Sage Journals: Discover world-class research

Abstract

Autologous fat grating is a widely-accepted method to correct soft tissue deficiency. Although fat transplantation shows excellent biocompatibility and simple applicability, the relatively low retention rate caused by fat necrosis is still a challenge. The vasculature is integral after fat grafting, serving multiple crucial functions. Rapid and effective angiogenesis within grafts is essential for supplying oxygen necessary for adipocytes’ survival. It facilitates the influx of inflammatory cells to remove necrotic adipocytes and aids in the delivery of regenerative cells for adipose tissue regeneration in fat grafts. The vasculature also provides a niche for interaction between adipose progenitor cells and vascular progenitor cells, enhancing angiogenesis and adipogenesis in grafts. Various methods, such as enriching grafts with diverse pro-angiogenic cells or utilizing cell-free approaches, have been employed to enhance angiogenesis. Beige and dedifferentiated adipocytes in grafts could increase vessel density. This review aims to outline the function of vasculature in fat grafting and discuss different cell or cell-free approaches that can enhance angiogenesis following fat grafting.

Keywords

fat grafting cell-enrichment therapy angiogenesis vasculature function

Graphical Abstract

Introduction

Autologous fat grafting is a procedure performed by plastic surgeons with the primary goal of restoring soft tissue defects and correcting facial deficiency¹. In recent years, it has become a prevalent surgical method in cosmetic surgery for facial rejuvenation^2,3, gluteal augmentation³, and breast reconstructive surgery⁴. Compared with other biologically active gels such as hyaluronic acid, it shows better biocompatibility and is relatively economical for patients^3,5.

Suction-harvested adipose tissue, while effective for extraction, faces a challenge due to its lack of a natural vascular network essential for oxygenation and nutrient distribution. This deficiency can lead to undesired volume reduction, fibrosis, and the development of oil cysts due to fat necrosis^6–9. Therefore, timely vascularization reconstruction is a crucial prerequisite for achieving high survival rates after fat grafting¹⁰. Based on the different extent of ischemic, three distinct zones emerge after fat grafting: survival (periphery), regeneration (intermediate), and necrosis (central). Initially, the boundary between the necrotic and regenerating areas is not distinctly demarcated, and the regenerative potential of the regeneration area relies on the timely ingrowth of capillaries that provide oxygen^11,12. Except for providing essential oxygen, reconstructed vasculature exerts multiple functions in the fat grafting model, such as providing circulating stem cells and inflammatory cells from circulation to remodel the grafts¹³ and provide a niche for the adipogenesis of perivascular cells, such as adipose-derived stem cells (ASCs), preadipocytes and pericytes, and so on (Figure 1).

Figure 1.

Vasculature function in fat grafting. Vasculature provides a timely and enough supply of oxygen and nutrients to reduce the apoptosis of adipocytes in the early hypoxia and ischemic environment and transport unessential triglycerides to the blood to avoid lipo-toxin-induced inflammation. Vasculature delivers early-stage inflammatory cells that could clear the damaged cells and extracellular matrix to give regenerative cells a better regenerative microenvironment. Vasculature provides a perivascular niche for the mesenchymal stem cells (especially ASCs) to proliferate and differentiate into regenerated adipocytes and promote neovasculogenesis through paracrine pro-angiogenic factors or transdifferentiate into ECs. ASCs, adipose-derived stem cells; ECs, endothelial cells.

This review focuses on the vascular function that contributes to the survival and remodeling of grafts, summarizes the cellular-derived angiogenic stimulation that benefits the neovascularization after fat grafting, and provides a future perspective for the following exploration.

Vasculature Function After Fat Grafting

Delivering Essential Oxygen Crucial and Nutrient-Waste Exchange for Adipocyte Survival

Following grafting, adipose tissue enters a state of severe ischemic and hypoxia, characterized by low oxygen levels¹³. Most adipocytes in the graft begin to die within 24 h ASCs remain viable for up to 72 h but still demonstrate limited resistance to hypoxia. Hence, the angiogenesis process within the first 72 h after grafting assumes paramount importance. This period is crucial for ensuring the timely delivery of oxygen and nutrients to rescue the transplanted cells, ultimately influencing their fate^13,14.

Before angiogenesis occurs, grafted adipose tissue can only receive oxygen through plasmatic diffusion from surrounding vessels¹². It takes 3–7 days for the peripheral vessels to ingrow into the outer layer of the grafts¹⁵. Moreover, newly formed blood vessels in grafts provide nutrients and exchange metabolic products after 2 days of implantation^16,17. To guarantee sufficient blood and oxygen delivery to the graft, each fat graft droplet must interface with a capillary recipient site in a 1:1 ratio to establish a successful fat–recipient complex. Excessive fat droplets relative to capillary recipient sites can result in inadequate neovasculogenesis, potentially leading to fat resorption and necrosis¹⁸. Therefore, smaller grafts tend to exhibit better survival rates compared with larger grafts due to the higher surface-to-volume ratio, which allows for a larger area of the graft to be in contact with the vascular bed and facilitates more efficient vascularization^16,19. Adequate oxygen supply is also essential for macrophages to effectively clear dead adipocytes and lipid droplets by maintaining proper fatty acid oxidation²⁰. Moreover, timely vascularization provides a channel for transporting excess triglycerides into the bloodstream, thereby reducing lipotoxin-induced massive inflammation^21–24.

Supplying Circulating Stem Cells and Inflammatory Cells From Circulation to Adipose Tissue Graft

After grafting, macrophages infiltrate into the grafts in the early stage to promote remodeling by clearing free fatty acids, phagocytosing dead cells, and digesting damaged extracellular matrix (ECM) components. These activities create space for regenerative cells such as ASCs, endothelial progenitor cells (EPCs), and other stem cells, which can enhance angiogenesis and adipogenesis^25–27. Traditional perspectives considered that macrophages infiltrated into the inflammatory site are mainly from circulation²⁸. In recent investigations, it has been established that during the initial week post-surgery, the proportion of infiltrated macrophages derived from the fascia surpasses that of macrophages originating from circulation. Notably, experiments involving the removal of fascia from the back of recipient mice in the fat grafting model have revealed a subsequent reduction in the number of early-stage macrophage infiltrations²⁹.

New vasculature also provides channels for the circulating stem cells, new vessels within the graft predominantly arise from recipient vessels growing into the graft, rather than from the reassembly of donor endothelial cells (ECs) or the reconnection of recipient and donor vessels³⁰. In the nascent phase of angiogenesis within the grafts, conduits are established to facilitate the ingress of blood-derived hematopoietic cells and ASCs, thereby enabling their pivotal contribution to the initial stages of regeneration³¹. From a long-term perspective, most of the CD34+ cells are derived from the recipient during adipogenesis rather than in the initial grafts³⁰. Besides, bone marrow-derived endothelial progenitors are also capable of migrating to the ischemic location, then differentiate into ECs and finally become capillaries in the superficial graft zones^32,14.

Providing NICHE for Adipogenesis and Angiogenesis in Grafts

Mounting evidence indicates that ASCs along with other types of mesenchymal stem cells (MSCs) are closely linked to vascular and perivascular niches in different tissues³³. During human embryonic development, adipocytes first appear from emergent vascular networks³⁴. Lineage-tracing studies have demonstrated that adipocyte progenitors reside within the walls of adult mouse adipose tissue capillaries^35,36. Co-implantation of adipose tissue with ASCs differentiated toward an endothelial phenotype significantly enhances both the survival and angiogenesis of the transplanted adipose tissue³⁷. Coculturing EPCs with ASCs enhances EPCs’ tube formation capacities and angiogenesis markers such as tie2 and vascular endothelial growth factor (VEGF)-A. ASCs’ combination enhances the function of EPCs in the fat transplant rather than directly differentiating into them³⁸. ASCs, often found alongside microvessels, stabilize the perivascular niche by differentiating into pericytes and expressing key pericyte markers such as α-smooth muscle actin (α-SMA), neural/glial antigen 2 (NG2), or platelet-derived growth factor receptor β (PDGFRβ)³⁹. Indeed, perivascular cells such as vascular smooth muscle cells and pericytes have been proven to give rise not only to white adipocytes but also to differentiate into beige adipocytes^40,41, which are located around the vasculature, sharing the same location as pericytes⁴². Inducing more pericytes in transplanted adipose tissue could also enhance the adipocytes’ viability and increase the survival of adipocytes⁴³, which could also shed light on the function of pericytes as the orchestrators of vasculature and adipogenesis⁴⁰. Besides, the reciprocal interaction between preadipocytes and endothelial cells (ECs) was promoted by the infiltration of blood vessels⁴⁴. Preadipocytes induce and guide EC migration through fibroblast growth factor (FGF)- and vascular endothelial growth factor (VEGF)-dependent pathways⁴⁵. Conversely, the VEGF-triggered signaling pathway in ECs promotes preadipocyte differentiation via a paracrine mechanism⁴⁶. Differentiation from preadipocytes to mature adipocytes is associated with increased production of angiogenic factors that induce robust angiogenic responses⁴⁷.

Cell-Enrichment Therapy in Fat Grafting

Enhancing fat grafts with diverse cell types that demonstrate significant angiogenic activity is presently recognized as a highly effective approach for enhancing the neovasculogenesis of transplanted adipose tissue. To achieve this objective, macrophages, ASCs, and other cells from the stromal vascular fraction (SVF) have been proposed¹⁰.

Endothelial Progenitor Cell

EPCs obtained from humans can integrate into sites of active angiogenesis, thereby enhancing the growth of collateral vessels in ischemic tissues³². The sprouting of the peripheral vessels toward the graft typically spans a period of 3–7 days with the recruitment of EPCs^15,48–50. Bone marrow-derived EPCs migrate to a target ischemic site, then differentiate into ECs, and finally become capillaries for the superficial graft zones^32,14. However, the EPCs cannot migrate through the outer graft layers to access the ischemic center, meaning that the inner part depends on its progenitor cells to induce capillary formation^51,52. Combining EPCs with fat grafts has been shown to enhance early-stage angiogenesis while minimizing the development of oil cysts and fibrosis. In their pursuit of EPCs with heightened regenerative potential, Asahara et al. utilized a serum-free medium comprising five different vasculogenic components (VEGF, stem cell factor, Flt-3 ligand, thrombopoietin and interleukin 6)—designed as a Quality and Quantity culture (QQ culture) for cell expansion and vasculogenic of EPCs⁵³. Maxim et al. discovered that when transplanted with QQ cultured c-Kit+ Sca-1+ Lin− (KSL) EPCs, grafts get better angiogenesis and less fibrosis and inflammation levels⁵². Their further study proves that the QQ culture method also enhanced the vasculogenic potential of peripheral blood mononuclear cells (PBMNCs) by enriching them with EPCs and anti-inflammatory M2-monocyte/macrophages. Grafts enriched with MNC-QQ exhibit the highest vessel density and secrete increased levels of VEGF paracrine⁵¹.

Macrophage

In the normal response to injury of multiple tissue types, monocytes infiltrate the wound site and differentiate into macrophages that exhibit a highly pro-inflammatory phenotype, at later stages, the macrophage population undergoes a shift in phenotype to pro-regenerative status^54,55 (Figure 2). The conventional classification of macrophage phenotype can be delineated into two primary types: the pro-inflammatory M1-polarized phenotype and the regenerative, pro-angiogenic M2-polarized phenotype⁵⁶. Another article has challenged this viewpoint, both M1 and M2 macrophages can facilitate the vascular reconstruction and vascular remodeling process. Primary human M1 macrophages secrete high levels of VEGF and increase the expression of VEGFR2 on EC⁵⁷. In the M2 macrophage subset, M2a macrophages secrete PDGF-BB, stabilizing pericytes and enhancing EC sprouting. Meanwhile, M2c macrophages release MMP9, contributing to basement membrane degradation and ECM remodeling. Besides, M2c-derived osteopontin promotes EC proliferation and migration⁵⁸.

Figure 2.

Macrophage function in microenvironment after fat grafting. IL, interleukin; iNOS, inducible nitric oxide synthase; VEGF, vascular endothelial growth factor; VEGFR2, vascular endothelial growth factor receptor 2; ECM, extracellular matrix; TGF-β, transforming growth factor-β; CCL, C-C motif chemokine ligand; CXCL, C-X-C motif chemokine; PPARγ, peroxisome proliferatoractivated receptor γ; PDGF-BB, platelet-derived growth factor subunit B.

Monocyte infiltration into the grafts shows the same pattern as the common injury. During the first week after surgery, grafts enter an “inflammatory phase,” M1 macrophages emerge within the first week to facilitate the clearance of necrotic tissue and release pro-inflammatory factors^26,59,60. Then in the regeneration stage (weeks 2–4 and beyond), there is a gradual transition as M1 macrophages are gradually replaced by M2 macrophages, which shift toward regulating angiogenesis and suppressing inflammation^59,61,62. M2 macrophages facilitate the sprouting of pre-existing blood vessels and their subsequent fusion to form new functional vasculature. This process is mediated by the direct localization of M2 macrophages at the fusion sites, where they engage with integrins on the EC surfaces, thereby promoting the connection of adjacent tip cells⁶³. Moreover, as previously mentioned, osteopontin derived from M2 macrophages also contributes to promoting endothelial tip cell linkage by interacting with integrins on the surface of ECs⁶⁴. Numerous factors in fat grafting contribute to the enhancement of M2 macrophage polarization (as depicted in the figure), indirectly fostering angiogenesis and shifting the pro-inflammatory microenvironment toward an anti-inflammatory state post-fat grafting^65–67.

Manipulating macrophage infiltration by using liposome-clodronate/Macrophage-Colony Stimulating Factor (M-CSF) resulted in either incompetent or improved angiogenesis after grafting⁶⁸. Early macrophage infiltration in grafts demonstrates a robust correlation with the onset of early neovasculogenesis. The early presence of macrophages plays a pivotal role in attracting more Sca-1⁺/CD45⁺ stem cells capable of generating ECs and neovessels possibly through the CXCL12/CXCR4 axis⁶⁸, and CXCL12 are also capable of promoting EPCs mediated angiogenesis and reduce the fat apoptosis⁶⁹. Chen et al. highlight the reciprocal mechanism that early angiogenesis is also essential for CXCL12-dependent ASC recruitment. Besides, inhibiting CXCL12 expression at the donor site in the autologous fat grafting model promotes increased ASC infiltration into the recipient site, thereby raising graft retention rates³¹.

Macrophage cell-enrichment therapy increases the vascularization of grafts. Both M0, M1, and M2a could enhance the revascularized ability in transplanted vascular grafts or glutaraldehyde-crosslinked scaffolds^57,58. Mixing M2 macrophage with white adipose transplant also increases the angiogenesis after 3 months of grafting⁷⁰. QQ-cultured PBMNCs including EPCs and M2 macrophage also increase the neovascularization in the grafts both in 1 week or 7 weeks after grafting. Moreover, QQ-cultured PBMNCs promote angiogenesis through upregulating the expression of VEGF-A in the graft and QQ-cultured MNCs are capable of becoming a part of the newly formed vessel⁵¹.

ASCs’ Function in Cell-Assisted Lipotransfer (CAL)

Advanced regenerative therapies focus on harnessing the potential of stem cells, for direct application to damaged sites, known as cell therapy, or for tissue engineering by utilizing suitable scaffolds as carriers for these cells which holds distinct advantages over differentiated cells⁷¹. Adipose tissue has been historically considered a passive energy storage. However, currently, it is well known that it is also a rich source of multipotent ASCs^72,73. ASCs express the surface MSC markers CD73, CD90, CD105, and STRO-1 and display a broad range of regenerative properties including a high replication capacity and potential to differentiate into various cell lineages, as well as the ability to secrete numerous pro-angiogenic growth factors, which are essential for promoting the formation of vascular networks and supporting the vasculature hemostasis^74,75.

Yoshimura’s group pioneered the utilization of exogenous ASCs in fat grafting and named this procedure “cell-assisted lipotransfer (CAL).” By elevating the local concentration of ASCs during the initial phase of grafting, CAL enhances angiogenesis and consequently improves graft retention rates⁷⁶.

VEGF (also known as VEGF-A)⁷⁷ serves as an excellent target for enhancing the paracrine function of ASCs in promoting the formation and stabilization of vascular networks within grafts. VEGF induces proliferation, sprouting, migration, and tube formation of ECs and also keeps the ECs survive^77–79 (Figure 3). Oxygen deprivation activates hypoxia-inducible factor 1α (HIF1α) in macrophages and ASCs, leading to the expression of VEGF-A, which stimulates the specification of ECs into tip cells and stalk cells. These tip cells, characterized by their high motility, actively sense environmental guidance cues and guide the nascent sprouts by forming filopodia. Stalk cells provide structural support to the nascent capillary as part of the body of the nascent sprout. Mechanistically, VEGF binds to the VEGFR2 receptor expressed on tip cells, promoting their proliferation, migration, and adhesion. Concurrently, the phosphorylation of the VEGFR2 receptor activates the DLL4-NOTCH1 signaling pathway in adjacent ECs, which inhibits VEGFR2 expression in these neighboring cells. This inhibition promotes the stalk cell phenotype, which matures and stabilizes the nascent sprout by recruiting more pericytes and ECM⁷⁷.

Figure 3.

The function of VEGF in the angiogenesis. Under hypoxia conditions, macrophages and ASCs-derived VEGF specify the expression of endothelial cells into tip cells and stalk cells. Tip cells function as a guider to build connections with the adjacent vessels and stalk cells function as a supporter to give structural support.

HIF1α-VEGF axis in ASCs mediates the angiogenesis in grafts. C-reactive protein (CRP) affected the proliferation and pro-angiogenic paracrine activity of ASCs through upregulating VEGF-A expression by activating HIF-1α⁸⁰. Moreover, inhibiting the negative regulator of VEGF-A with a DII4-neutralizing antibody enhances the angiogenesis of grafts and exhibits synergistic effects when combined with concurrent ASC supplementation⁸¹.

Besides, the overexpression of miR-126 in ASCs results in increased expression of VEGF. Transplanting adipose tissue with exogenous ASCs that overexpress miR-126 enhances the angiogenic capacity of grafts. This effect is mediated through the Erk/Akt pathway rather than via only VEGF signaling⁸². Besides, transplanting white adipose tissue (WAT) with exogenous ASCs transduced with VEGF also increases the angiogenesis and capillary density in the grafts⁷⁵.

The trans-differentiation of ASCs into ECs and pericytes represents another mechanism contributing to angiogenesis in grafts following CAL. EC-derived basic FGF (bFGF) triggers endothelial differentiation in ASCs by inhibiting miR-145 through the PI3K/AKT/FOXO1 signaling pathway. This inhibition relieves suppression of the angiogenic transcription factor ETS1, thereby promoting the differentiation of ASCs into ECs⁸³. Downregulation of miR-145 in ASCs induced vascular network formation and neovessels incorporation in ischemic muscle in vivo⁸³. Combined treatment of ASCs with VEGF and bone morphogenetic protein-4 (BMP4) under hypoxia conditions induces the differentiation of ASCs into vascular EC lineage and acquires the ability of mature EC such as tube formation, low-density lipoprotein (LDL) uptake, and nitric oxide secretion^84,85. Besides, treating ASCs with TGF-β induces their differentiation into pericytes. Transdifferentiating more ASC into pericytes could enhance neovascularization in CAL⁴³. Moreover, culturing ASCs with endothelial growth medium (EGM-2) in vitro and transplanting them with human umbilical vascular ECs (HUVEC) subcutaneously into immunodeficient mice increases the HUVEC survival and enhances the vasculature function⁸⁶.

There remains uncertainty regarding the mechanism by which exogenous ASCs become involved in the angiogenesis process during CAL, particularly in the crucial early stages following surgery. Yuan et al. gave a hypothesis that exogenous ASCs in free fat transplantation may not directly engage in angiogenesis through differentiating into mature adipocytes or ECs. However, they are believed to enhance the survival rate of graft-resident interstitial cells through paracrine effects⁸⁷. Gan et al.³⁸ also demonstrated that ASCs combined with EPCs enhance the function of EPCs in the fat transplant rather than directly differentiate into them. Conversely, using a tracing method to reveal the differentiation of donor ASCs in the CAL at 4 weeks after transplantation, Hong et al.⁸⁸ demonstrated that donor-derived ASC can directly differentiate into ECs and adipocytes to take part in angiogenesis. Evidence suggests that ASC-derived ECs can be assembled into newly formed vessels⁸⁹. CD34+CD31– cells sorted from SVF cells can differentiate into ECs in vitro and incorporate into the ischemic hindlimb vasculature in vivo⁹⁰. While intravenously injecting ASCs immediately after surgery does increase vascular density in the graft, their primary mechanism is through paracrine signaling⁹¹. Yan Huang et al. further demonstrated that intravenous injection of ASCs immediately after surgery and again at 2 weeks post-surgery led to increased retention of graft volume and vascular density. Simultaneously, grafts that are intravenously injected with ASC exhibit higher expression levels of CXCL12 and CXCR4⁹².

ASC-exosome is another strategy to enhance early angiogenesis. Recent research findings indicate that exosomes derived from stem cells exhibit comparable or even superior performance to that of MSCs^93–95. In comparison with ASCs, ASC-exos offer several advantages. These exosomes are small extracellular vesicles (50–200 nm) that can be sterile filtered and stored as safe, ready-to-use biological products with high biological activity. Furthermore, their use circumvents issues associated with cell therapy, such as limited cell survival, senescence-induced genetic instability, and the risk of unfavorable differentiation¹⁰.

ASC-exos also facilitate angiogenesis within grafted adipose tissue through both direct and indirect means^96–98. Directly, ASC-exos promote angiogenesis and upregulate early-stage inflammation, which is beneficial for clearing dead cells and creating a pro-regenerative environment⁹⁶. ASC-exos enhance the proliferation, migration, and tube formation capacity of HUVEC through the HIF-1α/VEGF signaling pathway⁹⁹. Moreover, hypoxia-treated ASC-exo contains more pro-angiogenic growth factors than normal ASC-exos⁹⁸, which fits the microenvironment after grafting¹³. Injectable thermosensitive hydrogels show significant promise as carriers for exosomes¹⁰⁰. Exosomes encapsulated within hydrogels can be utilized to regulate their release over extended periods, enhancing their therapeutic effects and efficacy⁹⁹. Human ASCs-derived exosomes (hASC-Exos) encapsulated in PF-127 hydrogel have been shown to enhance the neovascularization of autologous fat grafts⁹⁹.

Indirectly, ASC-exosomes may increase the vessel density and browning of WAT in grafts by inducing more M2 macrophage differentiation and suppressing M1 macrophage differentiation^101–103.

Adipocytes’ Pro-angiogenic Function in Fat Grafting

Brown and Beige Adipocytes

Adipose tissue consists of three primary types of adipocytes. White adipocytes, predominant in WAT, are unilocular cells specializing in lipid storage. In contrast, beige and brown adipocytes are multilocular, mitochondria-rich cells specialized in energy dissipation through non-shivering thermogenesis¹⁰⁴.

Beige adipocytes arise from subcutaneous WAT (sWAT) depots in response to β-adrenergic stimulation, dietary factors, or exposure to cold, a reversible process known as browning or beiging¹⁰⁵. Although originating from myogenic transcription factor 5 (Myf5)-negative mesodermal stem cells like white adipocytes, beige adipocytes functionally resemble brown adipocytes. They possess the capacity to convert chemical energy into heat in response to specific stimuli¹⁰⁶.

Transplantation of brown adipose tissue has been widely utilized in the treatment of diabetes, obesity, and their associated complications^107–110, owing to the high secretion of autocrine and paracrine factors¹¹¹. Compared with the transplantation of white adipocytes, the transplantation of rosiglitazone-induced beige adipocytes also offers metabolic benefits by alleviating obesity-induced inflammation and increasing the proportion of high-density lipoprotein (HDL)¹¹². Moreover, the beigeing of perivascular adipose tissue caused by vascular injury alleviates inflammation by expressing neuregulin 4, which activates M2 macrophage polarization¹¹³.

Brown and beige adipocytes with a higher surface-to-volume ratio can facilitate the process of diffusion and increase the relative exposure area to oxygen^114–116 (Figure 4). In comparison with traditional white adipocytes, PPAR-γ-induced beige adipocytes exhibit higher tolerance to hypoxic and inflammatory environments¹¹⁵. Beige adipose grafts are demonstrated with increased angiogenesis, recruiting fewer macrophages in the early stages, and consequently, exhibiting enhanced survival after the transplantation¹¹⁵. Overexpressing VEGF-A in adipocytes enhances its angiogenesis and induces beige fat formation in the doxycycline(dox)-inducible, VEGF-A overexpressing transgenic (VEGF-Tg) mouse model. Additionally, transplantation of subcutaneous adipose tissue from dox-inducible VEGF-Tg mice into the interscapular region of wild-type C57/BL6J mice, followed by a 3-week high-fat diet regimen, demonstrates enhanced adipocyte survival and reduced macrophage infiltration in VEGF-A-overexpressing adipose grafts¹¹⁷. Moreover, transplantation of cold-stimulated induced beige adipose tissue reveals heightened angiogenesis levels and a relatively diminished inflammatory state¹¹⁶.

Figure 4.

The difference between beige adipocyte and white adipocyte. White adipocytes are unilocular cells specializing in lipid storage with one big lipid droplet and low mitochondria density. The beige adipocytes are multiple lipid droplets with the expression of UCP1 and a high level of mitochondria density. Under hypoxia conditions, beige adipocytes show better endurance and less apoptosis compared with white adipocytes. higher surface-to-volume ratio makes it more easily to get enough blood supply and therefore more easily to survive after fat grafting. Beige adipocytes could also induce more M2 macrophage polarization. UCP1, uncoupling protein 1.

However, brown adipocytes within grafts may struggle to withstand hypoxic and ischemic conditions, leading to sustained high levels of inflammation due to the “whitening” of brown adipose tissue which results in adipocyte death and apoptosis^95,118. Transplanting brown adipose tissue (BAT) typically exhibits lower retention rates compared with beige adipose tissue and even WAT, partially due to the heightened inflammatory state^115,119. However, Zheng et al.¹²⁰ found that transplanting grafts that contain a mixture of BAT and WAT increased the recruitment of M2 macrophage and the expression of VEGF-A. Moreover, inducing the browning of white adipose transplant after grafting could also increase the expression of VEGF in the grafts and enhance angiogenesis¹¹⁹.

Dedifferentiated Adipocytes (DAs)

Traditional views depict adipogenic differentiation as an irreversible process where MSCs transition into mature adipocytes. Mature adipocytes were believed to possess limited plasticity, primarily responding to stimuli with changes in size rather than undergoing further dedifferentiation^121,122. Through the “ceiling culture” method, mature adipocytes exhibit dedifferentiation, ultimately transforming into progenitor cells expressing markers characteristic of adipose stem cells. Similar to stem cells, these progenitor cells exhibit proliferative and redifferentiation capabilities. Additionally, they demonstrate multilineage differentiation potential, differentiating into various cell types both in vitro and in vivo^123–126 (Figure 5).

Figure 5.

The pro-angiogenic function of dedifferentiated adipocyte. DAs in was founded in the SVFs in the grafts after 1 week of fat transplantation. In vitro ceiling culture also induces adipocyte dedifferentiation. With enough vascular supply, DAs could redifferentiate into mature adipocytes. DAs acquire the ability to differentiate into ECs or pericytes and also enhance the proliferation, cell migration, and tube formation of ECs through secretory proteins such as VEGF and HGF. HGF, hepatocyte growth factor; VEGF, vascular endothelial growth factor.

DAs are demonstrated to have the same adipogenesis and angiogenesis potential compared with ASCs in fat grafts¹²⁷. DAs can promote EC tube formation in vitro and facilitate angiogenesis in ischemic hindlimbs, potentially through the secretion of factors such as VEGF-A and hepatocyte growth factor (HGF). Moreover, when co-cultured with ECs, DAs express pericyte markers. TGF-β can also induce DAs to differentiate into mural cells in vitro¹²⁸. In 1 week after fat grafting, the mature adipocytes in grafts were found to differentiate into DAs¹²⁹, which can also spontaneously undergo mesenchymal-to-endothelial transition and differentiate into ECs. This transition enables them to participate in neovascularization, particularly in response to BMP4 and BMP9 signaling¹³⁰. Interestingly, host-derived adipocytes contribute to fat graft regeneration by migrating from the recipient inguinal fat pad into the grafts and undergoing dedifferentiation, which can be augmented by internal expansion to enhance the retention rate of fat grafts¹³¹. Moreover, given a sufficient vascular supply, DAs have the potential to undergo redifferentiation into mature adipocytes^132,133, which might be a way for them to survive in the regeneration area.

Future Perspective of Vascularization in Fat Grafting

As mentioned above, timely and complete neovascularization in the grafts determines the fate of cells in the grafts. Therefore, numerous efforts have been made to enhance early-stage revascularization by adding different kinds of angiogenic cells such as EPCs, ASCs, or other in vitro QQ cultured ECs and peripheral mononuclear cells^31,51,52. A more detailed understanding of the precise mechanism through which these cells integrate into the microenvironment’s vasculature after grafting, and their interactions with other progenitor cells to support adipogenesis, ECM remodeling, and adipocyte regeneration, remains a crucial focus for future research. Finding a stable source of vessel-forming progenitor cells and adding them to the grafts provides a novel strategy to rescue the ischemic microenvironment after grafting¹³⁴. Enriching the ASCs filtered with more angiogenic potential might be another choice to get better vascularization after grafting. A subpopulation of human ASC marked by CD34⁺CD146⁺ enhances the induction of tube formation of HUVECs in vitro. Enriching fat with CD34⁺CD146⁺ ASCs shows better vascularization compared with CD34⁺ unsorted SVF cells¹³⁵. Another two populations of distinct vascular stem/progenitor cells (VSPCs) are identified from the ASCs subpopulation, marked respectively by CD45^–Ter119^–Tie2⁺PDGFRa^–CD31⁺CD105^highSca1^low and CD45^–Ter119^–Tie2⁺PDGFRa⁺CD31CD105^lowSca1^low. Cotransplantation of these two types of cells forms functional vessels that improve perfusion of the mouse hindlimb ischemia model. However, there is still little article reporting that applying the different angiogenic potential of ASCs in the CAL¹³⁴. Identifying novel progenitor cells that can develop into fully functional blood vessels, continues to be a significant challenge^136–138. The significant variability in angiogenic potential among identical types of progenitor cells, such as distinct subtypes of ASCs, underscores the need for further investigation^136,139.

In recent years, single-cell RNA sequence (scRNA-seq) has become a popular method to detect the deconvolve tissue heterogeneity^140–142. Quite a lot of studies have been applied in analyzing the cells of SVFs in the WAT and brown adipose tissue¹⁴³. Although susceptible to certain technical noise, single-cell RNA sequencing (scRNA-seq) offers vital insights into cellular populations, gene expression patterns, and variations in cellular states that may occur under diverse physiological conditions^141,144. It is quite useful to filter the subpopulations according to their functions in the tissue and identify more angiogenic subpopulations in the SVFs¹⁴⁵. The latest spatial transcriptomic to build a high-resolution cellular map for further exploration¹⁴⁶. However, there is still no research applying the scRNA-seq in fat grafting to reveal the dynamic cellular map and the function of different stem and progenitor cells.

Another emerging technology in recent years is 3D-culture in vitro, compared with cells cultured as monolayers (2D), 3D spheroids can more accurately mimic the in vivo cell growth environment while retaining their biological characteristics. 3D-cultured ASCs demonstrate increased proliferation potential and reduced senescence compared with conventional culture methods. Co-culturing 3D-cultured ASCs with HUVECs enhances their neovascularization capabilities. Employing 3D-cultured ASCs in CAL improves the survival and neovascularization of ASCs¹⁴⁷.

Conclusion

This review summarizes the current knowledge about vasculature function in fat grafting as well as the different cellular compositions that improve revascularization during the process of fat regeneration. The results provide a greater understanding of the application of different cell-enrichment therapies. However, there are still a lot of unknown mechanisms for vasculature function in fat grafting⁴⁴.

Footnotes

Acknowledgements

Figures in this review were created with . Thanks for the support of Dr. Luo for this manuscript.

Authors’ Note

Zhang Xining as an Independent First Author in this review.

Author Contributions

Z.X. performed the literature search and wrote the manuscript and figures. L.S. conceived the project and revised the manuscript.

Availability of data and material

The authors confirm that the data supporting the findings of this study are available within the article.

Ethical Approval

This study was approved by our institutional review board.

Statement of Human and Animal Rights

This article does not contain any studies with human or animal subjects.

Statement of Informed Consent

There are no human subjects in this article and informed consent is not applicable.

Declaration of Conflicting Interests

The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding

The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: The author(s) was supported by the National Natural Science Foundation of China (82102356), Outstanding Young Medical Talents Training Funding Project of The First Affiliated Hospital of Harbin Medical University (2021J05).

ORCID iDs

Zhang Xining

Luo Sai

References

Khouri

Jr Khouri

. Current clinical applications of fat grafting. Plast Reconstr Surg. 2017;140(3):466e–86e.

Coleman

. Facial augmentation with structural fat grafting. Clin Plast Surg. 2006;33(4):567–77.

Coleman

. Structural fat grafting: more than a permanent filler. Plast Reconstr Surg. 2006;118(suppl 3):108S–20S.

Kaoutzanis

Xin

Ballard

Welch

Momoh

Kozlow

Brown

Cederna

Wilkins

. Autologous fat grafting after breast reconstruction in postmastectomy patients: complications, biopsy rates, and locoregional cancer recurrence rates. Ann Plast Surg. 2016;76(3):270–75.

Illouz

. Body contouring by lipolysis: a 5-year experience with over 3000 cases. Plast Reconstr Surg. 1983;72(5):591–97.

Hörl

Feller

Biemer

. Technique for liposuction fat reimplantation and long-term volume evaluation by magnetic resonance imaging. Ann Plast Surg. 1991;26(3):248–58.

Carpaneda

Ribeiro

. Study of the histologic alterations and viability of the adipose graft in humans. Aesthetic Plast Surg. 1993;17(1):43–47.

Choi

Small

Levovitz

Lee

Fadl

Karp

. The volumetric analysis of fat graft survival in breast reconstruction. Plast Reconstr Surg. 2013;131(2):185–91.

Matsumoto

Sato

Gonda

Takaki

Shigeura

Sato

Aiba-Kojima

Iizuka

Inoue

Suga

Yoshimura

. Cell-assisted lipotransfer: supportive use of human adipose-derived cells for soft tissue augmentation with lipoinjection. Tissue Eng. 2006;12(12):3375–82.

10.

Limido

Weinzierl

Harder

Menger

Laschke

. Fatter is better: boosting the vascularization of adipose tissue grafts. Tissue Eng Part B Rev. 2023;29(6):605–22.

11.

Kato

Mineda

Eto

Doi

Kuno

Kinoshita

Kanayama

Yoshimura

. Degeneration, regeneration, and cicatrization after fat grafting: dynamic total tissue remodeling during the first 3 months. Plast Reconstr Surg. 2014;133(3):303e–13e.

12.

Khouri

Jr Khouri

Lujan-Hernandez

Khouri

Lancerotto

Orgill

. Diffusion and perfusion: the keys to fat grafting. Plast Reconstr Surg Glob Open. 2014;2(9):e220.

13.

Yoshimura

Eto

Kato

Doi

Aoi

. In vivo manipulation of stem cells for adipose tissue repair/reconstruction. Regen Med. 2011;6(suppl 6):33–41.

14.

Eto

Kato

Suga

Aoi

Doi

Kuno

Yoshimura

. The fate of adipocytes after nonvascularized fat grafting: evidence of early death and replacement of adipocytes. Plast Reconstr Surg. 2012;129(5):1081–92.

15.

Saunders

Keller

Dunsker

Mayfield

. Survival of autologous fat grafts in humans and in mice. Connect Tissue Res. 1981;8(2):85–91.

16.

Liao

Marra

Rubin

. Application of platelet-rich plasma and platelet-rich fibrin in fat grafting: basic science and literature review. Tissue Eng Part B Rev. 2014;20(4):267–76.

17.

Mashiko

Yoshimura

. How does fat survive and remodel after grafting. Clin Plast Surg. 2015;42(2):181–90.

18.

Shih

Davis

Winocour

. The science of fat grafting. Semin Plast Surg. 2020;34(1):5–10.

19.

Chang

Lanni

Mirzabeigi

Bucky

. Large-volume fat grafting: identifying risk factors for fat necrosis. Plast Reconstr Surg. 2022;150(5):941e–49e.

20.

Pietiläinen

Naukkarinen

Rissanen

Saharinen

Ellonen

Keränen

Suomalainen

Götz

Suortti

Yki-Järvinen

Oresic

, et al Global transcript profiles of fat in monozygotic twins discordant for BMI: pathways behind acquired obesity. PLoS Med. 2008;5(3):e51.

21.

Niu

Zhang

Lai

Huang

Guo

Yuan

Gao

Chang

. Lipolysis inhibition improves the survival of fat grafts through ameliorating lipotoxicity and inflammation. Faseb J. 2024;38(5):e23520.

22.

Niu

Lai

Chen

Gao

Yuan

. A short-term high-fat diet improved the survival of fat grafts in mice by promoting macrophage infiltration and angiogenesis. Front Cell Dev Biol. 2022;10:856839.

23.

Marra

Svegliati-Baroni

. Lipotoxicity and the gut-liver axis in NASH pathogenesis. J Hepatol. 2018;68(2):280–95.

24.

Mineda

Kuno

Kato

Kinoshita

Doi

Hashimoto

Nakanishi

Yoshimura

. Chronic inflammation and progressive calcification as a result of fat necrosis: the worst outcome in fat grafting. Plast Reconstr Surg. 2014;133(5):1064–72.

25.

Mok

Feng

Wang

Cai

. Decreased serum estrogen improves fat graft retention by enhancing early macrophage infiltration and inducing adipocyte hypertrophy. Biochem Biophys Res Commun. 2018;501(1):266–72.

26.

Martinez-Santibañez

Lumeng

. Macrophages and the regulation of adipose tissue remodeling. Annu Rev Nutr. 2014;34:57–76.

27.

Duffield

Lupher

Thannickal

Wynn

. Host responses in tissue repair and fibrosis. Annu Rev Pathol. 2013;8:241–76.

28.

Chakarov

Lim

Tan

Lim

See

Lum

Zhang

Foo

Nakamizo

Duan

Ting Kong

, et al Two distinct interstitial macrophage populations coexist across tissues in specific subtissular niches. Science. 2019;363(6432):eaau0964.

29.

Zou

Zhao

Chen

Wang

Dong

Gao

. Fascia promotes adipose tissue regeneration by improving early macrophage infiltration after fat grafting in a mouse model. Plast Reconstr Surg. 2023;152(3):446e–57e.

30.

Dong

Peng

Chang

Zhan

Zeng

Zhang

. The angiogenic and adipogenic modes of adipose tissue after free fat grafting. Plast Reconstr Surg. 2015;135(3):556e–67e.

31.

Chen

Wang

Zou

Zhao

Dong

. Early angiogenesis-dependent cxcl12 attracts adipose-derived stem cells to promote the repair of fat grafting in a mouse model. Plast Reconstr Surg. 2023;152(2):363–72.

32.

Asahara

Murohara

Sullivan

Silver

van der Zee

Witzenbichler

Schatteman

Isner

. Isolation of putative progenitor endothelial cells for angiogenesis. Science. 1997;275(5302):964–67.

33.

Cao

. Angiogenesis modulates adipogenesis and obesity. J Clin Invest. 2007;117(9):2362–68.

34.

Crandall

Hausman

Kral

. A review of the microcirculation of adipose tissue: anatomic, metabolic, and angiogenic perspectives. Microcirculation. 1997;4(2):211–32.

35.

Han

Lee

Jin

Lim

Kim

Chang

Shibuya

Kim

Koh

. The spatiotemporal development of adipose tissue. Development. 2011;138(22):5027–37.

36.

Gupta

Mepani

Kleiner

Khandekar

Cohen

Frontini

Bhowmick

Cinti

Spiegelman

. Zfp423 expression identifies committed preadipocytes and localizes to adipose endothelial and perivascular cells. Cell Metab. 2012;15(2):230–39.

37.

Harris

Plastini

Kappy

Ortiz

Chang

Brown

Carpenter

Zhang

. Endothelial differentiated adipose-derived stem cells improvement of survival and neovascularization in fat transplantation. Aesthet Surg J. 2019;39(2):220–32.

38.

Gan

Liu

Zhou

Huang

Zhao

. Effects of adipose-derived stromal cells and endothelial progenitor cells on adipose transplant survival and angiogenesis. PLoS ONE. 2022;17(1):e0261498.

39.

Rautiainen

Laaksonen

Koivuniemi

. Angiogenic effects and crosstalk of adipose-derived mesenchymal stem/stromal cells and their extracellular vesicles with endothelial cells. Int J Mol Sci. 2021;22(19):10890.

40.

Picoli

Birbrair

. Pericytes as the orchestrators of vasculature and adipogenesis. Genes. 2024;15(1):126.

41.

Min

Kady

Nam

Rojas-Rodriguez

Berkenwald

Kim

Noh

Kim

Cooper

Fitzgibbons

Brehm

, et al Human “brite/beige” adipocytes develop from capillary networks, and their implantation improves metabolic homeostasis in mice. Nat Med. 2016;22(3):312–18.

42.

Jiang

Berry

Tang

Graff

. Independent stem cell lineages regulate adipose organogenesis and adipose homeostasis. Cell Rep. 2014;9(3):1007–22.

43.

Cai

Wei

Yang

Wang

Lin

. MiR-24-3p regulates the differentiation of adipose-derived stem cells toward pericytes and promotes fat grafting vascularization. FASEB J. 2023;37(5):e22935.

44.

Cao

. Angiogenesis and vascular functions in modulation of obesity, adipose metabolism, and insulin sensitivity. Cell Metab. 2013;18(4):478–89.

45.

Borges

Müller

Momeni

Stark

Torio-Padron

. In vitro analysis of the interactions between preadipocytes and endothelial cells in a 3D fibrin matrix. Minim Invasive Ther Allied Technol. 2007;16(3):141–48.

46.

Fukumura

Ushiyama

Duda

Tam

Krishna

Chatterjee

Garkavtsev

Jain

. Paracrine regulation of angiogenesis and adipocyte differentiation during in vivo adipogenesis. Circ Res. 2003;93(9):e88–97.

47.

Castellot

Jr Karnovsky

Spiegelman

. Differentiation-dependent stimulation of neovascularization and endothelial cell chemotaxis by 3T3 adipocytes. Proc Natl Acad Sci U S A. 1982;79(18):5597–601.

48.

Rossatti

. Revascularisation and phagocytosis in free fat autografts: an experimental study. Br J Plast Surg. 1960;13:35–41.

49.

Pan

Zhen

Zhang

Shu

Han

Guo

. Enhancement of viability of fat grafts in nude mice by endothelial progenitor cells. Dermatol Surg. 2006;32(12):1437–43.

50.

Sunaga

Sugawara

Katsuragi-Tomioka

Kobayashi

. The fate of nonvascularized fat grafts: histological and bioluminescent study. Plast Reconstr Surg Glob Open. 2013;1(6):e40.

51.

Geeroms

Fujimura

Aiba

Orgun

Arita

Kitamura

Senda

Mizuno

Hamdi

Tanaka

. Quality and quantity-cultured human mononuclear cells improve human fat graft vascularization and survival in an in vivo murine experimental model. Plast Reconstr Surg. 2021;147(2):373–85.

52.

Geeroms

Hamdi

Hirano

Hagiwara

Fujimura

Mizuno

Tanaka

. Quality and quantity-cultured murine endothelial progenitor cells increase vascularization and decrease fibrosis in the fat graft. Plast Reconstr Surg. 2019;143(4):744e–55e.

53.

Masuda

Iwasaki

Kawamoto

Akimaru

Ishikawa

Shizuno

Sato

Ito

Horii

Ishida

, et al Development of serum-free quality and quantity control culture of colony-forming endothelial progenitor cell for vasculogenesis. Stem Cells Transl Med. 2012;1(2):160–71.

54.

Willenborg

Lucas

van Loo

Knipper

Krieg

Haase

Brachvogel

Hammerschmidt

Nagy

Ferrara

Pasparakis

. CCR2 recruits an inflammatory macrophage subpopulation critical for angiogenesis in tissue repair. Blood. 2012;120(3):613–25.

55.

Arnold

Henry

Poron

Baba-Amer

van Rooijen

Plonquet

Gherardi

Chazaud

. Inflammatory monocytes recruited after skeletal muscle injury switch into antiinflammatory macrophages to support myogenesis. J Exp Med. 2007;204(5):1057–69.

56.

Kong

Gao

. Macrophage polarization: a key event in the secondary phase of acute spinal cord injury. J Cell Mol Med. 2017;21(5):941–54.

57.

Graney

Ben-Shaul

Landau

Bajpai

Singh

Eager

Cohen

Levenberg

Spiller

. Macrophages of diverse phenotypes drive vascularization of engineered tissues. Sci Adv. 2020;6(18):eaay6391.

58.

Spiller

Anfang

Spiller

Nakazawa

Daulton

Vunjak-Novakovic

. The role of macrophage phenotype in vascularization of tissue engineering scaffolds. Biomaterials. 2014;35(15):4477–88.

59.

Wang

Chen

Zhu

Chen

Guan

Yao

Wang

Gao

Dong

. The effects of macrophage-mediated inflammatory response to the donor site on long-term retention of a fat graft in the recipient site in a mice model. J Cell Physiol. 2020;235(12):10012–23.

60.

Kim

Nair

. Macrophages in wound healing: activation and plasticity. Immunol Cell Biol. 2019;97(3):258–67.

61.

Cheng

Luan

Wang

. M1/M2 macrophages play different roles in adipogenic differentiation of PDGFRα(+) preadipocytes in vitro. Aesthetic Plast Surg. 2019;43(2):514–20.

62.

Satoh

Kidoya

Naito

Yamamoto

Takemura

Nakagawa

Yoshioka

Morii

Takakura

Takeuchi

Akira

. Critical role of Trib1 in differentiation of tissue-resident M2-like macrophages. Nature. 2013;495(7442):524–28.

63.

Arroyo

Iruela-Arispe

. Extracellular matrix, inflammation, and the angiogenic response. Cardiovasc Res. 2010;86(2):226–35.

64.

Senger

Ledbetter

Claffey

Papadopoulos-Sergiou

Peruzzi

Detmar

. Stimulation of endothelial cell migration by vascular permeability factor/vascular endothelial growth factor through cooperative mechanisms involving the alphavbeta3 integrin, osteopontin, and thrombin. Am J Pathol. 1996;149(1):293–305.

65.

Novak

Koh

. Macrophage phenotypes during tissue repair. J Leukoc Biol. 2013;93(6):875–81.

66.

Arabpour

Saghazadeh

Rezaei

. Anti-inflammatory and M2 macrophage polarization-promoting effect of mesenchymal stem cell-derived exosomes. Int Immunopharmacol. 2021;97:107823.

67.

Dang

Wang

Jiao

Wang

Dou

Song

. Effects of melatonin on fat graft retention through browning of adipose tissue and alternative macrophage polarization. Aesthetic Plast Surg. 2023;47(4):1578–86.

68.

Cai

Feng

Liu

Zhou

. Early macrophage infiltration improves fat graft survival by inducing angiogenesis and hematopoietic stem cell recruitment. Plast Reconst Surg. 2018;141(2):376–86.

69.

Hamed

Egozi

Dawood

Keren

Kruchevsky

Ben-Nun

Gilhar

Brenner

Ullmann

. The chemokine stromal cell-derived factor-1α promotes endothelial progenitor cell-mediated neovascularization of human transplanted fat tissue in diabetic immunocompromised mice. Plast Reconstr Surg. 2013;132(2):239e–50.

70.

Phipps

Gebremeskel

Gillis

Hong

Johnston

Bezuhly

. Alternatively activated M2 macrophages improve autologous Fat Graft survival in a mouse model through induction of angiogenesis. Plast Reconstr Surg. 2015;135(1):140–49.

71.

Weissman

. Stem cells are units of natural selection for tissue formation, for germline development, and in cancer development. Proc Natl Acad Sci U S A. 2015;112(29):8922–28.

72.

Zuk

Zhu

Mizuno

Huang

Futrell

Katz

Benhaim

Lorenz

Hedrick

. Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng. 2001;7(2):211–28.

73.

Zuk

Zhu

Ashjian

De Ugarte

Huang

Mizuno

Alfonso

Fraser

Benhaim

Hedrick

. Human adipose tissue is a source of multipotent stem cells. Mol Biol Cell. 2002;13(12):4279–95.

74.

Zhao

Zeng

Xiong

Zhang

. Deciphering the emerging roles of adipocytes and adipose-derived stem cells in fat transplantation. Cell Transplant. 2021;30:963689721997799.

75.

Gao

Ogawa

Yang

. Improvement of the survival of human autologous fat transplantation by using VEGF-transfected adipose-derived stem cells. Plast Reconstr Surg. 2009;124(5):1437–46.

76.

Yoshimura

Sato

Aoi

Kurita

Hirohi

Harii

. Cell-assisted lipotransfer for cosmetic breast augmentation: supportive use of adipose-derived stem/stromal cells. Aesthetic Plast Surg. 2020;44(4):1258–65.

77.

Pérez-Gutiérrez

Ferrara

. Biology and therapeutic targeting of vascular endothelial growth factor A. Nat Rev Mol Cell Biol. 2023;24(11):816–34.

78.

Ferrara

Gerber

LeCouter

. The biology of VEGF and its receptors. Nat Med. 2003;9(6):669–76.

79.

Benjamin

Keshet

. Conditional switching of vascular endothelial growth factor (VEGF) expression in tumors: induction of endothelial cell shedding and regression of hemangioblastoma-like vessels by VEGF withdrawal. Proc Natl Acad Sci U S A. 1997;94(16):8761–66.

80.

Chen

Yang

Zhang

Zuo

Wang

Chen

. C-reactive protein can upregulate VEGF expression to promote ADSC-induced angiogenesis by activating HIF-1α via CD64/PI3k/Akt and MAPK/ERK signaling pathways. Stem Cell Res Ther. 2016;7(1):114.

81.

Lee

Park

Kim

Chung

Kim

Park

Kim

. Dll4 inhibition promotes graft retention in fat grafting enriched with adipose-derived stem cells. Stem Cells Transl Med. 2022;11(7):742–52.

82.

Jie

Nie

Zhu

Jiang

Liu

. Effects of miR126 expressing adipose-derived stem cells on fat graft survival and angiogenesis. Aesthetic Plast Surg. 2023;47(2):825–32.

83.

Arderiu

Peña

Aledo

Juan-Babot

Crespo

Vilahur

Oñate

Moscatiello

Badimon

. MicroRNA-145 regulates the differentiation of adipose stem cells toward microvascular endothelial cells and promotes angiogenesis. Circ Res. 2019;125(1):74–89.

84.

Shang

Zhang

Cui

Guo

. Hypoxia promotes differentiation of adipose-derived stem cells into endothelial cells through demethylation of ephrinB2. Stem Cell Res Ther. 2019;10(1):133.

85.

Chen

Yan

Guo

Chen

Huang

Zhang

Chen

. Adipose-derived mesenchymal stem cells promote the survival of fat grafts via crosstalk between the Nrf2 and TLR4 pathways. Cell Death Dis. 2016;7(9):e2369.

86.

Souza

LEB

Beckenkamp

Sobral

Fantacini

DMC

Melo

FUF

Borges

Leopoldino

Kashima

Covas

. Pre-culture in endothelial growth medium enhances the angiogenic properties of adipose-derived stem/stromal cells. Angiogenesis. 2018;21(1):15–22.

87.

Yuan

Gao

Liu

. Role of adipose-derived stem cells in enhancing angiogenesis early after aspirated fat transplantation: induction or differentiation? Cell Biol Int. 2013;37(6):547–50.

88.

Hong

Yim

Kim

Jin

Lim

Chang

Minn

. The fate of the adipose-derived stromal cells during angiogenesis and adipogenesis after cell-assisted lipotransfer. Plast Reconstr Surg. 2018;141(2):365–75.

89.

AlZaim

de Rooij

LPMH

Sheikh

Börgeson

Kalucka

. The evolving functions of the vasculature in regulating adipose tissue biology in health and obesity. Nat Rev Endocrinol. 2023;19(12):691–707.

90.

Miranville

Heeschen

Sengenès

Curat

Busse

Bouloumié

. Improvement of postnatal neovascularization by human adipose tissue-derived stem cells. Circulation. 2004;110(3):349–55.

91.

Hong

Kim

Park

Jin

Chang

. Systemic administration of adipose-derived stromal cells concurrent with fat grafting. Plast Reconstr Surg. 2019;143(5):973e–82.

92.

Huang

Jin

Hong

Chang

. Enhanced effect of secondary administration of adipose-derived stromal cells concurrent with fat grafting. Plast Reconstr Surg. 2024;153(2):390–99.

93.

Yang

Zhu

Yin

Zhao

Wang

Dang

Zhang

Wen

. Integration of human umbilical cord mesenchymal stem cells-derived exosomes with hydroxyapatite-embedded hyaluronic acid-alginate hydrogel for bone regeneration. ACS Biomater Sci Eng. 2020;6(3):1590–1602.

94.

Liu

Jia

Liu

. Exosomes derived from atorvastatin-pretreated MSC accelerate diabetic wound repair by enhancing angiogenesis via AKT/eNOS pathway. Stem Cell Res Ther. 2020;11(1):350.

95.

Wei

Wang

Cao

Liu

Zheng

Tian

. Mesenchymal stem cell-derived exosomes: a promising biological tool in nanomedicine. Front Pharmacol. 2020;11:590470.

96.

Chen

Cai

Wei

Jiang

Desjardins

Adams

Kao

Guo

. Exosomes are comparable to source adipose stem cells in fat graft retention with up-regulating early inflammation and angiogenesis. Plast Reconstr Surg. 2019;144(5):816e–27e.

97.

Chen

Xiong

Xue

Wang

. Adipose-derived stem cells exosomes improve fat graft survival by promoting prolipogenetic abilities through Wnt/β-catenin pathway. Stem Cells Int. 2022;2022:5014895.

98.

Han

Bai

Yan

Ren

Zeng

Pei

Han

. Co-transplantation of exosomes derived from hypoxia-preconditioned adipose mesenchymal stem cells promotes neovascularization and graft survival in fat grafting. Biochem Biophys Res Commun. 2018;497(1):305–12.

99.

Yang

Cai

Wang

Lin

. Pluronic F-127 hydrogel loaded with human adipose-derived stem cell-derived exosomes improve fat graft survival via HIF-1α-mediated enhancement of angiogenesis. Int J Nanomedicine. 2023;18:6781–96.

100.

Ruel-Gariépy

Leroux

. In situ-forming hydrogels–review of temperature-sensitive systems. Eur J Pharm Biopharm. 2004;58(2):409–26.

101.

Zhu

Zhang

Wang

. Supplementation with extracellular vesicles derived from adipose-derived stem cells increases fat graft survival and browning in mice: a cell-free approach to construct beige fat from white fat grafting. Plast Reconstr Surg. 2020;145(5):1183–95.

102.

Mantovani

Biswas

Galdiero

Sica

Locati

. Macrophage plasticity and polarization in tissue repair and remodelling. J Pathol. 2013;229(2):176–85.

103.

Murray

Wynn

. Protective and pathogenic functions of macrophage subsets. Nat Rev Immunol. 2011;11(11):723–37.

104.

Wolfrum

Gerhart-Hines

. Fueling the fire of adipose thermogenesis. Science. 2022;375(6586):1229–31.

105.

Harms

Seale

. Brown and beige fat: development, function and therapeutic potential. Nat Med. 2013;19(10):1252–63.

106.

Stanford

Middelbeek

Goodyear

. Exercise effects on white adipose tissue: beiging and metabolic adaptations. Diabetes. 2015;64(7):2361–68.

107.

Zhang

Cai

Zhang

Zeng

Fan

Zou

Fang

Lin

, et al Brown adipose tissue transplantation ameliorates diabetic nephropathy through the miR-30b pathway by targeting Runx1. Metabolism. 2021;125:154916.

108.

Stanford

Middelbeek

Townsend

Nygaard

Hitchcox

Markan

Nakano

Hirshman

Tseng

Goodyear

. Brown adipose tissue regulates glucose homeostasis and insulin sensitivity. J Clin Invest. 2013;123(1):215–23.

109.

Quan

Cai

Liao

Zhang

. Transplantation of beige adipose organoids fabricated using adipose acellular matrix hydrogel improves metabolic dysfunction in high-fat diet-induced obesity and type 2 diabetes mice. J Cell Physiol. 2024;239:e31191.

110.

Quan

Zhang

. Use of brown adipose tissue transplantation and engineering as a thermogenic therapy in obesity and metabolic disease. Obes Rev. 2023;25:e13677.

111.

Villarroya

Cereijo

Villarroya

Giralt

. Brown adipose tissue as a secretory organ. Nat Rev Endocrinol. 2017;13(1):26–35.

112.

Zhang

Qin

Wang

Liu

Zhu

Zeng

Zhang

Long

. Adipose-derived stromal cells attenuate adipose inflammation in obesity through adipocyte browning and polarization of M2 macrophages. Mediators Inflamm. 2019;2019:1731540.

113.

Adachi

Ueda

Nomura

Ito

Katoh

Katagiri

Yamada

Hashimoto

Zhai

Numata

Otani

, et al Beiging of perivascular adipose tissue regulates its inflammation and vascular remodeling. Nat Commun. 2022;13(1):5117.

114.

Koseki

Suzuki

. Deformation dynamics of giant unilamellar vesicles in the large surface-to-volume ratio regime: the emergence of neuron-like morphology. Langmuir. 2020;36(22):6238–44.

115.

Jiang

Lin

Zhang

Liao

Cai

. The fates of different types of adipose tissue after transplantation in mice. FASEB J. 2022;36(9):e22510.

116.

Luo

Cheng

Yuan

Hao

Liu

Zeng

Pan

. Transplantation of cold-stimulated subcutaneous adipose tissue improves fat retention and recipient metabolism. Aesthet Surg J. 2024;44(7):NP486-NP500.

117.

Park

Kim

Sun

Scherer

. VEGF-A–expressing adipose tissue shows rapid beiging and enhanced survival after transplantation and confers IL-4–independent metabolic improvements. Diabetes. 2017;66(6):1479–90.

118.

Kotzbeck

Giordano

Mondini

Murano

Severi

Venema

Cecchini

Kershaw

Barbatelli

Haemmerle

Zechner

, et al Brown adipose tissue whitening leads to brown adipocyte death and adipose tissue inflammation. J Lipid Res. 2018;59(5):784–94.

119.

Lin

Zhu

Liao

Liang

Quan

Cai

. Spontaneous browning of white adipose tissue improves angiogenesis and reduces macrophage infiltration after fat grafting in mice. Front Cell Dev Biol. 2022;10:845158.

120.

Zheng

Bao

Jia

Cao

. Brown adipose tissue promotes autologous fat grafts retention possibly through inhibiting Wnt/β-catenin pathway. Aesthetic Plast Surg. 2024;48(9):1817–24.

121.

Sakers

De Siqueira

Seale

Villanueva

. Adipose-tissue plasticity in health and disease. Cell. 2022;185(3):419–46.

122.

Ghaben

Scherer

. Adipogenesis and metabolic health. Nat Rev Mol Cell Biol. 2019;20(4):242–58.

123.

Sugihara

Yonemitsu

Miyabara

Toda

. Proliferation of unilocular fat cells in the primary culture. J Lipid Res. 1987;28(9):1038–45.

124.

Sugihara

Yonemitsu

Miyabara

Yun

. Primary cultures of unilocular fat cells: characteristics of growth in vitro and changes in differentiation properties. Differentiation. 1986;31(1):42–49.

125.

Matsumoto

Kano

Kondo

Fukuda

Iribe

Tanaka

Matsubara

Sakuma

Satomi

Otaki

Ryu

, et al Mature adipocyte-derived dedifferentiated fat cells exhibit multilineage potential. J Cell Physiol. 2008;215(1):210–22.

126.

Liang

Wang

Tang

Cai

Liao

. Ceiling culture of human mature white adipocytes with a browning agent: a novel approach to induce transdifferentiation into beige adipocytes. Front Bioeng Biotechnol. 2022;10:905194.

127.

Zhu

Zhao

Chai

Jia

. Evaluating the efficacy of dedifferentiated fat cells (DFATs) vs adipose-derived stem cells (ASCs) in enhancing the viability of fat grafts. Aesthet Surg J. 2023.

128.

Watanabe

Goto

Kato

Komiyama

Nagaoka

Kazama

Yamamoto

Konuma

Hagikura

Matsumoto

. The neovascularization effect of dedifferentiated fat cells. Sci Rep. 2020;10(1):9211.

129.

Chai

Chen

Yin

Zhang

Han

Cai

Yin

. Dedifferentiation of human adipocytes after fat transplantation. Aesthet Surg J. 2022;42(6):NP423–31.

130.

Jumabay

Abdmaulen

Urs

Heydarkhan-Hagvall

Chazenbalk

Jordan

Roos

Yao

Boström

. Endothelial differentiation in multipotent cells derived from mouse and human white mature adipocytes. J Mol Cell Cardiol. 2012;53(6):790–800.

131.

Liang

Tang

Shi

Cai

Liao

. Physical expansion preconditioning promotes host-derived adipocyte dedifferentiation and migration into fat grafts in A murine model. Plast Reconstr Surg. 2023.

132.

Chan

McCulley

Macmillan

. Autologous fat transfer: a review of the literature with a focus on breast cancer surgery. J Plast Reconstr Aesthet Surg. 2008;61(12):1438–48.

133.

Ellenbogen

. Autologous fat injection. Plast Reconstr Surg. 1991;88(3):543–44.

134.

Zhao

Lee

Sasagawa

Sokol

Wang

Ransom

Zhao

Steininger

Koepke

Borrelli

, et al A combination of distinct vascular stem/progenitor cells for neovascularization and ischemic rescue. Arterioscler Thromb Vasc Biol. 2023;43(7):1262–77.

135.

Borrelli

Patel

Blackshear

Vistnes

Diaz Deleon

Adem

Shen

Sokol

Momeni

Nguyen

Longaker

, et al CD34+CD146+ adipose-derived stromal cells enhance engraftment of transplanted fat. Stem Cells Transl Med. 2020;9(11):1389–1400.

136.

Chan

Lindau

Jiang

Chen

Zhang

Chen

Seita

Sahoo

Kim

Lee

Park

, et al Clonal precursor of bone, cartilage, and hematopoietic niche stromal cells. Proc Natl Acad Sci U S A. 2013;110(31):12643–48.

137.

Wang

Dai

Wang

. Adult stem cells in vascular remodeling. Theranostics. 2018;8(3):815–29.

138.

Yoder

. Endothelial stem and progenitor cells (stem cells): (2017 Grover Conference Series). Pulm Circ. 2018;8(1):2045893217743950.

139.

Fang

Dai

Kurjiaka

Burt

Hirschi

. Connexin45 regulates endothelial-induced mesenchymal cell differentiation toward a mural cell phenotype. Arterioscler Thromb Vasc Biol. 2013;33(2):362–68.

140.

Papalexi

Satija

. Single-cell RNA sequencing to explore immune cell heterogeneity. Nat Rev Immunol. 2018;18(1):35–45.

141.

Kalisky

Oriel

Bar-Lev

Ben-Haim

Trink

Wineberg

Kanter

Gilad

Pyne

. A brief review of single-cell transcriptomic technologies. Brief Funct Genomics. 2018;17(1):64–76.

142.

Trapnell

. Defining cell types and states with single-cell genomics. Genome Res. 2015;25(10):1491–98.

143.

Burl

Ramseyer

Rondini

Pique-Regi

Lee

Granneman

. Deconstructing adipogenesis induced by β3-adrenergic receptor activation with single-cell expression profiling. Cell Metab. 2018;28(2):300–3094.

144.

AlJanahi

Danielsen

Dunbar

. An introduction to the analysis of single-cell RNA-sequencing data. Mol Ther Methods Clin Dev. 2018;10:189–96.

145.

Rondini

Granneman

. Single cell approaches to address adipose tissue stromal cell heterogeneity. Biochem J. 2020;477(3):583–600.

146.

Massier

Jalkanen

Elmastas

Zhong

Wang

Nono Nankam

Frendo-Cumbo

Bäckdahl

Subramanian

Sekine

Kerr

, et al An integrated single cell and spatial transcriptomic map of human white adipose tissue. Nat Commun. 2023;14(1):1438.

147.

Zhang

Lin

Ding

Yang

Cai

Wang

, et al Three-dimensional spheroid formation of adipose-derived stem cells improves the survival of fat transplantation by enhance their therapeutic effect. Biotechnol J. 2023;18(10):e2300021.

The Evolving Function of Vasculature and Pro-angiogenic Therapy in Fat Grafting

Abstract

Keywords

Introduction

Vasculature Function After Fat Grafting

Delivering Essential Oxygen Crucial and Nutrient-Waste Exchange for Adipocyte Survival

Supplying Circulating Stem Cells and Inflammatory Cells From Circulation to Adipose Tissue Graft

Providing NICHE for Adipogenesis and Angiogenesis in Grafts

Cell-Enrichment Therapy in Fat Grafting

Endothelial Progenitor Cell

Macrophage

ASCs’ Function in Cell-Assisted Lipotransfer (CAL)

Adipocytes’ Pro-angiogenic Function in Fat Grafting

Brown and Beige Adipocytes

Dedifferentiated Adipocytes (DAs)

Future Perspective of Vascularization in Fat Grafting

Conclusion

Footnotes

Acknowledgements

Authors’ Note

Author Contributions

Availability of data and material

Ethical Approval

Statement of Human and Animal Rights

Statement of Informed Consent

Declaration of Conflicting Interests

Funding

ORCID iDs

References