Isolation and Differentiation of Primary White and Brown Preadipocytes from Newborn Mice

Summary

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.

Abstract

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.

Introduction

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 action^2,3. 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 properties^4,5,6,7. 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 homeostasis^8,9,10. 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 mice^11,12,13,14. However, because the adipose depots of adult animals consist primarily of fully mature adipocytes that have a very slow turnover rate^15,16,17, 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.

Protocol

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)

1. Prepare two 1.5 mL tubes for each pup: one for brown adipose tissue (BAT) and one for white adipose tissue (WAT). Add 250 µL of phosphate-buffered saline (PBS) + 200 µL of 2x isolation buffer (123 mM NaCl, 5 mM KCl, 1.3 mM CaCl₂, 5 mM glucose, 100 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), penicillin-streptomycin, and 4% fatty acid-free bovine serum albumin) to each tube. Keep solutions sterile and on ice.
2. Place the pups into small chambers (e.g., one well of a 6-well plate), and set them on ice until they become hypothermic. Make sure that there is no direct contact between the pups and the ice. Prick a paw with a tip to assure loss of consciousness, and euthanize the pups by decapitation using sharp scissors.
   NOTE: if working with different genotypes, prepare an additional tube to collect a tail biopsy (3 mm cut) for genotyping. If genotyping is performed immediately, keep the euthanized pups on ice until the dissection to collect adipose depots.
3. Calculate and weigh out the amount of collagenase needed to digest all the depots. Add 50 µL of 15 mg/mL collagenase type I in 2x isolation buffer to each tube. Do not resuspend the collagenase in isolation buffer until all the tissues have been collected.
4. To collect subcutaneous WAT, cut the skin around the pup's abdomen (avoid peritoneal rupture), and gently pull the skin down below the legs. Without taking the skin, carefully collect the fat depot, which will appear as a clear (P1 or younger) or white (P2 and older), thin, elongated tissue attached on the inside or on top of the quadriceps (Figure 1B). Rinse the fat depot in PBS, and place it in one of the tubes containing 250 µL of PBS + 200 µL of 2x isolation buffer. Keep on ice.
   NOTE: P0 and P1 mice give the best yield. In P0-1 pups, the subcutaneous WAT depot is nearly transparent. In P2 mice and older, the depot is easier to identify because it is already turning white, indicating the presence of fully mature, lipid-loaded, white adipocytes.
5. To collect interscapular BAT, pull the skin from the shoulder blades over the head. Lift the BAT - the deep red tissue between the shoulder blades - and carefully make incisions all around it to detach it from the body (Figure 1B). Check for consistency and color; rinse with PBS; place it in the other tube containing 250 µL of PBS + 200 µL of 2x isolation buffer; and keep on ice.
   NOTE: When harvesting BAT from P2 mice and older, carefully remove the white adipose tissue surrounding the BAT. It consists of a thin, soft, white sheet of adipocytes located between the skin and the BAT, which is deeper between the shoulder blades.
6. Once all depots have been collected, gently mince each depot (4-6 times) using small scissors directly inside the tubes.
7. Resuspend collagenase type I in the appropriate volume of 2x isolation buffer to obtain a 10x stock solution at 15 mg/mL. Add 50 µL of 10x collagenase to each tube, keeping the tubes on ice until collagenase has been added to all the tubes.
8. Invert the tubes quickly to mix, and transfer to a temperature-controlled mixer. Incubate the samples for 30 min at 37 °C on a shaker (1,400 rpm frequency for effective sample mixing).

2. Plating preadipocytes (day 1)

1. After digesting the tissues, place the tubes back on ice.
   NOTE: From this step onwards, all work is carried out in sterile conditions in a biosafety cabinet.
2. Strain the digested tissues through a 100 µm cell strainer into new 50 mL tubes. If working only with WT mice, or if genotypes are known, pool the relevant, dissociated BATs together; repeat this for the WATs. To maximize the cell yield, rinse each tube with 1 mL of isolation medium (Dulbecco's modified Eagle medium (DMEM) + 20% fetal bovine serum (FBS), 10 mM HEPES, 1% penicillin-streptomycin), and filter it through the cell strainer. If working with unknown genotypes, go to step 2.4
   NOTE: FBS is an essential determinant of both preadipocyte proliferation and differentiation. Rigorously test different lots of FBS to ensure high performance and consistency between lots.
3. Dilute BAT and WAT suspensions to plate 2-3 wells of a 6-well plate for each BAT sample and 4-6 wells of a 6-well plate for each WAT sample. For instance, if 6 BATs and 6 WATs were pooled together, dilute the BAT cell suspension up to 24-36 mL and the WAT cell suspension up to 48-72 mL. Plate 2 mL of the cell suspensions per well.
4. For samples with unknown genotypes, keep each sample separate; place a 100 µm cell strainer on top of each well of a 6-well plate, and filter one sample per well. Rinse the tube with 1.5 mL of isolation medium, and pass it through the strainer, making up the final volume to 2 mL per well. Discard the cell strainer, place the lid on the plate, and transfer the plate to the incubator.
   NOTE: The medium in the wells should look turbid due to floating blood cells, cell debris, and lipids from lysed cells.
5. 1-1.5 h after plating, aspirate media and wash with 2 mL of DMEM without serum. Gently agitate the plate to detach blood cells from the bottom of the wells. After 3 washes, add 2 mL of fresh isolation medium, and transfer the cells to the incubator (37 °C, 5% CO₂).
   NOTE: Check the cells under the microscope. Floating material (blood cells, cellular debris) should be minimal. Brown preadipocytes will appear as small non-translucent cells, whereas white preadipocytes will display a more elongated form. Both brown and white preadipocytes should be tightly attached to the plate.

3. Expansion of preadipocyte culture (day 2 to day 5)

1. On the next day, aspirate the medium, wash the cells with 2 mL of DMEM without serum, and add back 2 mL of the isolation medium.
2. Repeat step 3.1 once every 2 days until the cells reach 80-90% confluence.
   NOTE: For primary brown preadipocytes, reaching 80-90% confluence can take 4-5 days. For primary white preadipocytes, it usually takes 2-3 days. Once the preadipocytes reach 60% confluence, they can be efficiently infected with viral particles for knockdown or overexpression experiments. Infect preadipocytes with an appropriate viral load for up to 8 h. The use of cationic polymers to increase infection efficiency is discouraged, as it often results in toxicity and in a significant reduction of the adipogenic potential of infected preadipocytes.
3. To split the cells, coat new destination plates using a sterile solution of 0.1% w/v gelatin (dissolved in distilled water; do not use any heat). Use enough volume to cover the bottom of the well/dish. Incubate plates at 37 °C until the cells are ready to be seeded (at least 10 min).
   NOTE: This step is optional. Coating of cell culture plates does not affect the yield or differentiation potential, but it substantially simplifies the maintenance and handling of differentiating adipocytes.
4. When cells reach sub-confluent density (85-95%), aspirate the medium, wash with PBS, and add trypsin for 3 min to detach the cells. Block trypsin activity by adding 2.5x trypsin volume of isolation medium. Pipette the cell suspension up and down to maximize cell recovery, and transfer into a new tube. Wash the wells with 1 mL of isolation medium, and add it to the first collection.
5. Centrifuge the cells for 5 min at 800-1200 × g, aspirate the supernatant, and resuspend the cells in 3-5 mL of the isolation medium. Count the cells, and dilute them to the desired final density.
   NOTE: For instance, 50,000-80,000 cells/mL (2.5, 1, and 0.5 mL for 6-, 12- and 24-well plates, respectively) will result in fully confluent wells within 72-96 h.
6. Aspirate the coating solution in step 3.3, and wash off excess gelatin with PBS. Seed the cells, and return the plates to the incubator until fully confluent.

4. Differentiation of white and brown preadipocytes (days 7 - 12)

1. When preadipocytes become confluent, aspirate the medium, and replace it with differentiation medium consisting of 10% FBS in DMEM containing 170 nM insulin, 1 µM dexamethasone, and 0.5 mM 3-isobuthyl-1-methylxantine. If differentiating BAT preadipocytes, also add 1 nM triiodothyronine (T3). Mark the day on which adipogenic differentiation is induced as day 0 of differentiation.
   NOTE: Both white and brown preadipocytes can spontaneously differentiate upon reaching confluence. However, to maximize adipocyte differentiation, the traditional pro-adipogenic cocktail described above (plus T3 for BAT preadipocytes) should be used.
2. After 48 h (day 2 of differentiation), refresh the medium with maintenance medium consisting of 10% FBS in DMEM and 170 nM insulin (plus 1 nM T3 for BAT).
   NOTE: On day 2, small lipid droplets accumulating in the cells under the microscope can be observed using standard bright field.
3. Repeat step 4.2 once every 2 days until cells are used for experiments.
   NOTE: On day 4 or 5 of differentiation, the majority of cells will appear loaded with lipids. Terminally differentiating adipocytes can be kept in culture longer or used at this stage for experiments.

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.

1. Prior to trypsin digestion, prepare coating for 96-well plates. Add 50 µL of 0.1% gelatin to each well. Leave the plate in a tissue culture incubator until the cells are ready to be seeded.
2. On day 4 or 5 of differentiation, aspirate the medium, wash with 1 mL of PBS, and add 300 µL of 0.25% trypsin-ethylenediamine tetraacetic acid (for 1 well of a 6-well plate). Gently tilt the plate to ensure trypsin completely covers the bottom of the well; incubate for 2 min at room temperature. Block the action of trypsin with 700 µL of maintenance medium, pipette up and down to maximize cell recovery, and transfer the cell suspension into a new 1.5 mL tube.
3. Centrifuge the cells at 1200 × g for 5 min. Remove the supernatant, resuspend the pellet in 1 mL of maintenance medium, and count the cells.
   NOTE: One well of a 6-well plate should yield approximately 1.5-1.8 million cells.
4. Dilute the cells to have a final concentration of 60,000 to 80,000 cells/mL for brown adipocytes and 100,000 to 120,000 cells/mL for white adipocytes. Plate 100 µL of the cell suspension per well in gelatin-coated 96-well plates.
   NOTE: One well of a 6-well plate provides enough cells for up to two 96-well plates. For bioenergetics experiments, low-density plating (30-40% confluence) is needed to enable the accurate measurement of oxygen consumption and avoid oxygen depletion in the wells during the assay.
5. Allow the cells to adhere to the bottom of the plate for at least 48 h. If cells are kept in the plate for more than 48 h, refresh the medium after 2 days.
6. On the day of the assay, aspirate the medium, and replace it with assay medium. Incubate for 1 h at 37 °C in a non-CO₂-controlled incubator, and perform the assay as per the manufacturer's instructions.

Results

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 (Figu...

Discussion

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 depots^21,22...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
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