Isolation and adipogenic differentiation of murine mesenchymal stem cells harvested from macrophage-depleted bone marrow and adipose tissue

ABSTRACT

Introduction and Purpose

Mouse mesenchymal stem cells (MSCs) provide a resourceful tool to study physiological and pathological aspects of adipogenesis. Bone marrow-derived MSCs (BM-MSCs) and adipose tissue-derived MSCs (ASCs) are widely used for these studies. Since there is a wide spectrum of methods available, the purpose is to provide a focused hands-on procedural guide for isolation and characterization of murine BM-MSCs and ASCs and to effectively differentiate them into adipocytes.

Methods and Results

Optimized harvesting procedures for murine BM-MSCs and ASCs are described and graphically documented. Since macrophages reside in bone-marrow and fat tissues and regulate the biological behaviour of BM-MSCs and ASCs, we included a procedure to deplete macrophages from the MSC preparations. The identity and stemness of BM-MSCs and ASCs were confirmed by flow cytometry using established markers. Since the composition and concentrations of adipogenic differentiation cocktails differ widely, we present a standardized four-component adipogenic cocktail, consisting of insulin, dexamethasone, 3-isobutyl-1-methylxanthine, and indomethacin to efficiently differentiate freshly isolated or frozen/thawed BM-MSCs and ASCs into adipocytes. We further included visualization and quantification protocols of the differentiated adipocytes.

Conclusion

This laboratory protocol was designed as a step-by-step procedure for harvesting murine BM-MSCs and ASCs and differentiating them into adipocytes.

Introduction

Mesenchymal stem cells (MSCs) are multipotent stem cells that self-renew and differentiate into various cell types, including adipocytes, osteoblasts, chondrocytes, and myocytes [1]. MSCs can be isolated from tissues such as bone marrow, fat, lungs, liver, placenta, umbilical cord, and skin [2]. Bone marrow and white adipose tissue are two widely used sources of bone marrow-derived MSCs (BM-MSCs) and adipose tissue-derived MSCs (ASCs), respectively [3,4]. BM-MSCs and ASCs are commonly employed to study adipogenic differentiation in cell culture because of their robust stemness and proliferative capacity [3–5]. Wild-type mice and genetically engineered mouse models provide a useful source of BM-MSCs and ASCs for studying various molecular aspects of adipogenic differentiation. While many protocols have been published to isolate BM-MSCs and ASCs from mice and to differentiate them in cell culture into adipocytes, detailed hands-on protocols for these procedures are lacking. Therefore, this study aimed to provide an effective, highly reproducible step-by-step protocol to isolate murine BM-MSCs and ASCs and to differentiate them into adipocytes.

Murine bone marrow consists of heterogeneous cell types, including mesenchymal stem cells, haematopoietic stem cells, and resident macrophages. Similarly, the stromal vascular fraction (SVF) derived from adipose tissue comprises a heterogeneous population of cells, including mesenchymal stem cells, preadipocytes, adipocytes, and macrophages [4]. Most of the contaminating cells were removed during the isolation procedures described in this article. However, BM-MSCs isolated from bone marrow are often contaminated by macrophages [5,6]. Although somewhat less of a problem, macrophage contamination can also occur when ASCs are isolated from SVF. Macrophage contamination is a confounding factor for downstream differentiation. For example, incubation of BM-MSCs in medium containing macrophages leads to enhanced osteogenic differentiation, whereas inhibition of macrophages results in impaired osteogenic potential of BM-MSCs [7,8]. Pro-inflammatory cytokines produced by macrophages inhibit adipogenic differentiation of ASCs [9]. Since the presence of macrophages can modulate the differentiation potential of BM-MSCs and ASCs, it is necessary to deplete macrophages from MSC preparations for adipogenic differentiation. Macrophages can be depleted using different methods, such as magnetic bead-based separation or the liposomal chlodronate approach [10–12]. In this paper, we focus on magnetic bead separation for effectively depleting macrophages from isolated MSCs, as it is a simple and reproducible method that allows precise and targeted removal of macrophages while preserving stem cell integrity and function.

The formation of mature adipocytes from precursor stem cells, termed adipogenesis, involves a complex and highly coordinated program of gene expression [13]. MSCs give rise to white adipocyte precursor cells, known as adipoblasts, which, under appropriate stimulatory conditions, commit to form white preadipocytes [14]. These committed preadipocytes first maintain their proliferative capacity and then undergo differentiation. During the differentiation phase, cells acquire spherical morphology and form mature adipocytes. This is accompanied by dramatic changes in the extracellular matrix [15]. A full understanding of the complex and highly orchestrated program of adipocyte formation is required to understand adipose tissue-associated metabolic diseases and genetic disorders. Exposure of MSCs to a three-component adipogenic stimulus, including insulin, dexamethasone, and 3-isobutyl-1-methylxanthine (IBMX) is frequently used [16]. A growing body of evidence also highlights the use of a four-component adipogenic medium containing insulin, dexamethasone, IBMX, and indomethacin [17–20]. However, the components and concentrations of the major inducers vary among studies. Many researchers have altered the concentrations and components to obtain higher differentiation efficiency [21]. Other studies have used proprietary formulations for adipogenic induction with undisclosed components [22]. Considering this wide spectrum of methods, it is important to standardize adipogenic differentiation cocktails and procedures so that results can be readily compared and interpreted.

This method paper provides detailed experimental steps to harvest BM-MSCs and ASCs from mouse bone marrow and adipose tissues, respectively. We explain the steps involved in depleting macrophages from MSC preparations, after which they can be used for adipogenic differentiation experiments. We outlined the phenotypic characterization of the harvested BM-MSCs and ASCs by flow cytometric analysis. Furthermore, we demonstrated that the four-component cocktail at the indicated concentrations induced both freshly isolated and frozen BM-MSCs and ASCs towards the adipogenic lineage. Finally, we explain the details of the detection method for differentiated BM-MSCs and ASCs using Oil Red O staining and by quantifying the stained cells for statistical analysis.

Protocols

Please refer to Supplemental Table S1 for details on the materials and instruments used in the protocols.

1. Harvesting BM-MSCs

A schematic of the steps to harvest BM-MSCs from the tibiae and femurs of mice is provided in Figure 1.

Figure 1.

Schematic of the steps to harvest BM-MSCs from tibiae and femurs of mice. Schematics demonstrating the steps required to harvest bone marrow derived mesenchymal stem cells (BM-MSCs) from the tibiae and femurs of mice as explained in the protocol.

NOTE: All surgical tools should be either autoclaved or sterilized with 70% (v/v) ethanol.

• 1.1. Complete culture medium preparation

  1.1.1 Heat-inactivate foetal bovine serum (FBS) by placing a conical centrifuge tube containing FBS for 30 min in a water bath maintained at 56°C. Gently shake the tube every 10 min for equal heat distribution and to dissolve aggregates that might settle at the bottom.

• 1.1.2 Prepare complete culture medium by supplementing Minimum Essential Medium alpha (MEMα) with 10% heat-inactivated FBS supplemented with 1 µg/mL penicillin- streptomycin, and 0.02 mM glutamine.

• 1.2 Preparation of 1.5 mL reaction tubes containing the insert for extraction of bone marrow from bone shafts

• Schematic of the steps for customized tube preparation to extract BM-MSCs is provided in Figure 2.

• 1.2.1 Cut the bottom end of 1 mL pipette tips so that they can fit well in 1.5 mL reaction tubes and autoclave both the tube and cut tip separately before use.

• 1.2.2 Under a sterile laminar flow hood, place the cut tips inside 1.5 mL reaction tubes and ensure that the lids close well.

• NOTE: Alternatively, 0.75 mL microcentrifuge tubes with both ends cut can be used for this step.

• 1.3 Euthanizing the experimental mice

  1.3.1 Young donor mice of 4–6 weeks of age weighing about 18–20 g are ideal for BM-MSC isolation.

• NOTE: Young mice of this age tend to have higher numbers of multipotent MSCs within their bone marrow cavities than older mice.

• 1.3.2 Place the mouse selected for dissection in an anaesthesia chamber and euthanize the mouse in a humane manner by CO₂ asphyxiation under isoflurane anaesthesia followed by cervical dislocation.

• 1.4 Dissecting the experimental mice

  1.4.1 Pin down the mouse on a dissection board with the ventral side facing up.

  1.4.2 Swab the mouse with 70% (v/v) ethanol or other appropriate disinfectants, which has the added benefits of wetting the fur.

  NOTE: Washing the carcass with ethanol or disinfectant is necessary to reduce hair and other possible contamination.

Figure 2.

Schematic of the steps for customized tube preparation to extract BM-MSCs. Steps to prepare the customized tubes required to collect bone marrow after the bones are centrifuged. Briefly, 1 mL pipette tips were cut using a scalpel so that they fit properly in 1.5 mL reaction tubes. Both the tube and the cut tip were separately before use.

• 1.4.3 Using forceps, grab and hold the skin anteriorly to the urethral opening. Using sterile dissecting scissors, insert a small incision (about 1 cm) along the ventral midline from the groin to the chin, being careful to only cut the skin and not the underlying muscle wall.

• 1.4.4 Cut the abdominal skin and remove the skin from the animal in a rapid motion that pulls it off the torso and down onto the extremities to expose the lower abdomen and legs.

• 1.4.5 Remove the whole bone shaft by detaching it from the hipbone at femur head from both the right and left legs.

• 1.4.6 Remove the tibia and femur from each side and scrape off the adjacent adherent tissue using a scalpel.

• NOTE:a) Cutting the tendons and muscles from the bones as much as possible makes cleaning faster and easier using lint-free wipes (example: Kimwipes).

• b) Care must be taken when manipulating the bones, as the cleaning procedures tend to break the bones easily and the extent of bone fragility may vary depending on the genotype, age, or sex.

• 1.4.7 Remove the end of the bones by cutting at the epiphyses using sterile surgical scissors.

• 1.4.8 Once harvested, hold the shaft of the tibia and femur by sterile forceps and place them into a 10 mL centrifuge tube containing sterile isotonic phosphate-buffered saline (PBS) maintained on ice.

• NOTE: Process the collected bones quickly (within 30 min) as any delay could result in compromised sub-optimal MSCs.

• 1.5 Processing of bones to collect bone marrow

• 1.5.1 Move the collected bone samples to a sterile laminar flow hood for further steps of isolation. NOTE: Work aseptically from this step onwards to minimize the possibility of contaminations in the cell culture.

• 1.5.2 Rinse the tibia and femur thoroughly with 10 mL PBS to remove any remaining adherent particles, hair, or blood by transferring the bone shafts in a sterile petri plate.

• 1.5.3 Cut bones horizontally in two halves using pre-sterilized surgical scissors to ensure that the bone marrow is released from the bones as described in the subsequent steps.

• NOTE:a) Alternatively, bone marrow can be also effectively harvested by a ‘Bone flush method’ using a liquid-filled syringe as described in [23].

  b) Keep the surrounding area sterile and clutter-free so that in case the bone slips off while cutting it into two halves, it is still possible to use it.

• 1.5.4 Carefully place one bone cut into two halves with the cut end facing down for each bone in one 1.5 mL reaction tube containing the insert (Figure 2).

• 1.5.5 Centrifuge the tubes at 16,000 × g at ambient temperature three times for 30 s each. Ensure that the entire marrow has pelleted at the bottom of the tubes while the bones are held inside the inserts.

• NOTE: The bones from which the entire marrow is flushed out will appear almost white. If some bones still retain some marrow evident by a more red colour, those tubes should be centrifuged again to flush out the remaining marrow as much as possible.

• 1.5.6 Add 200 µL of the complete culture medium to the outside of each insert contained in the tube and then carefully remove the insert using sterile forceps.

• NOTE: Add the complete culture medium first before removing the inserts from the tube as the marrow is sticky and tends to be removed with the insert.

• 1.5.7 Pipette the cell pellets up and down several times using a 1 mL pipette to slowly suspend the marrow until a homogenous suspension is obtained. Pool the marrow suspension from femur and tibia of one mouse into a single 15 mL centrifuge tube.

• NOTE: It is important to cut the ends of the 1 mL of pipette tips to reduce shear forces that may damage the cells.

• 1.5.8 Add 10 mL of the culture medium for BM-MSCs to the homogenous cell suspension.

• NOTE: Do not let the marrow dry as the cells will lose their potential to differentiate.

• 1.5.9 Centrifuge the marrow suspension at 180 × g for 5 min which results in a cell pellet. Aspirate the supernatant and resuspend the pellet in 10 mL complete culture medium by pipetting it repeatedly using 1 mL pipette tips (with cut ends) to obtain the homogenous cell suspension.

• 1.5.10 Plate the cell suspension containing the marrow cells onto the surface of 75 cm² flasks and incubate at 37⁰C in a 5% CO2 atmosphere.

• 1.5.11 Change medium every 2–3 days until the cells become confluent. It typically takes about one week to reach 80–90% confluency.

• 1.6 Depletion of macrophages

• 1.6.1 Once the isolated BM-MSCs reach 80–90% confluency, trypsinize the cells (detailed steps of trypsinization described below) and suspend 1 × 107 BM-MSCs in 100 µl cold isolation buffer (PBS with 0.5% bovine serum albumin, 2 mM ethylenediamine tetra acetic acid (EDTA)).

• 1.6.2 Add 50 µl Fc blocking antibody (49.5 µl of isolation buffer with 0.5 µl purified anti-mouse CD16/32 antibody) to reduce binding of antibodies to CD16 (Fc receptor III) and CD32 (Fc receptor II) both expressed by macrophages. Mix gently and incubate at 4°C for 15 min.

• 1.6.3 Add 50 µL of biotinylated CD45 antibody. Mix gently and incubate at 4°C for 15 min.

• 1.6.4 Wash cells with 1.6 mL isolation buffer.

• 1.6.5 Centrifuge at 500 × g for 5 min at ambient temperature.

• 1.6.6 Discard supernatant and resuspend cells in 1.8 mL isolation buffer. Centrifuge at 500 × g for 5 min at ambient temperature.

• 1.6.7 Resuspend the pellet in 500 µL isolation buffer and add 80 µL of streptavidin ferrofluid. Mix gently and incubate at 4°C for 20 min. The streptavidin-coated particles will bind to the cells labelled with the biotinylated CD45 antibody.

• 1.6.8 Add 1.3 mL of isolation buffer, mix gently to ensure all the cells are in suspension.

• 1.6.9 Place tube in the magnetic field using commercially available magnet that has been positioned horizontally and incubate for 7 min at ambient temperature. The biotin-labelled cells will be captured on the side of the tube adjacent to the magnet, and the macrophage depleted BM-MSCs will remain in suspension.

• 1.6.10 With a 1 mL transfer pipette, carefully collect the cells to a new 2 mL tube without disturbing the magnetically trapped macrophages on the side (brown colour). Discard the magnetically trapped macrophages.

• 1.6.11 Repeat the magnetic macrophage depletion (steps 1.6.8 and 1.6.9) in a new tube containing the recovered BM-MSCs.

• 1.6.12 The cell suspension obtained after these steps contains the macrophage-depleted BM-MSCs.

2. Harvesting ASCs

Schematic of the steps to harvest ASCs from various adipose tissue depots of mice is provided in Figure 3.

Figure 3.

Schematic of the steps to harvest ASCs from various adipose tissue depots of mice. Schematic demonstrating the steps required to harvest the adipose tissue derived mesenchymal stem cells (ASCs) from subcutaneous, gonadal, and retroperitoneal fat depots of mice as explained in the protocol.

• NOTE: All surgical tools should be either autoclaved or sterilized with 70% (v/v) ethanol.

• 2.1 Complete culture medium preparation

• 2.1.1 Heat-inactivate foetal bovine serum (FBS) by placing a sterile tube containing FBS for 30 min in a water bath maintained at 56°C. Gently shake the tube every 10 min for equal heat distribution and to dissolve aggregates that might settle at the bottom.

• 2.1.2 Prepare complete culture medium by supplementing Minimum Essential Medium alpha (MEMα) with 10% heat-inactivated FBS supplemented with 1 µg/mL penicillin- streptomycin, and 0.02 mM glutamine.

  Preparation of collagenase, type I

• 2.1.3 Dissolve 2 mg/mL of collagenase, type 1 lyophilized powder (Source: Clostridium histolyticum) in the complete culture medium. Filter-sterilize the collagenase solution with a sterile 0.22 µm syringe filter.

• 2.2 Euthanizing the experimental mouse

• Young donor mice of 6–8 weeks of age weighing about 20–22 g are ideal for isolation of ASCs.

• NOTE: Young mice of this age tend to have higher numbers of multipotent MSCs in the adipose tissues than older mice.

• 2.2.1 Place the mouse selected for dissection in an anaesthesia chamber and euthanize the mouse in a humane manner by CO2 asphyxiation under isoflurane anaesthesia followed by cervical dislocation.

• 2.3 Dissecting the experimental mouse

• 2.3.1 Pin down the mouse on a dissection board with the ventral side facing up. 2.3.2 Swab the mouse with 70% (v/v) ethanol or other appropriate disinfectants, which has the added benefits of wetting the fur.

• NOTE: Washing the carcass with 70% (v/v) ethanol or disinfectant is necessary to reduce hair and other possible contamination.

• 2.3.3 Using forceps, grab and hold the skin anteriorly to the urethral opening. Using sterile dissecting scissors, insert a small incision (about 1 cm) along the ventral midline from the groin to the chin, being careful to only cut the skin and not the underlying muscle.

• 2.3.4 Separate the skin and attached inguinal fat pad from the body by pulling the skin in a rapid motion using blunt tip scissors or simply by using gloved fingers.

• NOTE: The adipose tissue exposed underneath the skin is the subcutaneous fat, also called inguinal fat.

• 2.3.5 Locate and hold the end of one white inguinal fat depot and gently cut it out along its entire length with scissors. The tissue can be removed either as a single long pad or in several small pieces.

• 2.3.6 Similarly remove the inguinal fat from the other side and pool the fat pads from both sides using sterile forceps. Transfer them to a 10 mL centrifuge tube containing sterile isotonic PBS maintained on ice.

• 2.3.7 To access the gonadal adipose tissue, cut the abdominal muscle using clean and sterile surgical scissors and forceps along the linea alba from the genitals to the rib cage. The fat tissue surrounding the reproductive organs is the gonadal fat.

• 2.3.8 Harvest the gonadal fat pads from both the right and the left side and pool them in the same centrifuge tube containing PBS maintained on ice.

• 2.3.9 To isolate the retroperitoneal fat, expose the kidney by moving the gut away from the abdominal cavity. The adipose tissue beneath the kidneys represents the retroperitoneal fat depot. Pool the retroperitoneal fat tissue from both sides and transfer them to the same centrifuge tube containing PBS maintained on ice.

• NOTE: a) Process the harvested fat tissue as soon as possible. This is critical to maintain the functional status of the cells and is an important consideration if many samples are being harvested.

• b) To obtain a sufficient number of cells, it might be necessary to combine fat from all major fat depots of the mouse. However, this depends on the study requirements. If the experiment requires cells from only one specific adipose tissue, then cell numbers might pose a limitation for the planned experiments.

• 2.4 Processing the adipose tissue

• 2.4.1 Move the collected adipose tissue samples to a sterile laminar flow hood for further steps of isolation.

• NOTE: Work aseptically from this step onwards to minimize the possibility of contamination in the cell culture to be established.

• 2.4.2 Wash the adipose tissue prior to processing by sterile PBS to remove excess blood from the tissue. Repeat this washing step once.

• 2.4.3 Transfer the rinsed tissue to a 10 cm plastic or glass petri dish and mince the adipose tissue to a very fine consistency using sharp sterile surgical scissors. To optimize the subsequent enzymatic digestion of the tissue, the minced product should be homogenous and have an appropriate slimy consistency.

• NOTE: Do not touch the tip of surgical scissors on the surface of plastic or glass petri plate as it will damage the scissors.

• 2.4.4 Transfer the minced tissue to a sterile 15 mL centrifuge tube and further enzymatically digest the tissue using 2 mL of the filter-sterilized 2 mg/mL collagenase, type 1 solution. Incubate at 37⁰C in a 5% CO2 incubator for 1 h to ensure optimal collagenase activity.

• 2.5 Isolating the stromal vascular fraction (SVF)NOTE: SVF is the source of multipotent stem cells.

• 2.5.1 Separate the cells from the tissue by gravity through a 70 µm sterile cell strainer placed on the top of a 50 mL centrifuge tube. A uniform single-cell suspension of adipocytes will collect in the tube and the tissue is retained in the strainer. NOTE: a) Pre-wet the strainer with 1 mL of complete culture medium to ensure a smooth flow-through of cells. b) Tap the strainer on the tube to release remaining cells on the strainer.

• 2.5.2 Add 2 mL of complete culture medium to stop the action of collagenase, type 1.

• 2.5.3 Centrifuge the tubes at 180 × g for 5 min at ambient temperature. This will result in three phases: i) The top viscous, yellow-coloured layer containing mature adipocytes, ii) an interphase which contains medium, and iii) the SVF that appears as a dark red layered pellet at the bottom.

• 2.5.4 Carefully aspirate and discard the top layer with mature adipocytes and the medium-containing interphase, while retaining the bottom SVF pellet.

• 2.5.5 Since the SVF pellet contains the red blood cells, it is lysed in 1 mL of red blood cell lysing buffer (Supplemental Table S1). Incubate at ambient temperature for 5 min. 2.5.6 Wash the cells with 9 mL of sterile PBS to remove the lysis buffer. Centrifuge at 180 × g for 5 min.

• 2.5.7 Repeat the washing step with 10 mL of PBS. NOTE: Aspirate and discard the viscous layer using a 9-inch disposable glass pasture pipette. Care must be taken to not lose the SVF pellet during the two washing steps.

• 2.5.8 Dissolve the pellet in 4 mL complete culture medium. Pipette up and down several times using 1 mL pipette tips with cut ends to obtain a homogenous cell suspension.

• 2.5.9 Plate the cell suspension containing the ASCs in 75 cm² cell culture flasks.

• 2.5.10 Change medium on the next day. Thereafter, change medium every 2–3 days until ASCs become 80–90% confluent. NOTE: If it is required to deplete macrophages from ASCs, follow same procedure as described under Chapter 1.6 for BM-MSCs.

• 3. Trypsinizing and plating the population of confluent BM-MSCs and ASCs

• Once confluent, both the macrophage depleted BM-MSCs and the ASCs must be trypsinized using the following protocol.

• 3.1 After the cells have reached 80–90% confluency, aspirate the culture medium containing the non-adherent or dead cells using a 9-inch disposable glass pasture pipette.

• 3.2 Carefully rinse the cell layer in the flask using 10 mL of PBS pre-warmed at 37⁰C. Repeat this washing step.

• NOTE: The washing step is important as the serum in the medium inhibits trypsin activity, thus complete removal of serum containing medium is necessary.

• 3.3 Aspirate the PBS and then add 2 mL of 0.05% trypsin containing 0.53 mM EDTA per 75 cm² flask. Incubate at 37°C in a cell incubator for 3 min to optimize trypsin activity.

• NOTE: Ensure that the monolayer is thoroughly covered with trypsin.

• 3.4 Moderately tap the flasks against the palm of the hand to dislodge the cells. Monitor the process under the microscope where one can see the trypsinized cells floating. Alternatively, hold the flask against the overhead light in a vertical position and look for the cell sheet coming off from the surface.

• NOTE: If the monolayer is not dislodged, incubate again at 37°C and observe the flask in 3–5 min for cell dissociation.

• 3.5 At this point, it is important to add at least 3 times the volume of the complete culture medium (6 mL). Serum present in the medium will inactivate trypsin.

• 3.6 Rinse the surface of the flask by pipetting the medium containing the dislodged cells multiple times to detach all the remaining cells. NOTE: It is essential to aspirate the cells well to obtain individual, well dispersed cells. If the cells are not well separated, the new culture will contain many microcolonies.

• 3.7 Transfer the medium containing the cells to a 15 mL sterile centrifuge tube and centrifuge at 180 × g for 5 min. 3.8 Add 10 mL of the complete culture medium to the pellet and pipette it up and down several times using a 1 mL cut pipette tip to obtain a homogenous cell suspension.

• 4. Cell counting

• Viable BM-MSCs and ASCs can be counted using trypan blue and a haemocytometer, among other methods.

• 4.1 Mix an aliquot of 10 µL cell suspension and 10 µL of 0.4% (w/v) trypan blue for the viability count. NOTE: Generally greater than 95% viability is observed after the isolation procedures. If more than 10% of the cells are not viable, this indicates that the procedure did not work optimally, which requires troubleshooting.

• 4.2 Add 10 µL of the above cell suspension mixture to one side of the haemocytometer. Count the total number of cells per mL and calculate the total yield of cells per mouse. For seeding BM-MSCs and ASCs intended for Oil Red staining, it is ideal to use 20,000 cells/well of a 96-well plate.

• 5. Phenotypic characterization

• If required, the harvested BM-MSCs and ASCs can be analysed and validated for the appropriate cell surface markers by flow cytometric analysis.

• 5.1 Transfer 10 × 104 BM-MSCs or ASCs in a 2 ml tube.

  5.2 Centrifuge the cells at 500 × g for 5 min at 4°C.

• 5.3Carefully remove the supernatant using a transfer pipette.

• 5.4 Disperse the cell pellet in 100 µL of Fc blocking antibody (99 µL of isolation buffer with 1 µL of purified anti-mouse CD16/32 antibody) and mix gently. Incubate at 4°C for 10 min.

• 5.5 Prepare a master mix containing all required antibodies with appropriate fluorescent dyes (Table S1). There are many products on the market. Examples of MSC markers are anti-CD29 eFluor450, anti-CD44 PE, and anti-CD90.2 PE-Cyanine 7, and examples of haematopoietic stem cell markers are anti-CD34 AlexaFluor 700 and anti-CD45 FITC. Add 3 µL of each antibody to 100 µL of isolation buffer.

• NOTE: Switch off the light to protect the respective fluorophores.

• 5.6 Add 100 µL of antibodies master mix. Mix the samples well. Place on ice for 45 min and cover with foil to protect from light.

• 5.7 Centrifuge at 500 × g for 5 min at 4°C. Remove the supernatant using transfer pipette and resuspend the cells in isolation buffer.

• 5.8 The cells are then ready for analysis by flow cytometry (Table S1) (example: and FlowJo software for data analysis).

• 6. Differentiation of BM-MSCs and ASCs into adipocytes

• Schematic of the critical time points of the adipogenic induction and maintenance is provided in Figure 6. Representative results of the differentiated and non-differentiated BM-MSCs and ASCs are shown in Figure 7.

• Here, we use a four-component cocktail for adipogenic induction which consists of insulin, dexamethasone, IBMX, and indomethacin (concentrations below).

• NOTE:

• a) Care must be taken while changing the medium. The plates must be tilted at an angle of 45⁰ to avoid disruption of the cell monolayer. 20000 cells per well are seeded in a 96-well plate. Once the cells are seeded, the time point is considered as day 0.

• b) It is advisable to seed the cells in triplicates or quadruplicates to account for variability of the results.

• 6.1 On day 1, the adipogenic induction medium is prepared for both BM-MSCs and ASCs by mixing the complete medium with the adipogenic components at the following concentrations: 10 µg/mL insulin (17 µM), 1 µM dexamethasone, 500 µM IBMX and 100 µM indomethacin. Refer to Supplemental Table 2 for the stock concentration and the solvents for each component.

• 6.2 On day 3, switch from adipogenic induction medium to adipogenic maintenance medium containing 10 µg/mL insulin and 1 µM dexamethasone.

• 6.3 Change adipogenic maintenance medium on day 5 and 7 (if end point is day 10) and additionally on day 9 (if end point is day 12).

• 6.4 From day 10 to day 12, observe under inverted microscope (100× or 200× magnification) that cells have accumulated maximum lipid droplets and display a mature adipocyte phenotype. Day 10–12 is the experimental endpoint.

• GENERAL NOTE: The differentiated adipocytes can be used for any further experimental setup, such as immunofluorescence staining, quantification of mRNA by quantitative polymerase chain reaction, Western blotting after protein extraction, RNA sequencing, as well as metabolomic and genomic experiments. Below we describe Oil Red O staining as a detection method of mature adipocytes.

• 7. Oil red O staining

• 7.1 Preparation of stock solution

• 7.1.1 Weigh 0.7 g Oil Red O powder and dissolve it in 200 mL 100% (v/v) isopropanol.

• 7.1.2 Stir the dye overnight at 4⁰C to dissolve it completely.

• 7.1.3 Next day, filter the solution using a Whatman grade 3 MM chromatography paper.

• NOTE: This stock solution can be stored at 4°C and can be used for about one year.

• 7.2 Preparation of working stock

• 7.2.1 Dilute 6 parts of Oil Red O stock solution with 4 parts of distilled water.

• 7.2.2 Allow it to stand for 20 min at ambient temperature. This ensures that the small remaining particles settle down.

• 7.2.3 Fine filter the working solution using a 0.45 µm non-sterile filter and cover immediately.

• NOTE: Working solution must be prepared fresh each time because of reduced solubility of Oil Red O in water.

  7.3 Oil Red O staining procedure

• 7.3.1 Aspirate the medium from the wells and wash each well once with 200 µL PBS.

• 7.3.2 Add 100 µL of 10% (v/v) formalin (~ 4% formaldehyde) to each well containing the cells and incubate at ambient temperature for 1 h to fix the cells. NOTE: One can stop the procedure at this stage. Cells can be stored in 10% (v/v) formalin for several days before proceeding with the staining steps. For this interruption, wrap the 96-well plate with all-purpose laboratory film (example: parafilm) to prevent it from drying.

• 7.3.3 Remove the formalin.

• 7.3.4 Wash each well gently once with 200 µL of 60% (v/v) isopropanol for 2–5 min.

• 7.3.5 Allow the wells to dry completely.

• NOTE: This is a critical step as any residue of isopropanol will interfere with the Oil Red O staining.

• 7.3.6 Add 100 µL of Oil Red O working solution and incubate at ambient temperature for 15 min.

• 7.3.7 Remove the Oil Red O working solution and wash the wells with four exchanges of distilled water for 15–20 s each.

• 7.3.8 Cover the cells with 100 µL of distilled water and analyse the sample promptly (see Chapter 8).

  NOTE: If desired, counter stain with haematoxylin for 1 min and capture the images using an inverted microscope. This step is skipped if quantification will be performed as outlined below.

Figure 6.

Timeline of adipogenic induction and maintenance in cells culture. BM-MSCs or ASCs were seeded on day 0. On day 1, the cells were treated with adipogenic induction medium consisting of 10 µg/mL (17 µM) insulin, 1 µM dexamethasone, 500 µM IBMX, and 100 µM indomethacin. This treatment resulted in the formation of preadipocytes during the commitment phase. On days 3, 5 and 7, the cells were treated with adipogenic maintenance medium consisting of 10 µg/mL (17 µM) insulin and 1 µM dexamethasone. During this expansion phase the cells differentiate into early and mature adipocytes. On day 10–12, analysis of differentiated adipocytes can be performed (for example oil red O staining to visualize and quantify lipid droplets).

Figure 7.

Analysis of differentiated BM-MSCs and ASCs by oil red O staining. Freshly isolated or freeze/thawed macrophage-depleted BM-MSCs and ASCs differentiation in the presence of the adipogenic induction and maintenance cocktail (see Figure 6) was assessed on day 10 using oil red O staining and compared to non-differentiated cells without the adipogenic cocktail. Cells were imaged at 200× magnification using an inverted microscope. The scale bar represents 100 µm. Data are presented as means ± SE, n = 3–5 mice per group, *p < 0.05, assessed by one-way ANOVA with Bonferroni post-test. Oil red O staining was quantified as detailed in the protocol and showed significant differences between the treated and non-treated cells. Freshly isolated and freeze-thawed cells were similarly potent in adipogenic differentiation efficiency.

• 8. Quantification of BM-MSCs and ASCs

  8.1 Lipid bodies in mature adipocytes will be stained bright red. Take at least five representative images for each well using an inverted microscope under 200× magnification.

  8.2 The differentiated adipocytes stained with Oil Red O can be quantified as described below:

  8.2.1 Remove the water and allow the wells to dry completely.

  8.2.2 Dissolve the Oil Red O stain by adding 100 µL of isopropanol containing 4% IGEPAL CA 630 which is a non-ionic detergent.

  8.2.3 Incubate the plates on the orbital shaker at 500 rpm for 15 min.

  8.2.4 Pipette up and down several times to ensure the homogenous solubilization of Oil Red O. Measure the absorbance at 520 nm using a microplate reader.

Results and discussion

Isolation of murine BM-MSCs and ASCs

We have provided comprehensive steps for harvesting BM-MSCs and ASCs from mice. Figure 1 depicts the isolation procedure for BM-MSCs from the tibiae and femurs of the mice. Figure 2 demonstrates the customized tube preparation required to flush the bone marrow from the tibiae and femurs of mice. Figure 3 illustrates the procedure for isolating ASCs from various murine adipose tissue depots (subcutaneous, gonadal and retroperitoneal). It is widely appreciated in the literature that the proliferation and differentiation potential of stem cells decline with age. Chaudhary et al. demonstrated that proliferation was higher in cultures of MSCs isolated from young mice (2–3 months) than in those isolated from aged mice (23–24 months) [24]. As per our experience, the most appropriate age of mice for the extraction of BM-MSCs is between 4–6 weeks and that of ASCs is between 6–8 weeks. If one aims to extract both BM-MSCs and ASCs from the same mice, we suggest using 6-week old mice as it allows efficient harvesting of MSCs from both sources. To ensure that the functional status of the isolated stem cells is maximally preserved, it is essential to process the harvested bone marrow and adipose tissue quickly, ideally within 30 min. Several methods have been described for harvesting bone marrow from bones. Bone marrow can be efficiently removed from long bones by either flushing the bones with a liquid-filled syringe, or by applying a centrifugal force to the bones [25,26]. We favour and describe here the centrifuge-based method as it is very convenient, considerably faster, and results in increased total bone marrow recovery [26].

Characterization of mouse BM-MSCs and ASCs and differentiation into adipogenic lineage

Freshly harvested macrophage-depleted BM-MSCs as well as ASCs proliferated efficiently, gradually expanded in cell size and reached confluency at ~ 5–6 d (ASCs) or at ~ 10–12 d (BM-MSCs). We analysed and validated both freshly isolated cells and cells that were first frozen in liquid nitrogen and then thawed. Both freshly harvested and frozen/thawed BM-MSCs and ASCs showed a typical spindle-shaped fibroblast-like appearance (Figure 4). The presence of macrophages was apparent in BM-MSC cultures that were not depleted for macrophages, and these BM-MSCs proliferated less than the macrophage-depleted BM-MSCs (Supplemental Figure 1A,B). To analyse typical stem cell markers, we performed flow cytometry using antibodies against MSC markers CD29, CD44, and CD90.2, and haematopoietic stem cell markers CD34 and CD45 (Figure 5). Both freshly harvested and frozen/thawed macrophage-depleted BM-MSCs were strongly positive for CD44 and CD29 but negative for CD34 and CD45 (Figure 5A,B). As expected, non-macrophage-depleted BM-MSCs were positive for CD34 and CD45 (Supplemental Figure 1C). Thus, the flow cytometric analysis demonstrated effective macrophage depletion to an extent that we do not expect an effect of any residual macrophages on adipogenic differentiation and function. Similarly, freshly isolated and frozen/thawed ASCs were strongly positive for CD29, CD44, and CD90.2 but negative for CD34 and CD45, indicating the absence of macrophages (Figure 5C,D). Therefore, macrophage depletion was not required for the ASC preparations. The data clearly demonstrate that the described cell harvesting procedures yield the expected mesenchymal stem cells from the bone and fat tissue. The results also showed that the isolation method produced cells that were sufficiently robust to be frozen and thawed without losing their characteristics and stemness. Other methods to isolate BM-MSCs and ASCs include fluorescence-activated cell sorting after specific cell surface marker labelling, but small amounts of MSCs present in bone marrow or adipose tissue can make harvesting of sufficient amounts of cells challenging [27].

Figure 4.

Microscopic images of isolated BM-MSCs and ASCs. Macrophage-depleted BM-MSCs (A,B) and ASCs (C,D) are shown either directly after isolation (A,C) or after one freeze-thaw cycle (B,D). Note the typical spindle-shaped morphology. The magnification was 100 × . The scale bar represents 200 µm.

Figure 5.

Analysis of mesenchymal stem cells markers for BM-MSCs and ASCs using flow cytometry. Flow cytometry analysis of freshly isolated (A,C) and frozen/thawed (B,D) macrophage-depleted BM-MSCs (A,B) and ASCs (C,D) for expression of MSC markers CD29, CD44, and CD90.2, and haematopoietic stem cell markers CD34 and CD45. Freshly isolated and frozen/thawed BM-MSCs and ASCs were negative for CD34 and CD45 and positive for CD29 and CD44. ASCs were also positive for CD90.2. Unstained cells were used as negative control cell populations for freshly isolated and frozen/thawed BM-MSCs as well as ASCs and are represented by green peaks. The respective marker-positive cell populations are indicated in red for BM-MSCs and cyan for ASCs. The x-axis shows the fluorescence intensity, and the y-axis represents the cell count.

In vitro differentiation capacities of freshly isolated and frozen/thawed BM-MSCs and ASCs

Adipogenic differentiation is a complex process that involves a transcriptional cascade of several transcription factors. For example, CCAT/enhancer binding protein (C/EBP) and PPAR family of transcription factors play important roles in adipogenic differentiation [21]. The exposure of MSCs to adipogenic stimuli is necessary to induce adipogenic differentiation. Three-component cocktails (insulin, dexamethasone, and IBMX) and four-component cocktails, including additionally indomethacin, are commonly used to induce adipogenesis in MSCs [16,20,22]. However, the concentration of each component in the adipogenic cocktails varies greatly between studies. In this protocol, we used an optimized four-component cocktail consisting of 10 µg/mL (17 µM) insulin, 1 µM dexamethasone, 500 µM IBMX, and 100 µM indomethacin supplemented 1 d after cell seeding until day 3. This was followed by adipogenic maintenance medium consisting of 10 µg/mL (17 µM) insulin and 1 µM dexamethasone supplemented on days 3, 5 and 7 (Figure 6). Insulin induces proliferation and differentiation of preadipocytes. It is the most potent inducer of adipogenic differentiation that has a strong anti-lipolytic action and stimulates lipogenesis which triggers and promotes adipocyte differentiation in vitro [28]. Dexamethasone stimulates both adipogenic and osteogenic differentiation in a cell-, time-, and concentration-dependent manner [29]. IBMX is a competitive, non-selective phosphodiesterase inhibitor that increases intracellular cAMP levels, which in turn elevates protein kinase A (PKA) activity. PKA signalling is required for the transcriptional activation of PPAR-γ and, thus, adipogenic gene expression [30]. Indomethacin is a ligand of PPAR-γ, which is a master regulator of adipogenesis. Indomethacin promotes adipogenesis in stem cells in vitro and upregulates the expression of adipogenic genes [31]. This is supported by data from Styner et al., who demonstrated that indomethacin accelerated adipogenesis in MSCs [32].

Freshly isolated and frozen/thawed BM-MSCs and ASCs were differentiated using an optimized four-component adipogenic induction medium in combination with a two-component maintenance medium. Adipocyte differentiation was assessed after 10 d using Oil Red O staining (Figure 7). Freshly isolated and frozen/thawed macrophage-depleted BM-MSCs cultivated in complete culture medium without the adipogenic cocktail were not stained with Oil Red O, whereas BM-MSCs treated with adipogenic cocktail showed large Oil Red O stained lipid-rich vacuoles, indicating effective adipogenic differentiation (Figure 7A,B). BM-MSC preparations without macrophage depletion showed relatively similar adipogenic differentiation properties under these experimental conditions (Supplemental Figure 1D). Similarly, freshly isolated and frozen/thawed ASCs treated with the adipogenic cocktail were strongly stained with Oil Red O compared to the untreated control, demonstrating effective adipogenic differentiation (Figure 7C,D). Differentiated ASCs resulted in the typical smaller differentiated adipocytes compared to those originating from the BM-MSCs. We recently used the described adipogenic differentiation scheme for murine BM-MScs and identified that the cells differentiated into the adipogenic lineage as evidenced by the upregulation of adipogenic markers including CEBPα, PPAR-γ and fatty acid binding protein [33].

In addition to the described differentiation cocktails, there are a variety of other suitable culture media available for adipogenic differentiation, such as alternative standard cell culture media [29] or various proprietary media formulations with undisclosed components. In this study, we used MEMα medium supplemented with heat-inactivated FBS, penicillin, streptomycin, and glutamine as it is an inexpensive culture medium that can be tailored with the addition of adipogenic components at optimized concentrations. Using proprietary media can be convenient but is associated with two disadvantages: i) they are typically much more costly than the culture medium mentioned above, and ii) the lack of knowledge of some or all of the components imposes difficulties in the standardization of the experimental procedure and interpretation of the data.

Conclusion

This protocol details hands-on experimental steps to effectively harvest murine BM-MSCs and ASCs that have full potential, either freshly isolated or after freeze/thawing, to effectively differentiate into adipocytes using a cost-effective optimized four-component adipogenic differentiation cocktail. The protocol also explains the steps to deplete macrophages from the MSC preparations and validate the quality of the extracted BM-MSCs and ASCs. This protocol provides a useful laboratory tool for establishing or optimizing MSC harvesting and differentiation into adipocytes.

Funding Statement

The work was supported by the Canadian Institutes of Health Research [PJT-479415].

Disclosure statement

No potential conflict of interest was reported by the author(s).

Data availability statement

The data that support this study are openly available in Figshare at https://doi.org/10.6084/m9.figshare.24999746.

Author contributions

IFSS wrote the manuscript, performed the experiments, and designed the figures. MLM performed experiments and edited the manuscript. DPR conceived the project and edited the manuscript and the figures.

Animal ethics

All experimental procedures were approved by the McGill University Animal Care Committee in accordance with the guidelines of the Canadian Council on Animal Care (Protocol #7561). The authors have nothing to disclose.

Supplementary materials

Supplemental data for this article can be accessed online at https://doi.org/10.1080/21623945.2024.2350751