This report describes a protocol for the simultaneous isolation of primary brown and white preadipocytes from newborn mice. Isolated cells can be grown in culture and induced to differentiate into fully mature white and brown adipocytes. The method enables genetic, molecular, and functional characterization of primary fat cells in culture.
The understanding of the mechanisms underlying adipocyte differentiation and function has greatly benefited from the use of immortalized white preadipocyte cell lines. These cultured cell lines, however, have limitations. They do not fully capture the diverse functional spectrum of the heterogenous adipocyte populations that are now known to exist within white adipose depots. To provide a more physiologically relevant model to study the complexity of white adipose tissue, a protocol has been developed and optimized to enable simultaneous isolation of primary white and brown adipocyte progenitors from newborn mice, their rapid expansion in culture, and their differentiation in vitro into mature, fully functional adipocytes. The primary advantage of isolating primary cells from newborn, rather than adult mice, is that the adipose depots are actively developing and are, therefore, a rich source of proliferating preadipocytes. Primary preadipocytes isolated using this protocol differentiate rapidly upon reaching confluence and become fully mature in 4-5 days, a temporal window that accurately reflects the appearance of developed fat pads in newborn mice. Primary cultures prepared using this strategy can be expanded and studied with high reproducibility, making them suitable for genetic and phenotypic screens and enabling the study of the cell-autonomous adipocyte phenotypes of genetic mouse models. This protocol offers a simple, rapid, and inexpensive approach to study the complexity of adipose tissue in vitro.
Obesity results from a chronic imbalance between energy intake and energy expenditure. As obesity develops, white adipocytes undergo a massive expansion in cell size that results in hypoxia in the microenvironment, cell death, inflammation, and insulin resistance1. Dysfunctional, hypertrophied adipocytes cannot properly store excess lipids, which accumulate instead in other tissues where they dampen insulin action2,3. Agents that improve adipocyte function and restore normal lipid partitioning amongst tissues are predicted to be beneficial for the treatment of obesity-associated conditions characterized by insulin resistance such as type 2 diabetes. Phenotypic screens in adipocytes using immortalized cell lines, such as 3T3-L1, F442A, and 10T ½, have proven useful to identify genetic factors that regulate adipogenesis and to isolate pro-adipogenic molecules with anti-diabetic properties4,5,6,7. These cell lines, however, do not fully reflect the heterogeneity of cell types present in adipose depots, which includes white, brown, beige, and other adipocyte subtypes with unique characteristics, all of which contribute to systemic homeostasis8,9,10. Further, cultured cell lines often show a diminished response to external stimuli.
In contrast, cultures of primary adipocytes recapitulate more accurately the complexity of in vivo adipogenesis, and primary adipocytes show robust functional responses. Primary preadipocytes are typically isolated from the stromal vascular fraction of adipose depots of adult mice11,12,13,14. However, because the adipose depots of adult animals consist primarily of fully mature adipocytes that have a very slow turnover rate15,16,17, this approach yields a limited quantity of preadipocytes with a low proliferation rate. Therefore, isolation of preadipocytes from newborn mice is preferable to obtain large quantities of rapidly growing cells that can be differentiated in vitro. Here, a protocol has been described, inspired by the initial work with primary brown adipocytes of Kahn et al.18 to efficiently isolate both white and brown preadipocytes that can be expanded and differentiated in vitro into fully functional primary adipocytes (Figure 1A). The advantage of isolating primary cells from newborn, as opposed to adult mice, is that the adipose depots are rapidly growing and are thus a rich source of actively proliferating preadipocytes17. Cells isolated using this protocol have high proliferative capacity, enabling rapid scale-up of cultures. In addition, preadipocytes from newborn pups display higher differentiation potential than adult progenitors, which reduces well-to-well variability in the extent of differentiation and thus increases reproducibility.
This protocol follows all IACUC guidelines of The Scripps Research Institute and the University of Wisconsin – Madison School of Medicine and Public Health.
1. Collection and digestion of adipose depots (day 1)
2. Plating preadipocytes (day 1)
3. Expansion of preadipocyte culture (day 2 to day 5)
4. Differentiation of white and brown preadipocytes (days 7 – 12)
5. Re-plating for bioenergetics experiments
NOTE: If the intention is to perform bioenergetics studies in mature adipocytes in the 96-well format, the following steps need to be taken. Ideally, cells that have just fully differentiated (day 4 or 5) should be used. The procedure described below starts from 1 well of a 6-well plate.
Section 1 of the protocol will yield a heterogeneous suspension of cells that are visible under a standard light microscope. Filtering of digested tissues with a cell strainer (section 2) will remove undigested tissue. However, some cellular debris, blood cells, and mature adipocytes will pass through (Figure 1C). Gentle washes 1 h after plating will remove non-relevant cells as preadipocytes attach rapidly to the bottom of the well (Figure 1C). In section 3, adipocyte precursors are expanded to obtain the number of cells required for the experimental plan. Although both white and brown preadipocytes isolated from newborn pups have high proliferative capacity, the yield of white preadipocytes is generally twice that of brown preadipocytes on a per-depot basis (Figure 2A). Therefore, if synchronized cultures are desired, the starting density of white preadipocytes must be calculated accordingly. Section 4 offers guidelines to obtain fully mature adipocytes.
At the end of differentiation, cells will appear loaded with lipid droplets and express classical markers of white and brown adipocytes, respectively (Figure 2B,C). Both white and brown adipocytes can be used for bioenergetics studies as described in section 5. A comparative analysis of oxygen consumption in a mitochondrial stress test of primary white and brown adipocytes under basal conditions, as well as in response to known stimulators of mitochondrial function (e.g., norepinephrine) is shown in Figure 2D. Upon isolation, it is also possible to differentiate primary white and brown adipocytes on the plates that will be directly used for bioenergetic experiments19. In this case, preadipocytes are isolated and plated as described in sections 1 and 2. When cells become confluent, they are induced to differentiate as described in section 4 until they reach terminal differentiation and are ready for bioenergetics analysis. This procedure is common for a 24-well plate format, but less so for 96-well plates.
Figure 1: Collection and processing of fat pads. (A) Schematic representation of primary white (top) and brown (bottom) adipocyte isolation. (B) Subcutaneous white (top) and brown (bottom) adipose depots. In P0 mice, subcutaneous WAT is almost invisible, but becomes distinguishable on ~day 2 after birth. In contrast, BAT has a distinct dark color even at P0. In older pups, the BAT is surrounded by a thin superficial layer of WAT, which requires removal when the tissue is dissected. (C) Representative images of primary white and brown precursor cells after filtration through the 100 µm cell strainer, after the initial washes, and 24 h after isolation. Scale bars = 100 µm. Abbreviations: WAT = white adipose tissue; BAT = brown adipose tissue. Please click here to view a larger version of this figure.
Figure 2: Differentiation of adipocyte progenitors. (A) Average number of subcutaneous WAT and BAT preadipocytes obtained per newborn (P0) pup. (B) Representative images of terminally differentiated white and brown adipocytes. For fluorescence imaging, cells were incubated with Nile Red and Hoechst 33342 to stain neutral lipids and nuclei, respectively. Scale bars = 100 µm.(C) Gene expression analysis of classical adipocyte markers, white-beige markers, and brown-specific markers in primary white and brown adipocytes differentiated for 6 days (n=3). For each gene, BAT expression is relative to WAT levels (set to 1). (D) OCR of white and brown adipocytes in a mitochondrial stress test and in response to norepinephrine (n=3). Brown adipocytes show uncoupled respiration and a robust response to norepinephrine, whereas white adipocytes show little uncoupled respiration and no significant response to adrenergic stimulation. *p<0.05; **p<0.01, determined by two-tailed Student's t-test. Abbreviations: WAT = white adipose tissue; BAT = brown adipose tissue; OCR = oxygen consumption rate; Oligo = oligomycin; FCCP = carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone; RAA = rotenone, antimycin A; PPARγ = peroxisome proliferator-activated receptor gamma; Acrp30 = adipocyte complement-related protein of 30 kDa; Fabp4 = fatty acid-binding protein 4; Glut4 = glucose transporter 4; Cd36 = cluster of differentiation 36; Retn = resistin; Slc27a1 = solute carrier family 27 member 1; Ear2 = eosinophil-associated, ribonuclease A family 2; Pgc1a = PPARγ coactivator 1alpha; Prdm16 = positive regulatory domain I-binding factor 1 (PRDI-BF1) and retinoblastoma protein-interacting zinc finger gene 1 (RIZ1) homologous domain containing 16; Eva1 = epithelial V-like antigen 1; Cidea = cell death-inducing DNA fragmentation factor-alpha-like effector A; Ucp1 = uncoupling protein 1; Dio2 = type 2 deiodinase. Please click here to view a larger version of this figure.
Adipose tissue is critical for systemic insulin sensitivity and glucose homeostasis20. Obesity-linked adipocyte dysfunction is tightly associated with the onset of type 2 diabetes. Therefore, greater understanding of the basic biology and physiology of adipose tissue may enable the design of new treatments for metabolic disorders. As a complement to direct functional and transcriptional analysis of mature adipocytes isolated from fat depots21,22, cultured primary adipocytes have been shown to recapitulate many aspects of adipose tissue pathophysiology, including secretion of adipokines, resistance to insulin in response to pro-inflammatory stimuli, and induction of the thermogenic program23,24,25. Although previous protocols have described the isolation of adipocyte precursors from adult mice11,12, this protocol provides a method for the efficient isolation of cells from newborn pups. This strategy yields a significantly larger population of white and brown adipocyte progenitors with higher differentiation potential, as postnatal adipose depots are still relatively undifferentiated compared to the adipose depots of adult mice26. Moreover, cells isolated using this method are already committed and yield fully functional differentiated adipocytes that express markers of mature cells and exhibit their unique physiological features, including lipid storage and thermogenic capacity.
Primary cells cannot be expanded indefinitely in vitro. In addition, primary preadipocytes start to lose their adipogenic potential after several passages in culture. This is likely because continual cell culture results in an intrinsic enrichment of less committed, more proliferative cells. Thus, one limitation of this protocol is the window of time during which preadipocytes can be used. Adipogenic potential is fully preserved if cells are induced to differentiate within 7-8 days from the time of isolation. Adipocyte progenitors are particularly resistant to enzymatic digestion, but it is nonetheless important to correctly time the collagenase treatment. Overdigestion of tissues may result in reduced cell survival and decreased ability of cells to adhere to the plate. Throughout the expansion phase, both white and brown preadipocytes are strongly adherent and can tolerate vigorous washes and frequent media exchange. However, the use of coated plates is recommended when preadipocytes are induced to differentiate. Mature, lipid-laden adipocytes have decreased surface adhesion and cell-cell interactions, resulting in a tendency to detach during differentiation unless gently handled. The most delicate step of the protocol is the induction of differentiation. FBS, insulin, T3, and other drugs must be tested, and their final concentrations optimized, to obtain the highest extent of differentiation. A PPARγ agonist (e.g., rosiglitazone) can also be added to further stimulate adipogenesis.
It is important to note that the differentiation conditions can be adapted to the experimental needs. For instance, in screens with genetic or chemical libraries designed to identify proteins/compounds that enhance white and/or brown adipocyte differentiation, preadipocytes can be induced to differentiate using a minimal permissive medium that includes 10% FBS in DMEM and 170 nM insulin only. Assessment of each component of the differentiation cocktail is recommended to determine ideal assay windows for differentiation assays. T3 and dexamethasone are dispensable for primary white and brown adipocyte differentiation. These conditions will ensure a low rate of differentiation in control cells, thus increasing the window of the assay and maximizing the ability to detect pro-adipogenic factors. Cultures of primary brown and white adipocytes are a powerful tool to interrogate adipocyte cell-autonomous function in response to genetic manipulation and metabolic stresses to complement the study of brown and white adipose tissue in vivo. Hence, protocols for isolation and culture of primary preadipocytes are needed to enable reproducible, high-throughput investigations of adipocyte function in vitro. The strategy described here allows study of primary white and brown preadipocytes that can be differentiated into fully mature adipocytes and tested under a variety of experimental manipulations.
The authors have nothing to disclose.
The authors are grateful to Cristina Godio at Centro Nacional de Biotecnología in Madrid, Spain, Mari Gantner at The Scripps Research Institute, La Jolla, and Anastasia Kralli at Johns Hopkins University, Baltimore, for assistance optimizing this protocol based on the initial work of Kahn et al.18. This work was funded by NIH grants DK114785 and DK121196 to E.S.
3-Isobutyl-1-methylxanthine (IBMX) | Sigma-Aldrich | I7018 | |
6-well plates | Corning | 353046 | |
AdipoRed (Nile Red) | Lonza | PT-7009 | |
Antimycin A | Sigma-Aldrich | A8674 | |
BenchMark Fetal Bovine Serum | Gemini Bioproducts LLC | 100-106 | |
CaCl2 | Sigma-Aldrich | C4901 | |
Cell strainer | Fisher Scientific | 22363549 | |
Collagenase, Type 1 | Worthington Biochemical Corp | LS004196 | |
ddH2O | Sigma-Aldrich | 6442 | |
Dexamethasone | Sigma-Aldrich | D4902 | |
DMEM | Sigma-Aldrich | D5030 | For Bioenergetics studies |
DMEM, High Glucose, Glutamax | Gibco | 10569010 | |
DPBS, no calcium, no magnesium | Gibco | 14190144 | |
Fatty Acid-Free BSA | Sigma-Aldrich | A8806 | |
FCCP | Sigma-Aldrich | C2920 | |
Gelatin | Sigma-Aldrich | G1890 | |
Glucose | Sigma-Aldrich | G7021 | |
HEPES | Sigma-Aldrich | H3375 | |
Hoechst 33342 | Invitrogen | H1399 | |
Insulin | Sigma-Aldrich | I6634 | |
KCl | Sigma-Aldrich | P9333 | |
NaCl | Sigma-Aldrich | S7653 | |
Norepinephrine | Cayman Chemical | 16673 | |
Oligomycin | Sigma-Aldrich | 75351 | |
Pen/Strep | Gibco | 15140122 | |
Rosiglitazone | Sigma-Aldrich | R2408 | |
Rotenone | Sigma-Aldrich | 557368 | |
Seahorse XFe96 FluxPak | Agilent Technologies | 102416-100 | For Bioenergetics studies |
Surgical forceps | ROBOZ Surgical Instrument Co | RS-5158 | |
Surgical Scissors | ROBOZ Surgical Instrument Co | RS-5880 | |
ThermoMixer | Eppendorf | T1317 | |
triiodothyronine (T3) | Sigma-Aldrich | 642511 |