Cosmetic: Special Topic

Technical Precision with Autologous Fat Grafting for Facial Rejuvenation: A Review of the Evolving Science

Strong, Amy L. MD, PhD1; Rohrich, Rod J. MD2,3; Tonnard, Patrick L. MD4; Vargo, James D. MD1; Cederna, Paul S. MD1

Author Information

Ann Arbor, MI; Dallas and Houston, TX; and Gent, Belgium

From the 1Section of Plastic and Reconstructive Surgery, Department of Surgery, University of Michigan

2Dallas Plastic Surgery Institute

3Baylor College of Medicine

4Coupure Centrum voor Plastische Chrirugie.

Received for publication February 7, 2021; accepted November 29, 2022.

Disclosure statements are at the end of this article, following the correspondence information.

Related digital media are available in the full-text version of the article on www.PRSJournal.com.

Paul S. Cederna, MD, Section of Plastic and Reconstructive Surgery, University of Michigan, 2130 Taubman Center, 1500 East Medical Center Drive, Ann Arbor, MI 48109-0340, [email protected]

Plastic and Reconstructive Surgery 153(2):p 360-377, February 2024. | DOI: 10.1097/PRS.0000000000010643

Abstract

Summary: 

The scientific study of facial aging has transformed modern facial rejuvenation. As people age, fat loss in specific fat compartments is a major contributor to structural aging of the face. Autologous fat grafting is safe, abundant, readily available, and completely biocompatible, which makes it the preferred soft-tissue filler in the correction of facial atrophy. The addition of volume through fat grafting gives an aging face a more youthful, healthy, and aesthetic appearance. Harvesting and preparation with different cannula sizes and filter-cartridge techniques have allowed for fat grafts to be divided based on parcel size and cell type into three major subtypes: macrofat, microfat, and nanofat. Macrofat and microfat have the benefit of providing volume to restore areas of facial deflation and atrophy in addition to improving skin quality; nanofat has been shown to improve skin texture and pigmentation. In this article, the authors discuss the current opinions regarding fat grafting and how the evolving science of fat grafting has led to the clinical utility of each type of fat to optimize facial rejuvenation. The opportunity exists to individualize the use of autologous fat grafting with the various subtypes of fat for the targeted correction of aging in different anatomic areas of the face. Fat grafting has become a powerful tool that has revolutionized facial rejuvenation, and developing precise, individualized plans for autologous fat grafting for each patient is an important advancement in the evolution of facial rejuvenation.

Autologous fat grafting has become increasingly popular in facial rejuvenation, with 40,000 to 55,000 procedures being performed each year in the United States.1 Fat is safe to use, readily available, and biocompatible, which makes it a highly valuable soft-tissue filler. Harvesting fat for facial fat grafting is performed using either a dry technique or tumescent solution, followed by aspiration of the fat through handheld syringe or liposuction devices, and processed using different techniques to concentrate the graft material, sequester high-density adipocytes and other cellular components, and remove cellular debris. Each step introduces variables that affect the character of the fat obtained, viability of the cells, and overall fat graft survival. Studies have been performed to help delineate the optimal anatomic location for fat graft harvesting, cannula and port size, multiperforated or single-perforated cannulas, tumescent technique (dry or wet technique), harvest technique (handheld or powered device), and fat-processing technique (Table 1).2 However, even with the extensive body of literature in this area, the results related to autologous fat grafting remain highly variable. Therefore, additional comprehensive basic science and clinical studies are necessary to determine the optimal approach to maximize fat graft retention.

Table 1. - Current Studies on Harvesting and Processing of Macrofat
Topic Study Primary Objective Detailed Methods Results
Donor site Rohrich et al. (2004) Assessed abdomen, thigh, flank, and knee Dry technique; handheld syringe aspiration; centrifugation at 500 g for 2 minutes No differences in cell viability between harvest sites
Ullmann et al. (2005) Assessed abdomen, lateral thigh, and breast Tumescent solution; handheld aspiration; centrifugation at 1500 rpm for 5 minutes No difference in donor site in volume retention when injected into nude animals with minor differences in the structural integrity and vascularity of the grafts between the donor sites
Padoin et al. (2008) Evaluated upper abdomen, lower abdomen, thigh, flank, and knee Tumescent solution; SAL; centrifugation at 450 g for 5 minutes; SVF cells isolated with collagenase for 1 hour at 37°C Greatest number of viable cells was harvested from lower abdomen and inner thigh
Lim et al. (2012) Assessed abdomen and nonabdominal regions Tumescent solution; handheld syringe aspiration; centrifugation at 1200 rpm for 3 minutes No difference in donor site with respect to volume retention and ability to correct craniofacial defects
Li et al. (2013) Assessed upper and lower abdomen, lateral and inner thigh, and flank Tumescent solution, handheld syringe aspiration; centrifugation at 1000 rpm for 3 minutes; SVF cells isolated with collagenase for 30 minutes at 37°C No difference in SVF cells concentration between harvest sites; no difference in volume retention of the grafts when injected into nude mice
Small et al. (2014) Assessed abdomen and lateral thigh Tumescent solution; SAL; centrifugation at 3000 rpm for 3 minutes No difference in volume retention between donor sites
Geissler et al. (2014) Assessed lower abdomen, inner thigh, and flank Dry technique; handheld syringe aspiration; centrifugation at 1200 rpm for 3 minutes No different in adipocyte viability between donor sites
Tsekouras et al. (2017) Assessed abdomen, inner and outer thigh, waist, and inner knee Dry technique; SAL; centrifugation at 400 g for 15 minutes; SVF cells isolated with collagenase for 25 minutes at 37°C Outer thigh exhibited higher SVF cells compared with other donor sites
Cannula characteristics Schiffman et al. (2001) Assessed effects of 2.5-mm and 3-mm cannula size Dry technique, handheld aspiration, centrifugation at 250 g and 300 g for 5 minutes No difference in adipocyte viability in the different cannula sizes
Ozsoy et al. (2006) Compared 2-mm, 3-mm, and 4-mm aspiration cannula Dry technique, handheld aspiration; centrifugation at 1000 rpm for 5 minutes Highest adipocyte viability with largest diameter (40 mm)
Erdim et al. (2009) Compared 2-mm, 4-mm, and 6-mm aspiration cannula Dry technique; handheld syringe aspiration; gravity separation; adipocytes isolated with collagenase Greatest cell adipocyte viability in the 60-mm aspiration cannula
Kirkham et al. (2012) Compared 3-mm and 5-mm aspiration cannulas Tumescent solution; SAL; centrifugation at 200 g for 5 minutes Increased fat retention (25%) in the 5-mm group compared with the 3-mm group; more intact adipocytes in the 5-mm group compared with the 3-mm group
Alharbi et al. (2013) Compared the 3-mm aspiration cannula with the 2-mm multiperforated cannula Tumescent solution; handheld syringe aspiration; centrifugation at 300 rpm for 3 minutes; SVF cell isolation with collagenase for 45 minutes at 37°C No difference in SVF concentration or cell count with different aspiration cannula; increased in vitro SVF/ADSC viability after 24 and 48 hours with the 2-mm multiperforated cannula
Trivisonno et al. (2014) Assessed multiperforated 2-mm and single-port 3-mm-diameter cannulas Tumescent solution; handheld syringe aspiration; centrifugation at 300 g for 2 minutes; SVF cells/ADSCs isolated with collagenase for 30 minutes at 37°C No difference in cell viability between the two groups; slightly more SVF cells isolated from 20 multiperforated cannula
Tambasco et al. (2014) Compared 3-mm and 5-mm aspiration cannulas Tumescent solution; handheld syringe aspiration Less adipocyte rupture (25%) with the 5-mm cannula group compared with the 3-mm cannula group
Rubino et al. (2015) Compared 2-mm aspiration cannula with 3-mm Mercedes cannula Tumescent solution; handheld syringe aspiration; centrifugation at 3000 rpm for 3 minutes or sedimentation Higher adipocyte yield with the 3-mm Mercedes cannula, with less alteration in cell membrane and leakage of oily material
Aspiration technique Rohrich et al. (2000) Compared SAL, internal UAL, external UAL, and massage Tumescent solution; SAL, internal UAL, external UAL, or massage for 7 minutes; centrifugation at 500 g for 2 minutes SAL and external UAL resulted in minimal histologic disruption of adipose tissue; significant (70% to 90%) damage to adipocytes with internal UAL
Leong et al. (2005) Compared handheld syringe aspiration and SAL Tumescent solution; handheld syringe aspiration and SAL; centrifugation at 500 g for 5 minutes; SVF/ADSCs harvested with collagenase for 30 minutes at 37°C No difference in cell viability between handheld syringe aspiration and SAL
Smith et al. (2006) Compared handheld syringe aspiration and SAL Tumescent solution; handheld syringe aspiration or SAL; no treatment, centrifugation at 500 g for 2 minutes, manual washing with saline or lactated Ringer’s solution, manual washing with centrifugation at 500 g for 2 minutes No difference in cell viability between handheld syringe aspiration and SAL; no difference in graft retention in the groups when injected into nude mice
Oedayrajsingh-Varma et al. (2006) Compared SAL and UAL Tumescent solution; SAL, UAL, centrifugation at 1000 rpm for 5 minutes, collagenase for 1 hour at 37°C Reduced SVF cells in UAL compared with SAL
Pu et al. (2008) Compared handheld syringe aspiration and SAL Tumescent solution; handheld syringe aspiration or SAL; handheld syringe lipoaspiration processed with 3000 rpm for 3 minutes; SAL aspirates were processed by centrifugation at 500 rpm for 10 minutes Handheld syringe lipoaspirates had greater adipocyte viability; no difference in histologic analysis of handheld syringe aspiration and SAL
Crawford et al. (2010) Compared handheld syringe aspiration and SAL Tumescent solution; handheld syringe aspiration with the Viafill system or SAL; processed by centrifugation at 50 g for 2 minutes Handheld syringe aspiration resulted in greater adipocyte count
Chung et al. (2013) Compared SAL and LAL Tumescent solution; SAL, LAL; centrifugation at 1000 rpm for 5 minutes; stem cells harvested by collagenase for 1 hour at 37°C Cell yield, viability, proliferation, and frequency of stem cells less with LAL compared with SAL
Fisher et al. (2013) Compared SAL and UAL Tumescent solution; SAL, UAL; filtration for 30 minutes No difference in SVF cell count and graft retention when injected into nude mice
Keck et al. (2014) Compared handheld syringe aspiration and SAL Dry technique; handheld syringe aspiration or SAL; processed by centrifugation at 380 g for 5 minutes No difference in cell counts or cell viability between handheld syringe aspiration and SAL; handheld syringe aspiration resulted in slightly greater oil release
Charles-de-Sa et al. (2015) Compared handheld syringe aspiration and SAL Tumescent solution; handheld syringe aspiration or SAL; centrifugation at 3000 rpm for 3 minutes; SVF cells isolated with collagenase for 30 minutes at 37°C No difference in adipocyte count or ADSC viability
Duscher et al. (2016) Compared SAL and UAL Tumescent solution; UAL, SAL, processed by centrifugation at 1000 rpm for 5 minutes; stem cells harvested by collagenase for 1 hour at 37°C No difference in cell yield, viability, proliferation, surface marker phenotype, biology of stem cells
Yildiz et al. (2016) Compared SAL and LAL Tumescent solution; SAL, LAL, centrifugation at 3000 rpm for 3 minutes; SVF/ADSCs harvested by collagenase for 1 hour at 37°C Lower number of viable ADSCs and higher apoptosis in vitro after LAL compared with SAL
Processing technique Rohrich et al. (2004) Compared with and without centrifugation Dry technique; handheld syringe aspiration; processed by centrifugation at 500 g for 2 minutes Centrifugation reduced cell proliferation
Ramon et al. (2005) Compared centrifugation and towel filtration Tumescent solution; handheld syringe aspiration; centrifugation at 1500 rpm for 5 minutes, or towel filtration for 10 minutes No difference in graft size when injected into nude mice; histological analysis revealed less fibrosis of the grafts with towel filtration
Rose et al. (2006) Compared gravity separation, centrifugation, and manual washing with centrifugation Tumescent solution; handheld syringe aspiration; processed by gravity separation, centrifugation at 3000 rpm (6000 g) for 3 minutes, or manual washing with centrifugation at 3000 rpm for 3 minutes Centrifugation and washing with centrifugation reduced cell number, nucleated adipocytes, and cross-sectional area of adipocytes; gravity separation had higher adipocyte count
Smith et al. (2006) Compared centrifugation, manual washing, and manual washing with centrifugation Tumescent solution; handheld syringe aspiration or SAL; no treatment, centrifugation at 500 g for 2 minutes, manual washing with saline or lactated Ringer’s solution, manual washing with centrifugation at 500 g for 2 minutes No difference in adipocyte cell viability between the groups; no difference in graft retention when injected into nude mice
Kurita et al. (2008) Compared with and without centrifugation at different speeds Tumescent solution; SAL; no centrifugation, centrifugation at 0 g, 400 g, 800 g, 1200 g, 3000 g, 4200 g for 3 minutes; SVF cells isolated with collagenase for 30 minutes at 37°C No difference in SVF cell count with or without centrifugation at the different speeds, except greater than 3000 g centrifugation resulted in larger grafts compared with no centrifugation
Khater et al. (2009) Compared centrifugation and manual washing Dry technique; handheld syringe aspiration; centrifugation at 3400 rpm for 3 minutes or manual washing Greater patient satisfaction with washing compared with centrifugation; more preadiopcytes with washing compared with centrifugation
Conde-Green et al. (2010) Compared gravity separation, centrifugation, and manual washing Tumescent solution; SAL; processed by gravity separation or centrifugation at 3000 rpm for 3 minutes Centrifugation and manual washing reduced adipocyte count but was better at removing cellular debris
Minn et al. (2010) Compared centrifugation, cotton gauze rolling, and metal sieve filtration Tumescent solution; handheld syringe aspiration; centrifugation at 1800 g for 3 minutes, metal sieve filtration for 3 minutes, cotton gauze rolling for 3 minutes No difference between centrifugation and cotton gauze rolling; metal sieve resulted in reduced cell viability; no difference in graft retention rates among the three groups when injected into nude mice
Botti et al. (2011) Compared centrifugation and filtration Tumescent solution; handheld syringe aspiration; centrifugation at 3000 rpm for 3 minutes, washing No difference in volume retention in patients with centrifugation or filtration; increased nodule formation in filtration group
Fisher et al. (2013) Compared centrifugation, cotton gauze rolling, and filtration Tumescent solution; handheld syringe aspiration and centrifugation at 3000 rpm for 3 minutes, handheld syringe aspiration and cotton gauze rolling for 5 minutes, or SAL and filtration Cotton gauze rolling resulted in the greatest SVF yield and in vivo graft retention when injected into nude mice
Salinas et al. (2014) Compared centrifugation at different speeds and mesh gauze rolling Tumescent solution; handheld syringe aspiration; centrifugation at 50 g, 1200 g, 5000 g, 10,000 g, or 23,000 g for 3 minutes and mesh gauze rolling; ADSC isolated with collagenase for 30 minutes at 37°C No difference in percentage of concentrated fat or ADSC count between centrifugation at 1200 g and mesh gauze rolling
Pfaff et al. (2014) Compared centrifugation and Telfa gauze processing Tumescent solution; handheld syringe aspiration; processed by centrifugation at 1500 rpm for 3 minutes or Telfa gauze rolling with large pieces of nonadherent dressing for 30 second; SVF cells harvested with collagenase for 1 hour at 37°C Cotton gauze rolling resulted in greater SVF/ADSC cell viability and count
Asilian et al. (2014) Compared centrifugation with sieve filtration Tumescent solution; handheld syringe aspiration; centrifugation at 3400 rpm for 1 minute or metal sieve filtration No difference between centrifugation and metal sieve filtration in patient outcomes
Rubino et al. (2015) Compared gravity separation and centrifugation Tumescent solution; handheld syringe aspiration; centrifugation at 3000 rpm for 3 minutes or gravity separation for 30 minutes No difference in adipocyte yield but higher number of morphologic altered cells in centrifugation compared with sedimentation
Cucchiani et al. (2016) Compared gravity separation, centrifugation, cotton gauze rolling, and manual washing Tumescent solution; SAL; gravity separation for 20 minutes, centrifugation at 700 rpm (94 g) for 3 minutes, centrifugation at 3000 rpm (1421 g) for 3 minutes, cotton gauze rolling for 3 minutes, manual washing Cell viability was greatest in the gravity separation > manual washing > cotton gauze rolling > centrifugation at 700 rpm > centrifugation at 3000 rpm
Canizares et al. (2017) Compared centrifugation and Telfa gauze rolling Tumescent solution; SAL; processed by centrifugation at 3000 rpm for 3 minutes or rolled on Telfa gauze for 2 to 4 minutes; SVF harvested with collagenase for 30 minutes at 37°C Centrifugation had higher number of ADSCs; Telfa rolling resulted in higher number of functional adipocytes
Streit et al. (2017) Compared gravity separation, centrifugation, and membrane filtration Tumescent solution; handheld syringe aspiration; gravity separation for 20 minutes, centrifugation at 1200 g for 3 minutes, or membrane filtration with PureGraft No significant difference in overall SVF/ADSC cell viability
Wu et al. (2018) Compared gravity separation, centrifugation, and cotton gauze rolling Tumescent solution; handheld syringe aspiration; gravity separation for 15 minutes, centrifugation at 1000 rpm (161 g) for 3 minutes, or cotton gauze rolling for 5 minutes Cotton gauze rolling demonstrated higher fat retention in patients assessed by three-dimensional imaging and analysis
Li et al. (2020) Compared gravity separation, centrifugation, and cotton gauze rolling Tumescent solution; handheld syringe aspiration; gravity separation for 30 minutes, centrifugation at 1000 rpm (209 g) for 5 minutes, centrifugation at 2400 rpm (1200 g) for 3 minutes, cotton gauze rolling for 5 minutes Cotton gauze rolling and gravity separation had higher functional adipocytes than centrifugation; cotton gauze rolling > centrifugation > gravity separation was most efficient in removing oil and aqueous fractions
ADSC, adipose-derived stem cell; LAL, laser-assisted liposuction; SAL, suction-assisted lipectomy; SVF, stromal vascular fraction; UAL, ultrasonic-assisted liposuction.

Early fat graft studies primarily focused on the structural component of fat, whereas more recent studies have examined the functional aspects of adipose tissue for the purposes of enhancing the regenerative capabilities of fat. In 2001, Zuk and colleagues3 discovered that adipose-derived stem cells (ADSCs) from lipoaspirate had regenerative properties after enzymatic dissociation. These ADSCs have been shown to secrete growth factors that improve skin texture.4 However, the use of these cells has been limited because of the enzymatic dissociation process, which may alter the biology of the cells, and therefore, it has not been approved for clinical use by the U.S. Food and Drug Administration.5–7 In 2013, Tonnard et al.8 published findings of mechanically dissociating autologous fat that allowed for the isolation of stromal vascular fraction (SVF) cells. In addition to the ADSCs in SVF, endothelial cells, pericytes, smooth muscle cells, and immune cells, which are capable of undergoing neoangiogenesis, have been found to support fat grafts.9 Independently, SVF cells have been shown to significantly improve skin elasticity and hyperpigmentation through regeneration of aged skin and to alter melanin content in the skin.9 Since these discoveries, the use of autologous fat grafting has increased dramatically, with more predictable results and with a broader scope of use than volumetric expansion.

Autologous fat grafting has been defined by a number of surgeons by the cell type and graft size in the final product generated by the different harvesting and processing techniques (Table 2).10,11 The major types of fat that have been described are macrofat (or millifat), microfat, and nanofat. Macrofat and microfat have become the standard technique for structurally filling soft-tissue defects and are composed of mostly adipocytes and a small population of SVF cells.12 These two types of fat are further divided based on the parcel size generated from the harvesting cannula and port size, where parcel size is defined by the size of the cells and aggregates of cells.10 Generally, macrofat has a parcel size of 2 to 2.5 mm in diameter harvested with a 3-mm harvesting cannula, while microfat is characterized by a parcel size of 1 mm harvested with a 2.1- to 2.4-mm harvesting cannula (Table 3).10 Fractofat is a microfat subtype that is further refined with multiple passages through a 2-mm filter.13 Amid the adipocytes in either macrofat or microfat are fewer than 10% of SVF cells.14 Nanofat is processed mechanically with multiple passes through a 2.4-mm and 1.2-mm connector and one pass through a 400- to 600-µm emulsifier, resulting in a product composed of SVF cells, growth factors, and extracellular matrix, with no adipocytes (Table 3).10 Each component of macrofat, microfat, and nanofat varies in cell size and type, and as a result, each type of fat has a defined role in improving the volume, skin texture, and pigmentation of the aging face.

Table 2. - Comparison of Macrofat, Microfat, and Nanofat Harvesting and Processing Techniques
Variable Coleman Rohrich Marten Mashiko Rohrich Tonnard Tonnard Cohen
Type of fat Macrofat Macrofat (A) Microfat Microfat Fractofat (specialized microfat) Microfat (B) Nanofat Microfat and nanofat
Source Abdomen, thigh Medial thigh Lateral torso (hip, waist, flank, and lateral thigh) Thigh Processed macrofat (A) Lower abdomen Processed microfat (B) Abdomen, thigh
Donor-site preparation 0.1% With 1:400,000 epinephrine No tumescent 0.1% Lidocaine with 1:1,000,000 epinephrine; nontumescent infiltration: 1 cc injected for every 3 cc of anticipated fat harvest 1 Liter normal saline with 1:1,000,000 epinephrine No tumescent Modified Klein solution — 0.05% Lidocaine with 1:500,000 epinephrine
Liposuction pressure Handheld syringe aspiration Handheld syringe aspiration Handheld syringe aspiration (10 cc) using hand-applied gradual low-negative suction High negative-pressure SAL Handheld syringe aspiration High negative-pressure SAL — Handheld syringe aspiration
Cannula size Single-port 3.0-mm cannula Multiport 3-mm with 1-mm holes Multiport 2.1- to 2.4-mm cannula Multiport 3-mm with 2-mm holes Multiport 3-mm with 1-mm holes Multiport 2- to 3-mm with 1-mm holes — Multiport 2.5-mm cannula
Processing technique Centrifugation at 3000 rpm for 3 minutes Centrifugation at 2250 rpm for 1 minute (a) Centrifugation at 1000 rpm for 1 to 3 minutes; no washing or emulsification; high-density fat (bottom 2 cc of centrifuged fat) sequestered and used preferentially for critical areas Centrifugation at 1200 g for 3 minutes, followed by 30 passes between two 2.5-cc syringes connected through Tulip connector with three holes 50 Passes between two 10-cc syringes connected through Tulip connector with two 2-mm-diameter cannula Washed over sterile nylon cloth with 0.5-mm perforations (b) 30 Passes through two 10-mL syringes connected by a 2.4-mm connector; 30 passes through two 10-mL syringes connected by a 1.2-mm connector; final pass through a double 400- or 600-μm emulsifier Washed and decanted, followed by processing with Nano Cube kit
Injection technique 17- or 18-G blunt cannula 2-mm Blunt-tip Micrin cannula 0.7-mm (20-G), 0.9-mm (19-G), 1.2-mm (18-G) blunt single side-hole cannulas Unspecified 2-mm Blunt-tip Micrin cannula 0.7-mm Blunt cannula 27-G needle or microneedling Microneedling or topical biocrème
Recipient-site preparation Unspecified None Nerve blocks with 0.25% bupivacaine with 1:200,00 epinephrine for anterior face; light infiltration with same solution for posterior jawline and lateral face Unspecified None None None None
SAL, suction-assisted lipectomy.

Table 3. - Biological Characteristics and Clinical Indications for Each Type of Fat
Type of Fat Parcel Size Cells Biological Mechanism Indication Treatment Area
Macrofat 2.0 to 2.5 mm Adipocytes (90% to 95%); SVF (5% to 10%) Provides volumetric expansion Large volume loss in deep fat compartments of the face; significant laxity and descent of facial ligaments Temporal region, deep malar fat compartments
Microfat 1.0 mm Adipocytes (90% to 95%); SVF (5% to 10%) Provides volumetric expansion Moderate volume loss in superficial fat compartments of the face; partial laxity and descent of facial ligaments; deep static and dynamic rhytids Forehead, glabella, superior orbital rim, tear trough, suborbicular oculi fat, superficial malar fat compartment, nasolabial fold, perioral skin, marionette line, lips, chin, mandible
Nanofat 400 to 600 µm SVF (100%); adipose stem cells; endothelial cells; pericytes; smooth muscle cells Improved nutrient delivery and toxin clearance; neocollagenesis and reorganization of elastin; clearance of melanin and inhibition of melanin deposition Reduced skin elasticity; superficial static and dynamic rhytids; periorbital hyperpigmentation; skin discoloration Full face and neck
SVF, stromal vascular fraction.

CELLULAR COMPONENTS OF AUTOLOGOUS FAT

Adipocytes are terminally differentiated cells held together by a complex extracellular matrix that allows the tissue to maintain its form even during the harvesting and processing techniques.15 Relative to the other components, adipocytes are large and bulky cells and are thus a good candidate for increasing volume. All adipocytes initially rely on nearby vascular networks for nutrients until angiogenesis occurs.16,17 Beyond energy storage, adipocytes also secrete growth factors, cytokines, and hormones that contribute to vasculogenesis, matrix remodeling, and accelerated wound healing.18–21 Thus, these adipocytes that make up the majority of the cells in macrofat and microfat may not only add volume but also may secrete factors that ultimately support incorporation of the fat.

In comparison, SVF cells contribute to the regenerative properties of autologous fat and have been used in multiple arenas for their effects on inflammation and angiogenesis.22 The ADSCs within the SVF have been shown to have immunomodulatory properties, tissue-regenerative capacity, and angiogenic effects.3,4,23,24 These ADSCs secrete antiinflammatory cytokines and chemokines that improve the engraftment of cells by reducing the number of macrophages that circulate in the recipient site at the time of implantation.23 Additional studies have shown that these ADSCs inhibit melanocyte proliferation and melanin synthesis by downregulating melanogenic enzymes through complex inflammatory pathways. Ultimately, these cells restore normal pigmentation along the basement membrane of the dermis and epidermis.25–27 The secretion of angiogenic growth factors by the ADSCs also appears to act on pericytes, smooth muscle cells, and endothelial cells to undergo neoangiogenesis.28 Together, the SVF cells provide regenerative capacity to the recipient site.

BIOLOGICAL IMPLICATIONS OF THE CELLS IN AUTOLOGOUS FAT

Macrofat and microfat provide significant structural support and tissue bulk to the recipient site when placed precisely where there is volume deficiency. Macrofat is used to target deeper structural volume losses, whereas microfat can be placed more superficially. Nanofat has a high concentration of regenerative cells with the potential to improve skin elasticity and hyperpigmentation and is injected intradermally or subdermally. With facial aging, there is a loss of tissue volume in the fat compartments. The addition of volume in the face through macrofat or microfat grafting provides a more youthful and rejuvenated appearance by restoring the contours of the face. In contrast, the emulsification process to retrieve nanofat releases the SVF cells from the adipocytes and its matrix to interact with the new microenvironment.29 As such, “nanofat” is a misnomer and does not contain fat. The SVF cells are intimately associated with complex pathways that allow for collagen and elastin regeneration to improve skin texture and elasticity and inhibit melanin synthesis to improve skin pigmentation.4,30–32 Meanwhile, the endothelial cells, pericytes, and smooth muscle cells in SVF migrate in a coordinated fashion to undergo neoangiogenesis to generate new networks. The effect of these cells allows for improved clearance of toxins and improved delivery of nutrients to the recipient site to slow facial aging.32 Nanofat contains SVF cells that have the potential to restore normal dermal and epidermal matrix and reverse discoloration caused by the injuries of intrinsic and extrinsic aging.

CLINICAL INDICATIONS AND UTILITY OF AUTOLOGOUS FAT

Given the significant differences in the cellular and biological components of macrofat, microfat, and nanofat, deliberate decisions should be made to utilize the specific type of fat that will correct the specific deformity identified when seeking to restore the youthful facial appearance (Fig. 1). Facial aging has been characterized by the loss of volume in the fat compartments, descent of the soft tissues, and degradation of the facial skin over time.33,34 Recent studies have shown that the deep medial cheek fat compartment is one of the first fat compartments to atrophy during the fourth and fifth decade of life, resulting in superficial fat pseudoptosis, where the overlying superficial fat descends because of loss of support, amplifying the aged appearance35 (Fig. 1). Correction of the deep fat compartment with volume leads to significant improvement in anterior cheek projection and flattening of the nasolabial fold, restoring the youthful appearance of the face.36,37 Macrofat and microfat grafting should be considered in patients with significant atrophy from aging. In particular, patients with temporal hollowing, volume loss in the malar region, and presence of jowls would benefit from deep-plane macrofat or microfat grafting to the temple, deep malar fat compartment, perioral region, chin, and inferior mandibular border (Figs. 1 through ). With respect to the use of macrofat or microfat, it remains to be determined whether macrofat or microfat has greater longevity or better outcomes for improving facial rejuvenation, and additional studies are necessary. To improve deep static and dynamic rhytids in other areas of the face, microfat and its subtypes, such as fractofat, should be placed into the forehead, glabella, superior orbital rims, infraorbital area, superficial lateral fat compartments of the cheek, nasolabial folds, marionette line, lips, and chin (Figs. 2 and 3). A sharp-needle (21 G or 23 G) microfat-grafting technique has been described to inject microfat intradermally or subdermally to fill rhytids observed in the forehead, glabella, lips, and neck.38 The shear bulk of the volume added restores fullness and provides a more youthful appearance. Microfat and its subtype, fractofat, can be used in both superficial and deep planes to add volume and improve contours (Figs. 2 and 3). However, macrofat should only be used in deep planes because of the risk of nodule formation when injected too superficially. Macrofat and microfat can be used alone or in combination depending on the extent of facial aging. One size does not fit all, and one fat grafting technique does not fix all. A personalized approach is required to achieve the optimal outcome. [See Figure, Supplemental Digital Content 1, which shows macrofat and fractofat use to augment deep malar fat deflation and to blend the lid–cheek junction. The patient presented with severe volume loss of the deep malar fat pad and loss of sub–orbicularis oculi fat (center, above and below). He underwent upper and lower lid blepharoplasty, face lift, macrofat grafting to the deep malar compartment (purple), and fractofat (subtype of microfat; green) grafting to the tear trough (above, left). The fat was grafted with a blunt cannula into the cheek–orbital rim junction to disrupt the orbital malar ligament below the orbicular oculi muscle and provide volume to the atrophied fat in the malar fat pad. Significant improvement is evident in the periorbital region at 6 years postoperatively (right, above and below). Procedure performed by Rod Rohrich, MD. Reprinted with permission from Rohrich RJ, Sinno S, Vaca EE. Getting better results in face lifting. Plast Reconstr Surg. 2015;136:27–38, https://links.lww.com/PRS/G401. See Figure, Supplemental Digital Content 2, which shows a patient with secondary face lift with microfat grafting to improve facial contour. A 75-year-old woman with a history of multiple previous face lifts and related procedures presented with concerns of a residual frail, unhealthy, and older appearance caused by uncorrected panfacial atrophy despite her previous surgical procedures (center, above and below). She is shown 1 year 7 months after panfacial microfat grafting (green; above, left) and secondary face lift, neck lift, forehead lift, upper and lower blepharoplasties, and canthopexy. A total of 90 cc of fat was placed. Volumes reflect nonemulsified microfat harvested with a 2.1-mm to 2.4-mm cannula that was centrifuged for 1 to 3 minutes at 1000 rpm. No skin resurfacing or implants were used. A significant improvement in her facial contour can be seen, with a healthy, fit, and feminine appearance secondary to restoration of facial volumes that could not be obtained by face lift alone (right, above and below). Procedure performed by Timothy Marten, MD. Reprinted with permission of the Marten Clinic of Plastic Surgery, https://links.lww.com/PRS/G402.]

F1
Fig. 1.:
Macrofat and microfat grafting to address facial aging. Significant atrophy of the malar eminence and sub–orbicularis oculi fat leads to associated descent of the fat pads over time. The volumetric effects of fat grafting with macrofat or microfat restore the contours of the face, with macrofat used in the deep malar fat pads in both younger and older patients and microfat used in more delicate areas in the superficial malar fat pads. Copyright © 2021 James D. Vargo, MD. Reprinted with permission.
F2
Fig. 2.:
Macrofat and fractofat grafting to improve significant volume loss and restore a youthful appearance. The patient presented with severe volume loss of the central face both in the deep and superficial malar fat and thinning of the suborbicular skin (center, above and below). She underwent face lift, upper and lower lid blepharoplasty, macrofat grafting to the deep and superficial fat compartments (purple), and fractofat (green) grafting to the tear trough and infrabrow area (above, left). Macrofat and fractofat injection improved youthful appearance by increasing the volume of the midface and in the suborbicular region, which camouflaged the thin skin and blended the lid–cheek junction. Results were maintained at 3 years postoperatively (right, above and below). Procedure performed by Rod Rohrich, MD. Reprinted with permission from Rohrich RJ, Mohan R. Role of ancillary procedures in facial rejuvenation. Plast Reconstr Surg Glob Open 2019;7:e2075.
F3
Fig. 3.:
Microfat grafting in a patient who underwent a secondary face lift to restore a youthful appearance. A 62-year-old woman presented with a history of face lift, neck lift, and upper and lower lid blepharoplasties with an incompletely rejuvenated appearance because of residual atrophy of her orbits, midface, perioral area, chin, and jawline (center, above and below). The patient is shown 12 months after panfacial microfat grafting (green; above, left), along with secondary face lift, neck lift, forehead lift, trichloroacetic acid lower eyelid peel, and perioral dermabrasion. A total of 70 cc of fat was placed, and volumes reflect nonemulsified microfat harvested with a 2.1- to 2.4-mm cannula that was centrifuged for 1 to 3 minutes at 1000 rpm. Note the improved facial contour and more feminine and healthy appearance (right, above and below). The synergistic effect of the face lift and fat grafting provides a result that arguably could not be achieved with either procedure alone. Procedure performed by Timothy Marten, MD. Reprinted with permission of the Marten Clinic of Plastic Surgery.

With nanofat, the biological activity of the mechanically dissociated SVF cells acts on the skin to improve rhytids and hyperpigmentation (Fig. 4).8 Nanofat has been shown to increase skin elasticity and reduce superficial rhytids.8 In addition, improved skin texture, wrinkles, pore size, firmness, and skin hydration were observed to still be present 12 months after nanofat grafting.31,39–42 Thus, nanofat should be injected intradermally or subdermally in the forehead, glabellar, and perioral region to improve skin texture and lessen rhytids. Nanofat also can be delivered with a microneedling device, where percutaneous microchannels are made into the papillary dermis of the skin and a manual pumping mechanism delivers small quantities of prepared nanofat simultaneously.43,44 Nanofat can improve skin quality and texture; the combination of macrofat or microfat with nanofat potentially can provide volumetric augmentation and target the overlying skin (Fig. 4). The number of studies objectively assessing nanofat is limited, and additional studies are warranted (Table 4). [See Figure, Supplemental Digital Content 3, which shows microfat and nanofat grafting to restore facial atrophy and improve skin quality in the periorbital and perioral region. A 61-year-old woman presented 15 years after a primary face lift with significant volume loss of the periorbital and perioral areas and loss of skin elasticity and hyperpigmentation (center, above and below). The patient underwent transconjunctival lower lid blepharoplasty, face lift, neck lift, lip lift, microfat grafting (green), sharp needle intradermal microfat grafting (green), and nanofat microneedling to the full face and neck (yellow; above, left). Sharp needle intradermal fat grafting with microfat was performed on the horizontal lines in the neck. The patient had improved volume in the periorbital and perioral region because of the microfat grafting, with less hyperpigmentation and improved skin quality secondary to the nanofat microneedling after 8 months (right, above and below). Procedure performed by Patrick Tonnard, MD, https://links.lww.com/PRS/G403.]

Table 4. - Clinical Studies Using Nanofat for Facial Rejuvenation
Study Level of Evidence No. of Patients Indication for Nanofat Isolation of Nanofat Macrofat or Microfat Grafting in the Same Procedure Face Lift and Blepharoplasty in the Same Procedure Nanofat Mixed with Other Solutions? Area Injected In Vitro Results Clinical Outcomes
Tonnard et al. (2013)8 4 67 Skin rejuvenation 30 Passes between 2- and 20-mL syringes connected to a Luer-Lok connector Yes Yes No Perioral skin (58%), glabellar skin (23%), breast cleavage (11%), scars (6%), lower eyelids (2%) No viable adipocytes in nanofat; nanofat was further enzymatically processed for in vitro studies with collagenase II for 45 minutes; most of the nanofat cells (95%) were nonhematopoietic stem cells (Cd34-) in nanofat with similar proliferation and multilineage differentiation capacity when compared with macrofat and microfat Remarkable improvement in skin quality (lightening of dark periorbital skin, reduced superficial perioral rhytids) 6 months postoperatively; no complications (eg, infection, fat cyst, granuloma, or unwanted side effects)
Oh et al. (2014)46 4 19 Dark periorbital hyperpigmentation 30 Passes between 2- and 1-mL syringes connected to Luer-Lok connector Yes Yes No Lower eyelid NA Improvement in skin darkening and skin texture of the lower lid; high patient and surgeon satisfaction; no visible lumps of fat, contour irregularities, or fat necrosis
Liang et al. (2018)31 2 231 (129 HA; 103 nanofat) Skin rejuvenation 3 Minutes between 2- and 20-mL syringes connected to a Luer-Lok connector No No Yes: PRP (percentage not specified) Not specified No additional enzymatic processing of nanofat; nanofat cells expressed adipose stem cell surface markers (CD29, CD44, CD49D, CD73, CD90, and CD105) without expression of CD34, CD45, or CD106 Compared nanofat with HA injection; higher patient and surgeon satisfaction in nanofat group; Visia skin image analyzer revealed greater improvement in skin wrinkles, texture, and pores in nanofat treatment group at 1, 12, and 24 months; SOFT5.5 skin test instrument demonstrated improved moisture and elasticity in nanofat treatment group at 1, 12, and 24 months; pigmentation complication was observed in one patient (1%) at 6 and 12 months
Menkes et al. (2020)39 4 50 Skin rejuvenation Tulip kit; 30 passes between 2- and 20-mL syringes connected to a 2.4-, 1.4-, and then 1.2-mm filter and a final pass through a Tulip nanotransfer 600-/400-µm filter Yes No Yes: 20% PRP Lower lid and orbital area, upper lip, lower lip, cheek, nasolabial fold, jaw line, parotid area, and temporal area Nanofat was further digested with Liberase MTF-S, which contains collagenase; cell viability was 90%; no changes in cell viability or number of adipose stem cells comparing nanofat with microfat; increased number of endothelial cells in the nanofat compared with microfat Clinical results apparent between 2 and 4 weeks after injection and continued until 6 months postoperatively; improvement in skin quality with respect to texture, elasticity, glow, firmness, fine wrinkles, and skin hydration; minor complications with redness and edema between 2 and 4 days, some bruises, and pain at the donor site; lower-lid nodule in one patient (2%) that improved spontaneously; no infection, fat cyst, or granulomas; biopsies showed an increased dermal cellularity, vascular density, and elastic and collagen fiber density; no changes in epidermal thickness or mucin content of the dermis

F4
Fig. 4.:
Microfat and nanofat grafting to improve volume loss and restore skin quality. A 54-year-old woman presented with significant fat atrophy of her central face, loss of skin elasticity, hyperpigmentation, and static facial rhytids (center, above and below). She underwent face lift, lip lift, extensive microfat grafting, and sharp-needle intradermal microfat grafting (green); erbium laser resurfacing; and nanofat microneedling to the full face and neck (yellow; above left). Sharp-needle intradermal fat grafting with microfat was performed on the white roll and philtrum, upper lip rhytids, frontal and glabellar lines, and neck lines. Nanofat microneedling was composed of 11 cc of nanofat, 2 cc of hyaluronic acid, and 100 U of botulinum toxin (2 cc) and 2 cc of vitamin C. The combination of skin resurfacing and nanofat microneedling appeared to work synergistically to improve healing time, reduce postoperative erythema, and improve skin quality. The patient had improved skin pigmentation and a natural, more youthful appearance after face lift with microfat and nanofat grafting after 10 months (right, above and below). Procedure performed by Patrick Tonnard, MD.

SPECIAL CONSIDERATIONS FOR THE PERIORBITAL REGION

In patients with periorbital hyperpigmentation, it is essential to define the underlying cause of the hyperpigmentation to ensure that the appropriate type of fat is used in the proper planes. With aging, the periorbital and malar cheek fat pads atrophy earlier in life compared with neighboring fat pads, resulting in the V-deformity. Periorbital hyperpigmentation can be caused by severe deflation in the deep malar fat compartments resulting in increased shadowing within the infraorbital region and enhanced definition of orbicularis retaining ligament. Injection of macrofat into the deep malar fat compartments can fill the void caused by the volume loss and reduce the shadowing that results in the illusion of hyperpigmentation.45

In contrast, visibility of the orbicularis oculi muscle secondary to atrophy of the sub–orbicularis oculi fat can give the illusion of hyperpigmentation in the infraorbital area. In this case, microfat, and its subtypes such as fractofat, could be used to increase bulk in the delicate infraorbital space to conceal the orbicularis oculi muscle.13 The smaller parcel size of microfat minimizes nodule formation and complications that can be associated with macrofat grafting. Microfat can be injected deep in the periosteal plane or in small aliquots in the midplane to provide volume while minimizing irregularities that can develop in this region. Nanofat should be considered for patients with true hyperpigmentation of the skin in the infraorbital region caused by increased deposition of melanin in the lower eyelid skin. Nanofat has been shown to improve hyperpigmentation of the lower eyelids and can be used for skin rejuvenation in the infraorbital region.46 Defining the underlying etiology of the hyperpigmentation is necessary to determine the type of fat required to correct the deformity. Expectation management is essential because complications, such as ecchymosis and edema, may present in the immediate postoperative period and can persist for up to 6 weeks.46 The periorbital region is a challenging area to address aesthetically and requires an individualized approach to treatment, including critical decisions regarding the type of fat to be used, the volume of fat to be grafted, and the location where the fat is placed.

SPECIAL CONSIDERATIONS FOR THE PERIORAL REGION

The perioral region can be particularly challenging to address because of the bony and soft-tissue changes that occur with aging and the static and dynamic considerations in this region. Patients with significant volume loss in the perioral region because of skeletal changes may present with maxillary retrusion and may benefit from macrofat grafting around the lips and oral commissure. However, it is inadvisable that macrofat be used in the lip mucosa because of the potential for nodule formation. In contrast, women with deep static and dynamic perpendicular perioral rhytids, flattening of the philtral columns, or loss of definition in cupid’s bow would benefit from microfat grafting to the perioral region. In these cases, sharp needle intradermal fat grafting with microfat is an option to improve deep static rhytides.38 Nanofat grafting would be most beneficial in patients with superficial perioral rhytids caused by loss of skin elasticity. In some cases, it is advantageous to perform perioral macrofat, microfat, and nanofat grafting in the same patient, if the patient presents with severe volume loss and skin changes. Again, an individualized approach should be adopted for autologous fat grafting so that the needs of each patient can be addressed through this form of precision medicine: the right type of fat graft needs to be given to the right patient, at the right time, in the right volume, and in the right plane.

DISCUSSION

The use of fat grafting during face-lift procedures has been popularized in the United States by surgeons such as Marten and Elyassnia,47,48 Cohen et al,49,50 Rohrich and colleagues,51,52 and Stuzin.53 In many cases, macrofat and microfat grafting performed with face lifts alone can achieve dramatic effects. Recent studies have demonstrated the efficacy of nanofat grafting in improving age-related changes of the face, including skin thinning, loss of skin elasticity, hyperpigmentation, and sun-damaged spots.54–56 These studies suggest that nanofat may be the missing link to improve skin texture and pigmentation, while the largest effect of macrofat and microfat grafting is to provide volumetric enhancement.43,44,57 Nevertheless, it is imperative to recognize that additional high-quality basic science and clinical studies are necessary to further understand the biology of the grafted fat and to optimize outcomes for patients.

While the majority of the regenerative effects of nanofat are likely from the SVF cells,58 the effect of nanofat should not be directly compared with enzymatically processed SVF or ADSCs. Since Zuk and colleagues discovered ADSCs, considerable interest in ADSCs exists because of their regenerative and restorative properties. However, methods to dissociate these regenerative cells enzymatically have harmful effects on the cell; these harmful effects not only affect cell survival but can also alter the biological processes, which limits their regenerative capacity and clinical efficacy. Therefore, we caution against direct comparison between nanofat and enzymatically harvested SVF cells or ADSCs and recommend considering nanofat separate from enzymatically dissociated cell products until appropriate validation studies have been performed.

Although technological advances have provided tools to maximize the efficacy of autologous fat grafting, a thorough understanding of the types of fat and their clinical utilization is necessary to achieve optimal outcomes. Furthermore, nothing can replace the importance of having an artistic eye when it comes to fat grafting, particularly when considering the volume to inject. In addition, the type of face-lift procedure performed along with the autologous fat grafting may affect the volume injected, and achievement of a balanced, more youthful appearance also requires the surgeon’s artistic eye. However, we highlighted several key concepts to provide the principles and foundation of fat grafting to achieve acceptable results. As we have observed, there are slight variabilities in the precise methodology among the authors of and contributors to this article; no superior protocol has emerged, as subtle differences among techniques do not appear to alter the overall graft take. In particular, the use of lidocaine does not appear to alter the overall graft take. Previous studies have shown that appropriate processing of the harvested fat appears to remove the lidocaine and does not affect the survival of the adipocytes and SVF cells. Additional studies comparing these methodologies will be necessary to determine whether there is an optimal processing method. In addition, experience and ongoing critique of one’s own outcomes can lead to further improvements for each patient. Autologous fat grafting delivered through an individualized approach allows for restoration of all the soft-tissue layers. Each patient still requires an individualized approach that identifies the underlying pathology and the type of autologous fat that is best suited to correct the concern. Improper injection of fat in the wrong plane can result in undercorrection or overcorrection of the soft tissue, leading to contour irregularities. Knowledge of the anatomy of the aging face and a clear definition of the aesthetic goals of the procedure will allow for a precise approach to fat grafting to achieve the desired aesthetic outcomes.

As we continue to observe the progress made in surgical and nonsurgical facial rejuvenation, the utilization of various fat-grafting techniques and types of fat grafting for individualized use will continue to improve outcomes. The ideal filler for aesthetic surgery is autologous fat grafting, because it is inexpensive, easily harvested, natural in appearance and texture, immunologically compatible, and long lasting. Furthermore, with the advent of nanofat grafting, we potentially have the next scientific advancement in facial rejuvenation, as nanofat can be delivered in a seemingly easy manner to restore the quality of the skin. Nonetheless, it is important to recognize that although additional studies are necessary to determine the precise mechanism by which nanofat improves skin texture and hyperpigmentation, the clinical outcomes have been promising. The combination of autologous macrofat and microfat grafting with nanofat grafting can serve as a standalone rejuvenation technique or as a powerful complement to face lifts and provide plastic surgeons with another tool to achieve even better aesthetic results for their patients.

DISCLOSURE

Dr. Rohrich receives instrument royalties from Eriem Surgical, Inc., and book royalties from Thieme Medical Publishing; is a clinical and research study expert for Allergan, Inc., Galderma, and MTF Biologics; is a medical monitor for Merz North America; and owns Medical Seminars of Texas, LLC. Dr. Tonnard receives book royalties from Thieme Medical Publishing and instrument royalties from Tulip Medical Products, Inc. The remaining authors have no financial interest to disclose in relation to the context of this article.

ACKNOWLEDGMENT

The authors would like to acknowledge Dr. Timothy Marten for his contributions and insightful discussions, which assisted with the formation of the ideas for this article.

REFERENCES

1. The Aesthetic Society. Procedural statistics (2015–2018). Available at: www.surgery.org/media/statistics.
2. Strong AL, Cederna PS, Rubin JP, Coleman SR, Levi B. The current state of fat grafting: a review of harvesting, processing, and injection techniques. Plast Reconstr Surg. 2015;136:897–912.
3. Zuk PA, Zhu M, Mizuno H, et al. Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng. 2001;7:211–228.
4. Kilroy GE, Foster SJ, Wu X, et al. Cytokine profile of human adipose-derived stem cells: expression of angiogenic, hematopoietic, and pro-inflammatory factors. J Cell Physiol. 2007;212:702–709.
5. Hwang C, Pagani CA, Das N, et al. Activin A does not drive post-traumatic heterotopic ossification. Bone 2020;138:115473.
6. Shaker RF, Abdel Aal ARM, El Gazzar KMA, Abu Zahra FAK, Elshahat A. In vitro comparative study of emulsified fat grafts. Eplasty. 2020;20:e1–e1.
7. Aronowitz JA, Lockhart RA, Hakakian CS. Mechanical versus enzymatic isolation of stromal vascular fraction cells from adipose tissue. Springerplus. 2015;4:713–713.
8. Tonnard P, Verpaele A, Peeters G, Hamdi M, Cornelissen M, Declercq H. Nanofat grafting: basic research and clinical applications. Plast Reconstr Surg. 2013;132:1017–1026.
9. Yoshimura K, Shigeura T, Matsumoto D, et al. Characterization of freshly isolated and cultured cells derived from the fatty and fluid portions of liposuction aspirates. J Cell Physiol. 2006;208:64–76.
10. Cohen SR, Hewett S, Ross L, et al. Regenerative cells for facial surgery: biofilling and biocontouring. Aesthet Surg J. 2017;37:S16–S32.
11. Pourang A, Rockwell H, Karimi K. New frontiers in skin rejuvenation, including stem cells and autologous therapies. Facial Plast Surg Clin North Am. 2020;28:101–117.
12. Coleman SR, Lam S, Cohen SR, Bohluli B, Nahai F. Fat grafting: challenges and debates. Atlas Oral Maxillofac Surg Clin North Am. 2018;26:81–84.
13. Rohrich RJ, Mahedia M, Hidalgo D, Shah N. The evolving role of blending of the lid-cheek junction in lower blepharoplasty. Plast Reconstr Surg. 2018;142:377–382.
14. Yoshimura K, Suga H, Eto H. Adipose-derived stem/progenitor cells: roles in adipose tissue remodeling and potential use for soft tissue augmentation. Regen Med. 2009;4:265–273.
15. Suga H, Matsumoto D, Inoue K, et al. Numerical measurement of viable and nonviable adipocytes and other cellular components in aspirated fat tissue. Plast Reconstr Surg. 2008;122:103–114.
16. Trayhurn P, Wang B, Wood IS. Hypoxia in adipose tissue: a basis for the dysregulation of tissue function in obesity? Br J Nutr. 2008;100:227–235.
17. Hausman GJ, Richardson RL. Adipose tissue angiogenesis. J Anim Sci. 2004;82:925–934.
18. Walther TC, Farese RV. Lipid droplets and cellular lipid metabolism. Annu Rev Biochem. 2012;81:687–714.
19. Hauner H. Secretory factors from human adipose tissue and their functional role. Proc Nutr Soc. 2005;64:163–169.
20. Halberg N, Wernstedt-Asterholm I, Scherer PE. The adipocyte as an endocrine cell. Endocrinol Metab Clin North Am. 2008;37:753–768.
21. Shibata S, Tada Y, Asano Y, et al. Adiponectin regulates cutaneous wound healing by promoting keratinocyte proliferation and migration via the ERK signaling pathway. J Immunol. 2012;189:3231–3241.
22. Gimble JM, Bunnell BA, Chiu ES, Guilak F. Concise review: adipose-derived stromal vascular fraction cells and stem cells: let’s not get lost in translation. Stem Cells 2011;29:749–754.
23. Kapur SK, Katz AJ. Review of the adipose derived stem cell secretome. Biochimie. 2013;95:2222–2228.
24. Sterodimas A, de Faria J, Nicaretta B, Pitanguy I. Tissue engineering with adipose-derived stem cells (ADSCs): current and future applications. J Plast Reconstr Aesthet Surg. 2010;63:1886–1892.
25. Kim DW, Jeon BJ, Hwang NH, et al. Adipose-derived stem cells inhibit epidermal melanocytes through an interleukin-6-mediated mechanism. Plast Reconstr Surg. 2014;134:470–480.
26. Jeong JH. Adipose stem cells and skin repair. Curr Stem Cell Res Ther. 2010;5:137–140.
27. Li P, Guo X. A review: therapeutic potential of adipose-derived stem cells in cutaneous wound healing and regeneration. Stem Cell Res Ther. 2018;9:302–302.
28. Suh A, Pham A, Cress MJ, et al. Adipose-derived cellular and cell-derived regenerative therapies in dermatology and aesthetic rejuvenation. Ageing Res Rev. 2019;54:100933.
29. Sesé B, Sanmartín JM, Ortega B, Matas-Palau A, Llull R. Nanofat cell aggregates: a nearly constitutive stromal cell inoculum for regenerative site-specific therapies. Plast Reconstr Surg. 2019;144:1079–1088.
30. Charles-de-Sá L, Gontijo-de-Amorim NF, Maeda Takiya C, et al. Antiaging treatment of the facial skin by fat graft and adipose-derived stem cells. Plast Reconstr Surg. 2015;135:999–1009.
31. Liang ZJ, Lu X, Li DQ, et al. Precise intradermal injection of nanofat-derived stromal cells combined with platelet-rich fibrin improves the efficacy of facial skin rejuvenation. Cell Physiol Biochem. 2018;47:316–329.
32. James IB, Coleman SR, Rubin JP. Fat, stem cells, and platelet-rich plasma. Clin Plast Surg. 2016;43:473–488.
33. Stuzin JM. MOC-PSSM CME article: face lifting. Plast Reconstr Surg. 2008;121:1–19.
34. Gosain AK, Klein MH, Sudhakar PV, Prost RW. A volumetric analysis of soft-tissue changes in the aging midface using high-resolution MRI: implications for facial rejuvenation. Plast Reconstr Surg. 2005;115:1143–1152; discussion 1153.
35. Rohrich RJ, Avashia YJ, Savetsky IL. Prediction of facial aging using the facial fat compartments. Plast Reconstr Surg. 2021;147:38S–42S.
36. Rohrich RJ, Pessa JE, Ristow B. The youthful cheek and the deep medial fat compartment. Plast Reconstr Surg. 2008;121:2107–2112.
37. Rohrich RJ, Durand PD, Dayan E. The lift-and-fill facelift: superficial musculoaponeurotic system manipulation with fat compartment augmentation. Clin Plast Surg. 2019;46:515–522.
38. Zeltzer AA, Tonnard PL, Verpaele AM. Sharp-needle intradermal fat grafting (SNIF). Aesthet Surg J. 2012;32:554–561.
39. Menkes S, Luca M, Soldati G, Polla L. Subcutaneous injections of nanofat adipose-derived stem cell grafting in facial rejuvenation. Plast Reconstr Surg Glob Open 2020;8:e2550.
40. Rihani J. Microfat and nanofat: when and where these treatments work. Facial Plast Surg Clin North Am. 2019;27:321–330.
41. Pallua N, Kim B-S. Microfat and lipoconcentrate for the treatment of facial scars. Clin Plast Surg. 2020;47:139–145.
42. Sese B, Sanmartin JM, Ortega B, Matas-Palau A, Llull R. Nanofat cell aggregates: a nearly constitutive stromal cell inoculum for regenerative site-specific therapies. Plast Reconstr Surg. 2019;144:1079–1088.
43. Verpaele A, Tonnard P, Jeganathan C, Ramaut L. Nanofat needling: a novel method for uniform delivery of adipose-derived stromal vascular fraction into the skin. Plast Reconstr Surg. 2019;143:1062–1065.
44. Tonnard P, Verpaele A, Carvas M. Fat grafting for facial rejuvenation with nanofat grafts. Clin Plast Surg. 2020;47:53–62.
45. Rohrich RJ, Mohan R. Role of ancillary procedures in facial rejuvenation. Plast Reconstr Surg Glob Open 2019;7:e2075.
46. Oh DS, Kim DH, Roh TS, Yun IS, Kim YS. Correction of dark coloration of the lower eyelid skin with nanofat grafting. Arch Aesthetic Plast Surg. 2014;20:92–96.
47. Marten T, Elyassnia D. Facial fat grafting: why, where, how, and how much. Aesthetic Plast Surg. 2018;42:1278–1297.
48. Marten TJ, Elyassnia D. Fat grafting in facial rejuvenation. Clin Plast Surg. 2015;42:219–252.
49. Cohen SR, Womack H, Ghanem A. Fat grafting for facial rejuvenation through injectable tissue replacement and regeneration: a differential, standardized, anatomic approach. Clin Plast Surg. 2020;47:31–41.
50. Cohen SR, Womack H. Injectable tissue replacement and regeneration: anatomic fat grafting to restore decayed facial tissues. Plast Reconstr Surg Glob Open 2019;7:e2293–e2293.
51. Ramanadham SR, Rohrich RJ. Newer understanding of specific anatomic targets in the aging face as applied to injectables: superficial and deep facial fat compartments: an evolving target for site-specific facial augmentation. Plast Reconstr Surg. 2015;136:49S–55S.
52. Pezeshk RA, Stark RY, Small KH, Unger JG, Rohrich RJ. Role of autologous fat transfer to the superficial fat compartments for perioral rejuvenation. Plast Reconstr Surg. 2015;136:301e–309e.
53. Stuzin JM, Rohrich RJ, Dayan E. The facial fat compartments revisited: clinical relevance to subcutaneous dissection and facial deflation in face lifting. Plast Reconstr Surg. 2019;144:1070–1078.
54. Lambros V. Observations on periorbital and midface aging. Plast Reconstr Surg. 2007;120:1367–1376; discussion 1377.
55. Gonzalez-Ulloa M, Flores ES. Senility of the face: basic study to understand its causes and effects. Plast Reconstr Surg. 1965;36:239–246.
56. Lambros V. Facial aging: a 54-year, three-dimensional population study. Plast Reconstr Surg. 2020;145:921–928.
57. Stuzin JM. Discussion: nanofat grafting: basic research and clinical applications. Plast Reconstr Surg. 2013;132:1027–1028.
58. Ghiasloo M, Lobato RC, Díaz JM, Singh K, Verpaele A, Tonnard P. Expanding clinical indications of mechanically isolated stromal vascular fraction: a systematic review. Aesthet Surg J. 2020;40:NP546–NP560.

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