NIHMS1515353-supplement-SDC_1.pdf (5.1MB, pdf)
SDC 2

Figure, Supplemental Digital Content 2. Immunohistochemical staining and quantification of CD31 at weeks 2, 4, and 6 (n=3 grafts per group per time point. (A) Representative CD31 stained sections of fat grafts from different age groups at weeks 2, 4, and 6 (20× magnification, scale bar = 100 μm). (B) Quantification of staining with percent CD31 (red) fluorescence per high-power field (median and range) for each age group (**p < 0.01), INSERT HYPER LINK.

NIHMS1515353-supplement-SDC_2.pdf (1.9MB, pdf)

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