Matrix-Assisted Transplantation of Functional Beige Adipose Tissue

The unabated growth of the obesity epidemic and associated diseases such as type 2 diabetes reflects the current lack of effective strategies for intervention and treatment. Since obesity results from an imbalance in the ratio of energy intake to energy expenditure, it can be treated with reduced caloric uptake and/or increasing energy expenditure. Brown adipose tissue (BAT) and inducible brown-like cells in white adipose tissue (WAT), currently referred to as beige or brite adipocytes (1), possess the innate ability to dissipate metabolic energy as heat through nonshivering thermogenesis. This is possible owing to the presence of the uncoupling protein (UCP) 1 (2), a long-chain fatty acid anion/proton symporter (3), in the inner mitochondrial matrix, which allows the return of protons after they have been pumped across the mitochondrial inner membrane by the electron transport chain, thereby bypassing ATP synthase. While the importance of BAT for thermogenesis in smaller mammals and human infants has been well documented (4), only recent observations have demonstrated the presence of BAT in adult humans (5,6). These observations have reversed the dogma that BAT is absent in adult humans and have presented a new route for the treatment of obesity-related disorders. Over 90% of the metabolic energy consumed by activated BAT will be derived from the β-oxidation of free fatty acids (FFAs) (7). Furthermore, many studies have suggested that the amount of BAT correlates inversely with BMI, raising the possibility that variations in the amount of BAT may drive alterations in body weight (8). Thus, increasing BAT mass may serve as a novel approach to combat obesity and related disorders such as type 2 diabetes. This concept is supported by recent studies that found evidence for metabolic enhancement, including the reversal of type 1 diabetes (9) and resistance to diet-induced obesity, in mouse models of BAT expansion (10) via transplantation of existing BATs.

Strategies for expanding BAT can be grouped into two general categories: pharmaceutical/genetic intervention to trigger endogenous BAT/beige differentiation pathways and the ex vivo generation of BAT for implantation (11–15). Gene therapy approaches, for example, the ectopic overexpression of the transcriptional regulator PRDM16 (14,16), are powerful tools to investigate BAT biology but are difficult to apply clinically owing to the risks associated with current gene therapy regiments (17). Pharmacological activation of differentiation pathways that drive a WAT-to-BAT transition (“browning”) run the risk of affecting the function of other tissues and offer little control over the location and temporal extent of BAT expansion. Similarly, heterologous transplantation of existing BAT into immune-compromised recipients has demonstrated the metabolic impact of BAT expansion (10), but this approach has no clinical translatability owing to a paucity of donor tissues and the expected host-versus-graft rejection. Thus, we propose an alternative approach that takes advantage of current progress in the field of bio-inspired hyaluronic acid (HyA) hydrogels to engineer a matrix-assisted cell transplantation (MACT) system for beige adipose tissue (BAT-MACT) derived from the readily available, multipotent stem cell (MSC)-containing, stromal vascular fraction of WAT (18).

Experiments were performed according to Association for Assessment and Accreditation of Laboratory Animal Care guidelines and approved by the University of California, Berkeley, Animal Care and Use Committee. Eight-week-old male C57BL/6J or FVB/NJ mice (The Jackson Laboratory) were group housed for a 1-week acclimation period (temperature 21 ± 2°C, humidity 30–70%) in Tecniplast cages filled with sani-chip bedding (Harlan) and then individually housed 1 week prior to implantation and administration of high-fat (60%) diet (Teklad TD.06414). Luciferase-expressing L2G85 (FVB/NJ) mice were used as a cell source for experiments using bioluminescent monitoring of implant viability and function.

Fat depots from L2G85 (FVB/NJ) or C57BL/6J mice, perigonadal (visceral) or inguinal (subcutaneous), respectively, were isolated from 3-month-old mice as previously described (19).

Adipose-derived MSCs (ADMSCs) (initial density ∼100 k cells/cm²) were cultured to postconfluence on tissue culture polystyrene (TCPS) or adsorbed peptide surfaces. Differentiation was induced for 3 days with 0.85 μmol/L insulin, 10 nmol/L triiodothyronine, 1 μmol/L dexamethasone, and 500 μmol/L isobutylmethylxanthine in high-glucose DMEM (Invitrogen) with 10% FCS (Gibco) and penicillin/streptomycin (P/S) (Gibco). The induced cells were then treated with 0.85 μmol/L insulin and 100 nmol/L rosiglitizone in high-glucose DMEM (Invitrogen) with 10% FCS and P/S for 12 subsequent days. Before harvest with TRIzol, cultures were treated with 5 μg/mL CL-316243 for 4 h. The initial peptide screening differentiation experiments were carried with the goal of determining the differentiation outcome of an adhesion-selected cell fraction, a process referred to as “panning.” To this end, primary ADMSCs were given 4 h to adhere onto the surface coating, cultures were then PBS washed twice to remove less adherent cells within the ADMSC population, and differentiation was induced as described above (Fig. 1A–C). Experiments to determine optimal proportions of selected peptides were carried out without washing off poorly adherent cells (Fig. 1D–F).

Immunohistochemistry was performed as previously described (20). Whole in vitro hydrogels were fixed with 4% paraformaldehyde for 30 min at room temperature and imaged within their MilliCell insert (Milipore) placed on coverslips.

Tissue samples were suspended in RNAlater ICE (Ambion) and stored at −20°C. mRNA was isolated from RNAlater ICE stabilized tissues or directly from in vitro cultures of monolayers or cell-laden hydrogels with TRIzol reagent (Ambion). Tissue samples were homogenized with a Polytron PT 2100. Assays were carried out on an ABI 7500 RT-PCR system with TaqMan Universal Master Mix II and validated PrimeTime primer probe sets (Integrated DNA Technologies). A first-strand cDNA synthesis kit (Fisher) was used to transcribe 5 μg RNA/20 μL. cDNA (100 ng) was used per RT-PCR reaction in triplicates. The Δ-Δ-CT method was used to comparatively assess mRNA quantity. All data are represented as a sample’s value normalized to GAPDH relative to source tissue–derived ADMSCs, source tissue, or endogenous BAT.

A Pierce BCA kit was used to measure protein concentrations. Western blotting was accomplished with Novex 4–20% tris-glycine gels, iBlot Transfer Stacks (Invitrogen), and a LI-COR Odyssey infrared imager with anti-UCP1 (Sigma-Aldrich) and anti–β-tubulin E7 clone (Developmental Studies Hybridoma Bank). Data are presented as UCP1 normalized to β-tubulin relative to endogenous BAT.

Modified HyA (500 kDa) was synthesized using previously established methods (21). The acrylated HyA (AcHyA)-C16 or AcHyA-Ag73 was synthesized by reacting peptides (CGGKAFDITYVRLKF [10 mg] or CGGRKRLQVQLSIRT [10 mg]) with AcHyA (25 mg, 10 mL deionized water) at room temperature. To synthesize hydrogels, we dissolved AcHyA (2 mg), AcHyA-C16 (6 mg), and AcHyA-Ag73 (2 mg) in 0.3 mL triethanolamine buffer (0.3 mol/L, pH 8), incubated for 30 min at 37°C, and subsequently matrix metalloprotease (MMP-13)–cleavable peptide (CQPQGLAKC) (3 mg in 50 µL triethanolamine buffer) was added to HyA peptide solution. The crosslinker concentration was varied to achieve complete crosslinking of available acrylate functionalities on the HyA macromers (∼200 mmol/L). The initial storage modulus of all hydrogels (in vitro and in vivo) was measured consistently to be ∼850 Pa.

ADMSCs were cultured in vitro in DMEM with 10% FBS and 1% P/S on TCPS. Two days postconfluency, differentiation was induced with 0.85 μmol/L insulin, 10 nmol/L triiodothyronine, 1 μmol/L dexamethasone, and 500 μmol/L isobutylmethylxanthine (for white adipocytes scaffolded by unmodified AcHyA [WAT-MACT], triiodothyronine was omitted from the cocktail). Three million cells per mL were suspended in AcHyA-Ag73:C16 (1:3 molar ratio) just before implantation (for WAT-MACT, cells were suspended in AcHyA). The cell suspension was mixed with the MMP-13–cleavable crosslinker at a final concentration of ∼200 mmol/L (sufficient to achieve a modulus of ∼850 Pa) and 100 μL of the forming BAT-MACT was immediately injected to recipient mice.

Experiments where bioluminescent imaging (BLI) was used to monitor the viability and function of the BAT-MACTs, in conjunction with metabolic measures, were performed in FVB/NJ mice with ADMSCs isolated from perigonadal fat depots of syngenic L2G85 donor mice. Experiments where metabolic parameters were assessed without BLI monitoring were performed with C57BL/6J mice implanted with BAT-MACTs generated with ADMSCs isolated from the inguinal adipose depot.

ADMSCs were cultured in vitro in DMEM with 10% FBS and 1% P/S on TCPS. Three million subconfluent cells per milliliter were suspended in the specific hydrogel and injected into a Millicell transwell chamber (Millipore) in a 24-well tissue culture plate (Falcon). After 2 days of culture in DMEM with 10% FBS and 1% P/S, the differentiation protocol described for the in vitro differentiation of ADMSCs was implemented.

Imaging was accomplished with an IVIS Spectrum. Luciferin (45 μL; 4 mg/mL i.p.) in PBS was injected into animals. FFA-SS-Luc 100 μL of (1.4 μg/μL) intraperitoneal compound was injected into mice. Once the BLI signal dropped off (∼40 min), the animals were removed to recover. Four hours later, once all original BLI signal was extinguished, 1 μg CL-316243/g body wt i.p. was injected followed by another dose of the FFA-SS-Luc.

Respirometry was performed with the Comprehensive Lab Animal Monitoring System (CLAMS) (Columbus Instruments). Measurements were taken over 24-h periods, and average values were assessed (22). CL-316243 was administered to animals, and the 4-h preinjection was compared with the 4-h postinjection time periods. Activity was monitored in 1-min intervals of infrared beam breaks in x-, y-, and z-axes and was found to be not significantly different for any of the groups.

A Bio-Rad Bio-Plex Pro Mouse Cytokine 23-Plex immunoassay kit was used per the manufacturer’s instructions with a Bio-Rad MAGPIX system.

Viscoelastic properties of the hydrogel were determined using an oscillatory rheometer with parallel-plate geometry (8 mm) and a gap height of 0.2 mm under 10% constant strain and frequency ranging from 0.1 Hz to 10 Hz at 37°C in a humidity-controlled chamber.

Purified peptides were suspended at 20 μmol/L in ultrapure water with 1% P/S. Peptide suspensions (20 μmol/L) were allowed to adsorb to the 24-well TCPS plates (BD Falcon) overnight at 4°C. Plates were then washed two times with PBS. Twenty thousand primary ADMSCs suspended in DMEM with 10% FBS and 1% P/S were added to each well and incubated at 37°C for 4 h. Media and nonadherent cells were removed and centrifuged to separate the cells from media. The adherent and nonadherent cells were then counted with the CyQUANT kit (Invitrogen).

A Physitemp BAT-12 Microprobe thermometer was used to measure core body temperature of mice rectally. Cold challenges took place in a 4°C walk-in refrigerator approved for use in accordance with Association for Assessment and Accreditation of Laboratory Animal Care guidelines.

Blood glucose was measured with a Nova Max blood glucometer and Nova Max glucose test strips (Sanvita). Mice were injected with an equivalent dose of glucose (167 μL of 300 g/L glucose). This dose was selected as a dose that the median mouse weight would warrant to achieve the standard glucose dose of 2 mg/g body wt generally used for a glucose tolerance test.

All data are presented as SEM analyzed using Prism (GraphPad). Statistical significance was determined by either one-way ANOVA followed by Tukey posttest or unpaired two-tailed Student t test. Significance presented at P < 0.05, P < 0.01, and P < 0.001.

Cell adhesion and matrix interactions exert significant control over MSC differentiation (23); however, no screens have examined how these interactions affect the browning of WAT-derived MSCs (ADMSCs). To this end, ADMSCs were extracted from WAT (19) and cultured on TCPS plates coated with specific peptides found to be ligands for integrins and syndecans (Fig. 1A and Supplementary Table 1). Differentiation was induced, and lipid droplet accumulation (Fig. 1B) and UCP1 expression were assessed (Fig. 1C). Our initial screens identified non-RGD peptide sequences such as AG73 (CGGRKRLQVQLSIRT), an adhesion ligand for α₆ integrins and syndecan-1 (24,25), as particularly efficient in driving UCP1 expression of ADMSCs, while C16 (CGGKAFDITYVRLKF), an α_5/vβ_1/3 integrin–engaging ligand (26), induced lipid accumulation. Interestingly, while all peptide-coated substrates outperformed uncoated TCPS in UCP1 expression assays, a combination of AG73 with C16 (1:3 molar ratio) had significant effects on UCP1 expression (Fig. 1D and E). Combining varied concentrations of AG73 with AG32 (CGGTWYKIAFQRNRK), another α₆β₁ integrin–engaging ligand (25), we observed attenuated UCP1 expression (Fig. 1F). The impressive enhancement of UCP1 expression achieved by optimizing adhesion ligands is highlighted by the fact that previous attempts to differentiate WAT-derived MSCs into BAT yielded UCP1 levels significantly lower than BAT-derived MSCs (11). In contrast, we find UCP1 expression in WAT-derived MSCs differentiated on optimized adhesion ligands to be comparable with those of differentiated BAT-derived MSCs in vitro (Supplementary Fig. 1).

Engagement with the extracellular matrix provides a crucial factor in driving MSC differentiation through biophysical properties and composition (27–29). Using our expertise in biomaterials (20,30–32), we synthesized AcHyA macromers (Fig. 2A) conjugated with the bioactive peptides. These functionalized AcHyA macromers (AcHyA-Ag73 and AcHyA-C16) were combined with specific MMP-sensitive peptide crosslinkers at defined concentrations to form hydrogels with precise physical characteristics and degradation kinetics (Fig. 2B). The synthesis of hydrogel networks takes advantage of the Michael-type addition reaction (i.e., attack of the acrylate group by available cysteine thiols to form a covalent bond) (33,34), which forms a defined hydrogel scaffold within 5 min (Fig. 2C and D).

We verified that the ADMSCs embedded in the optimized matrix were forming functional cell adhesions with the hydrogel. By suspending ADMSCs in uncrosslinked AcHyA-Ag73:C16 or unmodified AcHyA, we found that the inclusion of adhesion peptides resulted in a significant increase in the storage modulus of uncrosslinked hydrogels seeded with ADMSCs, indicating that the adhesion peptides enable cells to mechanically integrate the synthetic extracellular matrix (Fig. 2E). The optimized hydrogel scaffold allowed the ADMSCs to evenly distribute and differentiate into UCP1-positive adipocytes in vitro without direct cell-cell contact (Fig. 2F and Supplementary Fig. 2). Importantly, culturing cells in AcHyA-Ag73:C16 hydrogels significantly improved UCP1 expression of differentiated ADMSCs in vitro compared with monolayers exposed to adsorbed Ag73:C16 (Fig. 2G). Notably, unconjugated hyaluronan- or RGD (CGGNGEPRGDTYRAY)-conjugated hydrogels were not capable of inducing UCP1 expression independent of Ag73 and C16 (Fig. 2H).

To produce a functional BAT-MACT, we isolated ADMSCs from donor WAT, initiated differentiation in vitro, and suspended the cells in Ag73 and C16 presenting AcHyA macromers. BAT-MACTs were formed by mixing cells (3*10⁶/mL) with AcHyA macromers (3 wt%) and a MMP-13–sensitive crosslinker. The cell suspension was immediately injected with a 22.5-gauge needle into the subcutaneous inguinal fat pad (or as noted) of recipient animals. The viability of the implanted BAT-MACT was monitored via bioluminescence using luciferase-expressing donor cells delivered in a hydrogel or suspended in PBS. Subcutaneous injections of luciferin adjacent to the implant demonstrate luminescence for both conditions after implantation. Importantly, the PBS-delivered ADMSCs produce little luminescence when the bioluminescent substrate is introduced into general circulation. By delivering luciferin with intraperitoneal injections distant from the BAT-MACT implant site, we observed massive signal expansion during the first week, which declines to undetectable levels 4 weeks postimplantation (Fig. 3A).

Owing to the rapid crosslinking kinetics, BAT-MACTs form a distinct subcutaneous lobe upon injection that is clearly brown/beige in appearance (Fig. 3B). BAT-MACTs were highly vascularized on a macroscopic level, and thin sections of implants demonstrated an extensive and consistent vascular network throughout the implants (Fig. 3C). Likewise, UCP1-positive and lipid droplet–accumulating cells were evenly spread throughout the BAT-MACT (Fig. 3D). Interestingly, regions of the implant showed positive staining for tyrosine hydroxylase, the rate-limiting enzyme in the synthesis of catecholamines (Supplementary Fig. 3). Tyrosine hydroxylase is essential for the production of norepinephrine by sympathetic neurons and macrophages to not only activate respiration but also maintain differentiation status and function (35). UCP1 mRNA expression of the implanted MACT was >10-fold higher than that of comparable in vitro cultured MACTs (Fig. 3E) and reached 7–17% of the levels found in endogenous BAT (Supplementary Fig. 4). Importantly, PRDM16 and PPARγ expression in BAT-MACTs were comparable with those in endogenous BAT (Fig. 3F), highlighting the potential of the MACT.

Immense differences of UCP1 induction within white fat depots have been shown to be under genetic control (36). We confirmed these differences by generating BAT-MACTs from FVB/NJ and C57BL/6J mice. Similar to previously published findings, C57BL/6J visceral fat depots produce significantly less UCP1 protein than FVB/NJ or subcutaneously sourced C57BL/6J ADMSCs within our BAT-MACT system (Fig. 3G). To assess the effect of implant location, we analyzed two different implant sites in the thoracic and inguinal regions for lipid droplets and UCP1 expression. In BAT-MACTs from both implantation sites, we observed multilocular lipid droplets, UCP1-positive cells, and blood vessels with no apparent difference between the implantation sites. Impressively, UCP1 protein levels in both BAT-MACT sites reached ∼30% of BAT from the same animals (Fig. 3H). Importantly, inflammatory cytokines measured in serum were not upregulated whatsoever by the BAT-MACT system; in fact, interleukin-2 was dissipated (Fig. 3I). Therefore, the BAT-MACT system allows for the measurement of metabolic effects independent of inflammatory artifacts.

To determine the metabolic impact of BAT-MACTs, we examined the respiratory capacity of animals augmented with a BAT-MACT or WAT-MACT. HyA matrixes have been shown to aid in the development and persistence of WAT (37,38), and WAT-MACTs formed fatty implants that were well vascularized, contained extensive lipid stores, and had undetectable UCP1 expression (Supplementary Fig. 5). The BAT-MACT implanted animals displayed significantly enhanced VO₂ and reduced respiratory exchange ratio, recapitulating the elevated fatty acid utilization and respiration after classical BAT activation (Fig. 4A and B). Furthermore, BAT-MACT augmented animals were significantly more responsive to CL-316243, a β₃-specific adrenergic agonist (Fig. 4C). To verify that FFAs would be taken up by BAT-MACTs, we used a recently developed optical probe for fatty acid uptake (39) and observed that an intraperitoneal injection of the probe was robustly taken up by BAT-MACTs. Importantly, as previously shown for endogenous BAT (40), injection of a β₃-adrenergic agonist significantly enhanced fatty acid uptake rates by the BAT-MACTs (Fig. 4D).

To test the impact of BAT-MACTs on thermogenesis, we examined core body temperature at room temperature (21°C) and over 24 h of 4°C. Body temperatures of BAT-MACT recipients were higher throughout the cold challenge as well as at room temperature (Fig. 4E). Thermography revealed that the implant is distinctly thermogenic (Supplementary Fig. 6). Importantly, this thermogenic effect was dependent on the amount of cells implanted (Fig. 4F) and the environmental temperature (Fig. 4G). Using C57BL/6J mice as a model for diet-induced obesity, we measured the impact of BAT-MACT impact on weight gain and glucose homeostasis. Animals implanted with BAT-MACTs and housed at 21°C for 2 weeks gain significantly less weight when fed a 60% fat diet compared with PBS-delivered ADMSCs, cell-free matrix, or WAT-MACTs (Fig. 5A) in spite of slightly higher food consumption (Fig. 5B). Extended monitoring of weight gain identified that the reduction of weight gain was significant within 2 weeks post–BAT-MACT implantation and concomitant high-fat feeding. The difference in weight gain was sustained over a time period coinciding with the lifetime of the BAT-MACT cells as measured by bioluminescence (Fig. 5C).

After 2 weeks, serum triglyceride levels were found to trend lower and serum free fatty acids were reduced in animals augmented with BAT-MACTs (Fig. 5D and Supplementary Fig. 7). Accompanying the reduced weight gain and serum FFAs, resting blood glucose levels (Fig. 5E) and glucose tolerance (Fig. 5F) were improved in BAT-MACTs but not WAT-MACTs, cell-free implants, or matrix-free implants of BAT-like cells.

Expansion of UCP1-positive tissue holds exciting potential to combat obesity and related disorders (41). However, few practicable approaches for clinical deployment have been presented thus far. This novel strategy to integrate WAT-derived MSCs with bioactive hydrogels bespoke to enhance differentiation, support tissue formation, and survival upon implantation provides the required core technology for autologous expansion of BAT mass in patients. ADMSCs are easily extractable and abundant and are already used in regenerative medicine (18). ADMSCs used in this study were only partially purified; further gains in differentiation effectiveness could be anticipated by sorting for UCP1-enriching markers such as Sca-1 (13), CD24, CD29 (15,42), and CD137 (43).

Hydrogel-based therapies are minimally invasive therapeutic options that will complement pharmacological and cell-based approaches (44). We selected HyA as the core scaffold of our matrix for its biocompatibility, biodegradability, and nonimmunogenic properties and its role in tissue development and repair (30). Recently, HyA has been shown to augment integrin-mediated mechanotransductive signaling (45) and likely plays an essential role in the overall effect that our matrix has on the differentiation of ADMSCs. Similar hydrogel systems have been proposed for clinical use that used crosslinkers and adhesion ligands that directly compete for attachment to the available functional groups on core polymeric chains within the hydrogel. These systems have struggled with compromised crosslinking efficiencies, unbound adhesion ligands preventing cell association with the matrix, and low–molecular weight core structures that degrade too rapidly to be useful (46).

Matrix stiffness plays an important role in cell fate determination and should be a critical design feature of cell transplantation systems (27). We have developed a modular hydrogel system that allows for modulus and biological properties of the hydrogel to be tuned independently. This modular approach to tissue engineering may be essential to target an optimal net modulus for the development and maintenance of our BAT-MACT by allowing us to integrate and independently tune the crosslinking modulus and modulus that cells produce through ligand-specific mechanotransduction observed in Fig. 2E.

We have already shown that some ADMSCs spontaneously differentiate into UCP1-expressing cells in vitro using our bio-inspired hydrogel (Fig. 2F). In our future studies, generation of UCP1-positive adipocytes could be enhanced with matrices presenting heparin-binding growth factors, specifically, BMP7 and FGF21, which have been implicated in BAT differentiation (47). This objective is feasible, as heparin-binding growth factors can be associated with the matrix through the addition of thiolated heparin to the AcHyA (20). Ultimately, there is potential to generate a cell-free matrix that would orchestrate WAT transdifferentiation.

Our current generation of BAT-MACTs was viable for ∼30 days in vivo after ∼10 days in vitro culture. Duration of the implants is likely determined by a combination of the matrix susceptibility to biodegradation, through the MMP-sensitive crosslinker and HyA backbone, and the life span of brite/beige cells (∼50 days) (48). Importantly, we developed MMP-sensitive crosslinkers (30) to enable tissue remodeling and recruitment of host vasculature (Fig. 3C). MMP-13 has been shown to be essential for vascularization processes (49), and its expression is one of the most significantly upregulated MMPs during the differentiation of adipocytes (50). Additionally, crosslinkers designed to be sensitive to other MMPs appear to degrade too rapidly in vivo to be used for these applications.

In vivo experiments with BAT-MACTs validated prior notions about the metabolic impact of BAT expansion. Recipients exposed to cold demonstrated that a 100-µL BAT-MACT implant had a significant effect on thermogenesis in a cell-number–dependent manner. BAT-MACTs increased respiration rates and fatty acid utilization, reduced weight gain, and improved glucose homeostasis. These results are in line with recent reports demonstrating that transplantation of primary BAT fragments into mice can prevent diet-induced obesity (10). It remains to be determined whether these beneficial effects are solely due to the increased caloric expenditure afforded by the implant or if they also rely on an endocrine component such as interleukin-6 (10) or other factors secreted by UCP1-positive adipose.

In summary, beige adipose tissue can be engineered using an optimized bio-inspired HyA hydrogel system in conjunction with WAT-derived stem cells and can persist under defined spatio-temporal conditions to affect body temperature, energy homeostasis, weight gain, and insulin sensitivity. These data from our preclinical model demonstrate the feasibility of a novel autologous transplantation-based antiobesity approach with a clear path for ultimate translation into clinical practice. Importantly, the BAT-MACT approach will facilitate systematic evaluation of the metabolic effects of BAT expansion and the underlying biological mechanisms.

Acknowledgments. The authors thank the University of California, Berkeley, imaging core facility members Holly Aaron, Denise Schichnes, and Steve Ruzin for support. The authors also acknowledge Chris Baintner of FLIR for providing thermal imaging equipment.

Funding. This work was supported in part by National Institutes of Health grants R01-DK-101293 and R01-DK-089202 and the American Diabetes Association Basic Science Award 1-14-BS-191 awarded to A.S.

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. K.M.T. designed and carried out the main experiments with assistance from A.K.J., J.K., and A.Y. A.K.J. designed, produced, and assisted in biomaterials use. G.K., R.S., and E.A.D. designed and produced the FFA-SS-Luc probe. K.E.H. directed and guided bioengineering approaches. A.S. conceived the project, supervised and guided experimental approaches, and wrote the manuscript with assistance from K.M.T. A.S. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.