Isolation of Preadipocytes from Broiler Chick Embryos

All animal procedures were approved by the University of Tennessee Institutional Animal Care and Use Committee. Freshly fertilized commercial broiler eggs (Cobb 500) were obtained from a local hatchery. Eggs were incubated at 38 °C with 60% relative humidity until dissections at embryonic days 16-18 (E16-E18). Adipose tissue was collected from the subcutaneous (femoral) depot.

1. Preparation for isolation and culture

1. Prepare the culture hood and the instruments.
   1. Before starting dissections, set up a work area in the laminar-flow hood. Disinfect the working area and all the instruments by swabbing with 70% ethanol. Perform all procedures using sterile materials.
      NOTE: Always swab the containers and instruments with 70% ethanol before placing them back in the cell culture hood. It is recommended to place a benchtop instrument sterilizer in the hood so that instruments can readily be sterilized between embryos.
      CAUTION: When using an instrument sterilizer, cool down the hot instrument properly to prevent burn injury and tissue damage. A minimum cooling time of 3 min is recommended. Swab with 70% ethanol before use.
   2. Assemble the following instruments and vessels in the hood: straight forceps (120 mm), tweezers (110 mm), two pairs of curved forceps (100 mm), two pairs of straight scissors (140 mm), curved surgical scissors (115 mm), tissue strainer (250 µm nylon mesh), cell strainer (40 µm nylon mesh), conical centrifuge tubes (15 mL and 50 mL), Petri dishes (60 mm and 100 mm), small beaker (100 mL), 70% ethanol spray and sterile gauze, paper towel, and benchtop wiper (see Table of Materials).
2. Prepare enzymatic solution, collection media, and culture media following the steps below.
   NOTE: Prepare solutions in advance and store at 4 °C until use. Media needs to be placed on ice before tissue collection. All reagents used in this step are listed in the Table of Materials.
   1. Prepare media used for both collection of adipose tissue and preparation of enzymatic solution by supplementing DMEM/F12 (Dulbecco's Modified Eagle Medium with 2.50 mM of L-glutamine and 15 mM of HEPES buffer) with 2.5 µg/mL of Amphotericin B and 1x penicillin/streptomycin (P/S), 100 U/mL. Store at 4 °C until use.
      NOTE: This media remains stable for 1 year when stored at 4 °C.
   2. Prepare sterilization solution by diluting Betadine to 20% (v/v) in 1x PBS (Phosphate Buffered Saline having pH 7.4, without calcium, magnesium, or phenol red) with 2.5 µg/mL of Amphotericin B. Store at 4 °C until use.
      NOTE: The solution is stable for 2 years when stored at 4 °C.
   3. Prepare enzymatic solution by dissolving Type 1 collagenase (1 mg/mL) in DMEM/F12 with 2.5 µg/mL of Amphotericin B and 1x P/S (step 1.2.1). Make fresh collagenase solution and maintain it on ice during dissections. Approximately 10 min before use, pre-warm by incubating this solution at 37 °C in a water bath to initiate its enzymatic activity.
      NOTE: Approximately 1 mL of collagenase solution (1 mg/mL) is needed per 100 mg of adipose tissue.
   4. Prepare growth media used for plating and propagation by supplementing DMEM/F12 media with 1x P/S and 10% FBS. Store at 4 °C until use.
      NOTE: The solution is stable for 1 year when stored at 4 °C.
   5. Prepare washing solution by supplementing 1x PBS with 2.5 µg/mL of Amphotericin B. Adjust solution to desired pH 7.4. Store at 4 °C until use.
      ​NOTE: The solution is stable for 2 years when stored at 4 °C.

2. Adipose tissue collection and digestion

1. Euthanize the embryo.
   1. On embryonic day 16, remove the eggs from the incubator. The primary potential source of microbial contamination is the egg's surface; therefore, swab the eggs with a sterile gauze soaked in 70% ethanol prior to cracking them. After swabbing, place the egg vertically with the pointy end down onto a small beaker (100 mL) lined with paper towels to cushion the egg from the glass.
   2. Using the forceps' handle, break an egg by tapping the blunt end of the egg. Carefully remove the eggshell to create an opening sufficiently large to remove the embryo (step 2.1.3). Gently tear the white shell membrane to expose the embryo (Figure 1A). Pierce the amnion carefully using sterile tweezers (Figure 1B).
      NOTE: The amniotic sac is a transparent membrane filled with amniotic fluid that encloses the embryo.
   3. Remove the embryo from the egg by lightly gripping the embryo's neck using straight forceps. Sever the yolk sac to disconnect it from the embryo, and transfer the embryo to a 100 mm Petri dish.
   4. Decapitate immediately using surgical scissors and forceps.
2. Perform the adipose tissue collection.
   1. Swab the embryo body with 70% ethanol and scrub the skin surface gently with a sterile gauze to remove feathers, as they can interfere with filtration at later steps after digestion. Use benchtop wipers to keep the skin and feathers from touching collected tissue.
   2. Cut off the skin between the legs and abdominal region to reveal the pair of femoral adipose depots.
   3. Hold the skin around the leg using curved forceps with one hand. Gently remove the femoral subcutaneous fat with the other hand, using curved forceps to gently pull the depot away from the leg.
      NOTE: There is relatively little connective tissue that adheres to the fat pad to the leg, and the entire fat pad should be removable in one piece using only forceps. This can be facilitated by holding the forceps backward so that the curved portions (rather than the ends) clamp the fat pad for removal.
      1. If necessary, cut the fat pad away with curved scissors. Repeat with the other fat pad and additional embryos as needed.
         NOTE: A total of 80 mg of subcutaneous fat can be obtained from most embryos at E16 (Figure 1C). This typically yields ~1 x 10⁶ cells, sufficient to plate one T-25 flask. It might be useful at this step to weigh fat pads from a few embryos to assess the amount of starting material, as fat pad weights can vary across specific broiler lines, and due to uncontrollable factors, such as breeder hen age and diet. Subcutaneous fat was routinely collected from five eggs to ensure an adequate yield of cells for plating in multiple flasks.
   4. Transfer the tissues into ~5 mL of collection media in a 15 mL tube and repeat dissection for the remaining eggs.
   5. Briefly rinse the collected fat pads by transferring them to a 60 mm Petri dish containing sterilization solution and swirling the dish a few times.
   6. Rinse off the sterilization solution by transferring fat pads to a 60 mm Petri dish containing 1x PBS. Swirl gently. Repeat this step by transferring to a second 60 mm Petri dish containing 1x PBS.
3. Perform enzymatic digestion and preadipocyte isolation following the steps below.
   1. Transfer the adipose tissues to a 15 mL tube containing ~1 mL of pre-warmed enzymatic solution per 100 mg of tissue. Immerse a pair of long, straight scissors in the tube and finely mince the adipose tissues in the solution into as small pieces as possible (~1 mm³) (Figure 2A).
      NOTE: Cell yield will be reduced if the tissues are not thoroughly minced.
   2. Transfer the minced tissue and enzymatic solution to a 25 mL autoclaved flask. Wrap the flask with paraffin film. Place on an orbital shaker inside an incubator and shake at 37 °C during the digestion step.
      NOTE: Alternatively, use a shaking water bath. With either approach, the speed should be sufficient to prevent pieces of tissue from settling to the bottom of the flask, but must not be so fast that pieces are propelled to the top of the flask and stick to the glass, or that fluid accumulates on the paraffin film cover.
   3. After ~30 min, cut the end of a 1 mL pipette tip to a diameter of approximately 3 mm. Pipette the tissue mixture up and down a few times to help release the cells from the adipose tissue. Return the flask to the incubator/water bath and resume shaking.
   4. After an additional 15 min, check the flask for completeness of digestion. After removing the flask from the shaker, a layer of whitish cells will form at the top of the fluid layer. If tissue fragments still remain, cut the end from another pipette tip and gently pipette the mixture up and down, then continue shaking for an additional 15 min.
      NOTE: Completely digested tissue/collagenase mixture looks like milky chyme. Typically, tissue is sufficiently digested after 1 h of gentle shaking. If many fragments remain at this point, it may indicate an issue with the collagenase enzyme used. Constant shaking can increase yield; however, prolonged exposure to the enzyme and the physical stress may also damage the cells.
   5. After digestion, pipette gently up and down to mix well. Filter through a 250 µm tissue strainer into a 15 mL tube by pipetting to remove any bits of undigested tissue and debris.
      1. Rinse the flask with 4 mL of growth media by pipetting to remove cells that may be adhered to the glass, and filter into the same 15 mL tube. Rinse the strainer with additional growth media by pipetting to loosen any trapped cells, up to a total volume of 14 mL.
   6. Pellet the cell fraction by centrifuging at 300 x g for 5 min at RT (7 min for 50 mL tube).
      NOTE: Always swab the tubes with 70% ethanol before returning to the cell culture hood.
   7. Aspirate the supernatant. Be careful not to dislodge the cell pellet. Gently resuspend the pellet in 1 mL of red blood cell lysis buffer (see Table of Materials) by pipetting up and down, and incubate for 5 min at RT. The solution will turn reddish as red blood cells are lysed (Figure 2B).
      NOTE: Place the RBC lysis buffer at RT before use. Chickens have nucleated red blood cells⁹, and they attach to tissue culture dishes along with preadipocytes. RBC lysis buffer is used to lyse these cells to prevent their interference with accurate cell counts.
   8. Add 5 mL of growth media to the tube containing the cells to dilute the lysis buffer and mix gently by pipetting up and down. Using a pipette, filter through a 40 µm cell strainer into a new 50 mL tube and rinse the strainer with an additional 5 mL of growth media.
   9. Pellet cells by centrifuging at 300 x g for 7 min at RT. Carefully aspirate the supernatant and resuspend the remaining cell pellet in 1 mL of growth media by pipetting up and down.
      1. Use a hemocytometer, cell counter, and Trypan Blue stain to count cells and determine cell viability. Take 10 µL of sample and mix with 10 µL of Trypan Blue by pipetting. Load 10 µL of the mixture on to the hematocytometer and measure¹⁰.
         ​NOTE: Centrifugation speeds can be increased to 600 x g if sufficient cell pellets are not readily visible after the initial spin.

3. Seeding and culture of preadipocytes

1. Seed ~1 x 10⁶ cells in 4 mL of pre-warmed growth media in a T-25 flask. Follow the same plating density if using other types of culture vessels. Place in tissue culture incubator and allow to attach overnight.
   NOTE: Cells are cultured in a 38 °C incubator with a humidified atmosphere of 5% CO₂. The optimal temperature for the growth of avian cells¹¹ is 38 °C, and they grow slowly at 37 °C.
2. Next day, aspirate media and gently wash the cells with 1x PBS by pipetting to remove unattached or dead cells. Replace with 4 mL of fresh growth media. Check the cells under a microscope. Cells should be spindly shaped, like fibroblasts (Figure 2A).
   ​NOTE: Typically, preadipocytes attach fairly quickly (within a few hours). The success or failure of cell isolation can be confirmed at this time (after 24 h).
3. Replace with 4 mL of fresh growth media every 2 days and subculture or cryopreserve cells when they reach 70%-80% confluence (Figure 3C).

4. Subculturing and cryopreservation

1. Aspirate old media and gently wash cells with 4 mL of 1x PBS by pipetting. Aspirate PBS, add 2 mL of 0.1% trypsin to cover the cell surface in a T-25 flask (adjust the volume accordingly for other culture vessels), and then incubate for 3-4 min at 38 °C.
   NOTE: Observe if the cells are detached from the culture plate. Tap the culture plate gently to help cell detachment. Incubating cells with trypsin for too long will damage cells.
2. Add an equivalent volume of pre-warmed growth media to inhibit the trypsin reaction. Pipette over the cell surface several times, tilting the plate to loosen the remaining cells.
3. Transfer the cell suspension to a 15 mL tube and centrifuge at 300 x g for 5 min at RT (7 min for 50 mL tube). Aspirate the supernatant.
4. If subculturing, resuspend pellet in 1 mL of growth media by pipetting up and down. Count cells as described in step 2.3.9.1 and then replate using the same plating density used initially in step 3.1.
5. If cryopreserving, prepare 4 mL of freezing media for a T-25 flask with 90% confluency.
   NOTE: Freezing media consists of 10% DMSO, 30% FBS, and 60% DMEM/F12.
6. Resuspend cell pellet in freezing media and transfer 1 mL of freezing media into a cryovial. Freeze the cryovials slowly to -1 °C/min using a freezing container. Place the container in a -80 °C freezer overnight and then transfer it to liquid nitrogen for long-term storage.
   ​NOTE: Properly cryopreserved preadipocytes maintain their viability for at least 3 years and typically perform like freshly isolated cells when thawed and plated.

5. Adipogenic differentiation

NOTE: 2% gelatin-coated plates can be used to enhance cell adhesion.

1. Prepare a 2% (w/v) gelatin solution (see Table of Materials) in distilled water. Autoclave at 121 °C, 15 psi for 30 min to sterilize. Coat culture surface with 5-10 µL of gelatin solution/cm² (i.e., 100-200 µg/cm²). Swirl gently to evenly coat the surface.
2. Check plates for even spreading of the gelatin solution since some regions may remain uncoated initially. Allow the gelatin-coated plate to remain at RT for at least 1 h. Remove the entire volume of gelatin solution from the wells.
   NOTE: This will leave a thin gelatin coat at the bottom of the wells/dishes. Gelatin solution can be reused multiple times (at least 10 times) without altering cell adhesion and growth. Allow the gelatin-coated dishes to remain in the tissue culture hood for at least 30 min before plating the cells.
3. Induce the chicken preadipocytes to undergo adipogenic differentiation by supplementing growth media with fatty acids. To prepare adipogenic differentiation media (ADM), supplement DMEM/F12 with 10% chicken serum, 1x Linoleic Acid-Oleic Acid-Albumin (9.4 µg/mL), and 1x P/S (see Table of Materials). Make ADM fresh and warm before use.
   ​NOTE: Chicken preadipocytes are commonly induced to undergo adipogenic differentiation by supplementing media with fatty acids rather than hormonal cocktails typically used for adipocytes from other species¹².
4. Induce differentiation by replacing growth media with adipogenic differentiation media when cells reach ~90% confluence. Maintain cells in this media, replacing them every 2 days. Assess the differentiation visually under a microscope (20x) based on the formation of lipid droplets, which become readily visible within 48 h of inducing differentiation.

6. Assessing adipogenesis

1. Perform Oil Red O staining following the steps below.
   1. Plate the cells in a six-well plate and induce adipogenic differentiation at ~90% confluency.
   2. Prepare a working solution of Oil Red O by combining six parts of the Oil Red O stock solution with four parts of distilled water in a 50 mL tube. Gently mix by pipetting up and down and let stand for 10 min at RT, and then filter through a Grade 1 filter paper (see Table of Materials) inside the funnel by slowly pouring the solution into a 50 mL tube.
      NOTE: Oil Red O stock solution is made by dissolving 0.7 g of Oil Red O (see Table of Materials) in 200 mL of 100% isopropanol in an autoclaved bottle. Mix well and let it sit for 20 min. Store at 4 °C and keep away from light until use. This is stable for 1 year. The working solution can be used for 3 h; however, it is recommended to use it within 2 h.
   3. Remove media and gently wash the wells twice with 2 mL of pre-warmed 1x PBS using a pipette. Remove PBS completely by pipetting off. Fix cells with 2 mL of 10% buffered formalin and wrap the plate with paraffin film. Leave at RT for at least 1 h and up to 2 days before staining.
      NOTE: To make 1 L of 10% buffered formalin solution, mix 100 mL of 37% formaldehyde, 4.09 g of NaH₂PO₄, 6.5 g of Na₂HPO₄ (see Table of Materials), and 900 mL of distilled water. Do not pipette formalin directly onto the cells. Dispense gently on the sidewall near the bottom of the well.
   4. Remove formalin and gently wash the wells with 2 mL of distilled water. Replace with 2 mL of 60% isopropanol. After 5 min, remove isopropanol and let the wells dry completely for about 10 min. Perform all staining steps at RT.
   5. Add 1 mL of Oil Red O working solution to cells and incubate for 10-20 min at RT. Remove stain by pipetting off and wash five times by dipping in tap water until no excess stain is seen. After the final rinse, add 1 mL of water to cells prior to visualizing staining and collecting images under a microscope.
   6. To quantify lipid accumulation per dish, remove water and extract Oil Red O dye from cells by adding 1-2 mL of 100% isopropanol that is sufficient to cover cells in a well of six-well plate completely. Incubate with gentle shaking on a plate shaker for 10 min at RT.
   7. Transfer 200 µL of the extraction into a well of the 96-well assay plate. Quantify the relative amount of stain using a spectrophotometer plate reader to measure absorbance at 495 nm.
2. Perform the staining assay of the intracellular lipid droplets and the nucleus.
   1. Plate cells in black bottom 96-well plates and induce adipogenic differentiation as described in step 5.3.
   2. To stain the cells, add 200 µL of the staining solution containing a fluorescent lipid stain and a fluorescent DNA stain (see Table of Materials) in 1x PBS per well. After calculating the total volume of the stain required, add two drops of the DNA stain and 25 µL of the lipid stain per mL of pre-warmed 1x PBS using a foil-wrapped tube to protect the solution from light. Incubate at RT for 20 min and protect from light.
   3. Read the fluorescence using a fluorescent plate reader (see Table of Materials). Detect the lipid stain (Excitation: 485 nm/Emission: 572 nm) using a red filter and the DNA stain (Excitation: 359 nm/Emission: 450 nm) through a blue/cyan filter.
      NOTE: Staining can also be visualized and images captured under a fluorescent microscope.
   4. Normalize the lipid stain intensities to the DNA stain intensities to quantify lipid accumulation relative to cell number¹³.