Isolation, Primary Culture, and Cryopreservation of Human Neonatal Fibroblasts

Objective

A recommended procedure to isolate and establish a primary culture of human neonatal fibroblasts from foreskin tissue under animal origin free (AOF) or serum-containing conditions is described below. Subculture and cryopreservation procedures are also included.

Introduction

Primary human fibroblasts from skin (dermis) are useful for a number of scientific endeavors including the study of growth factor action, wound healing, toxicity/irritancy studies, and use as feeder cells for embryonic stem cells and induced pluripotent stem cells. We offer a complete range of products for the isolation, growth and cryopreservation of these cells in animal origin free conditions or serum-containing conditions. The following protocol describes the isolation of cells from neonatal tissue. Neonatal dermis tissue is more cellular and contains less extracellular matrix than dermal tissue from adult skin. In addition, neonatal cells have an inherently longer lifespan than cells from older individuals. If isolation from adult skin is desired, consider using larger amounts of the starting tissue and increasing the collagenase concentration.

General Guidelines

• Perform all protocols using sterile techniques in a Class II, Type A2 laminar flow hood
• Always wear double gloves, protective eyewear, and a lab coat during isolation procedures
• Use universal precautions when handling human tissue and dispose of contaminated materials appropriately

 

Preparing Tissue

This procedure describes the preparation of fibroblasts from neonatal foreskin tissue. If you are processing larger pieces of tissue, modify the protocol accordingly.

1. To prepare supplemented Fibroblast AOF medium, add the following to one 500 mL bottle of Fibroblast AOF Basal Medium (FABM):

• Defined Fibroblast AOF Supplement - 5 mL
• 100X Antibiotic-Antimycotic Liquid (AA) - 5 mL

To prepare serum-supplemented medium, add the following to one 500 mL bottle of D-MEM medium:

• Fetal Bovine Serum (FBS) - 55 mL
• 100X Antibiotic-Antimycotic Liquid (AA) - 5.5 mL

2. Place an absorbent underpad in the hood.
3. To a sterile 100 mm culture dish, add ~10 mL of the supplemented medium prepared in Step 1.
4. Obtain tissue and place the container with the tissue in the laminar flow hood.
5. Remove the lid from the 100 mm dish and place it upside down in the hood for use in Step 8, below.
6. Using sterile forceps transfer the tissue to the culture dish prepared in Step 3.
7. Wash the tissue by agitating with forceps in the medium contained in the 100 mm dish.
8. Using sterile forceps place and flatten the tissue onto the overturned lid of the 100 mm culture dish, epidermal side down. If the tissue is in a tubular configuration, use small sterile scissors to open the tissue and flatten onto the lid.
9. Trim away any fat and loose fascia using sterile scissors and forceps. To keep the tissue from drying, rinse every few minutes in the medium in the 100 mm dish.
10. After the trimming is complete, cut the tissue into strips approximately 0.5 cm × 1.5 cm using a sterile scalpel.

Dispase Digestion

1. Add 5 mL Dispase Solution to a sterile, 15 mL conical centrifuge tube.
2. Transfer the cut tissue from Step 10, above to the tube containing Dispase Solution. Ensure the tissue pieces are submerged in solution. Cap the tube securely.
3. Remove outer gloves and wipe the outside of the bottle with tuberculocidal solution. Label the tube appropriately.
4. Transfer the tube to a 4°C refrigerator.
5. Incubate the tube for 16–21 hours at 4°C.

Isolating and Plating Dermal Cells

1. After Dispase digestion, retrieve the tube containing the tissue and place the tube in the hood.
2. Obtain a sterile 100 mm culture dish, and remove and place the lid upside down in the hood.
3. Transfer the digested tissue and accompanying Dispase Solution into the bottom of the 100 mm culture dish, avoiding splashing. If any pieces of tissue remain in the bottle, use a sterile 1 mL pipette or sterile forceps and transfer the tissue pieces in the bottom of the100 mm culture dish.
4. Separate the epidermis from the dermis.
   
   
   • Handling a few pieces at a time, move the tissue pieces to the overturned lid of the dish. Orient the strips so the epidermis is facing up.
   • Hold the dermis of the tissue strip with one pair of sterile forceps and the edge of the epidermis with another pair of sterile forceps. Pull/peel the dermis and epidermis apart keeping pieces separated on the same lid. Working quickly, repeat the process for each tissue piece.
     
     
5. To a new sterile, 100 mm culture dish, add 10 mL of supplemented medium
6. Using sterile forceps transfer the separated dermal pieces to the 100 mm culture dish containing medium, keeping the dermal-epidermal interface facing up. To isolate and culture epidermal cells, refer to the Isolation, Primary Culture, and Cryopreservation of Human Keratinocytes protocol.
7. Working with one piece of dermis at a time, place a dermal piece into the sterile lid of the 100 mm tissue culture dish and scrape the dermal-epidermal interface surface firmly and thoroughly with a sterile scalpel blade to reduce the presence of microvasculature. Repeat the process for each piece of dermis.
8. Wash the dermal pieces in the 100 mm dish containing 10 mL of supplemented medium prepared in Step 5 and transfer the pieces to the bottom of a clean dry 100 mm tissue culture dish.
9. Add 10 mL supplemented medium to the dish and mince dermal pieces finely with sterile scissors or a scalpel such that the pieces are small enough to be drawn through the opening of a 10 mL pipette.
10. Transfer the tissue pieces to a 50 mL conical centrifuge tube. Use an additional 10 mL supplemented medium to wash the dish and add the wash medium to the conical tube. Add 10 mL 1,500 U/mL Collagenase Solution to the tube containing the tissue pieces (total 30 mL). Cap tube tightly.
11. Incubate the tube at 37ºC for 1 hour, and swirl the tube vigorously every 15 minutes. At the end of the incubation period, tissue should be almost completely digested and no longer visible. If dermal pieces are still visible after 1 hour, continue incubation, checking every 15 minutes, until dermal pieces are no longer visible (do not exceed 2 hours).
12. At the end of the collagenase digestion, centrifuge the cell suspension at 180 × g for 7–10 minutes.
13. Remove the supernatant from the tube carefully without dislodging the pellet. Remove any remaining supernatant from the pellet with 1,000 µL pipette tip.
14. Add 30 mL supplemented medium to the 50 mL tube and resuspend the cell pellet (it is not necessary to obtain a single cell suspension). Replace and tighten the cap.
15. Centrifuge the cell suspension again at 180 × g for 7–10 minutes.
16. Remove the supernatant from the tube carefully without dislodging the pellet. Remove any remaining supernatant from the pellet with 1,000 µL pipette tip. Aspirate all residual drops from inside of tube wall.
17. Resuspend the pellet in 3 mL supplemented medium.
18. Determine the concentration of viable cells/mL and calculate the culture surface area required for primary culture as described below. Place the remaining cell suspension at 4ºC until needed.
19. Add a 20 µL aliquot of the cell suspension from Step 17 to a sterile tube containing 20 µL Trypan Blue solution and determine the total number of viable cells in the preparation using a hemocytometer. For dermal cells isolated from neonatal tissue, plate 5 × 10³ viable dermal cells/cm².
20. For AOF cultures, coat culture surfaces with Coating Matrix as described below. For serum-containing medium, proceed to Step 21.
    
    
    • Obtain and label the required number of flasks.
    • You need 1 Coating Matrix kit per every 250 cm₂ of surface to be coated.
    • Supplement one bottle of Dilution Medium (50 mL) with the contents of one .tube (0.5 mL) Coating Matrix.
    • Add 5 mL diluted Coating Matrix for each 25 cm₂ flask to be coated
    • Swirl flasks thoroughly to coat the surface of each flask. Incubate at room temperature for 30 minutes.
    • Aspirate the Coating Matrix solution from flasks using a pasteur pipette under vacuum.
      
      
21. Alternatively, Coating Matrix can be added directly to the cell suspension before plating cells without the use of the Dilution Medium. Dilute Coating Matrix 1:100 into the cell suspension.
22. Retrieve the cell suspension from 4°C.
23. Dilute the cell suspension in supplemented medium to yield 5 × 10³ viable cells/cm² in an appropriate volume for the culture surface used. For example, seed a T-75 flask with 15 mL/flask of a 2.5 × 10⁴ cell/mL cell suspension
24. Incubate flasks at 37°C and 5% CO₂.

Primary Culture

After initial seeding of the primary culture, change the medium after 24 hours, and then at least once every 48 hours. Once the cultures are >50% confluent, change the medium daily. Once the cultures are ~90% confluent (7–13 days), subculture or cryopreserve the cells using SAF cryopreservation medium as described below. View the culture under a microscope to ascertain the condition of the culture (i.e., confluence, mitotic activity).


Subculture of Dermal Fibroblasts

This protocol is designed for the subculture of one 25 cm₂ culture flask of actively proliferating cells near confluence. If different-sized culture vessels are to be used, adjust the reagent volumes accordingly.
 
Note:  We do not recommend warming the reagents prior to use.

1. Remove all of the culture medium from the flask.
2. Add 3 mL Ca++ and Mg++-free PBS to the flask. Rock the flask to ensure that the entire surface is covered.
3. Immediately remove the PBS solution from the flask and add 1 mL TrypLE™ solution. Rock the flask back and forth to ensure even coating.
4. Incubate the flask at room temperature for 5 minutes. View the culture under a microscope. When the cells have become partially detached and rounded, gently rap the flask to dislodge the cells from the surface of the flask.
5. Add 4 mL complete medium to the flask and transfer the detached cells to a sterile 15 mL conical tube.
6. Add 4 mL additional complete medium to the flask and pipette the solution over the flask surface several times to remove any remaining cells. Add this solution to the 15 mL conical tube.
7. Centrifuge the cells at 180 × g for 7–10 minutes. Observe the cell pellet.

Note:   Stringy or loose cell pellets may be observed when culturing cells in FABM/dFAS conditions. Take care when aspirating medium from cell pellets.

5. Carefully remove the supernatant from the tube without dislodging the cell pellet.
6. Resuspend the cell pellet in 4 mL complete media. Pipette the cells up and down with a 1 mL pipette tip to ensure a homogeneous cell suspension. Determine the concentration of cells in the suspension using a hemocytometer.
7. Dilute the cells in supplemented media and seed new culture vessels at 2.5 × 10³ cells/cm_².
8. Incubate the cultures in a 37°C, 5% CO₂/95% air, humidified cell culture incubator.

Cryopreservation of Dermal Fibroblasts

1. View the culture under a microscope to ascertain the condition of the culture (i.e., confluence, mitotic activity). We recommend cryopreserving dermal fibroblasts when the culture is approximately 90% confluent and actively growing.

Note:   We do not recommend warming the reagents prior to use.

2. Follow Steps 1–8 in Subculture of Dermal Fibroblasts.
3. Resuspend the cell pellet in a small volume of cold (4°C) Synth-a-Freeze® cryopreservation medium to yield approximately 2–5 × 10⁶ cells/mL.
4. Determine the number of viable cells/mL using a hemocytometer and dilute to the desired final cell density (5–10 × 10⁵ viable cells/mL is recommended).
5. Cryopreserve the cells using a controlled-rate freezer or other appropriate device, then transfer to liquid nitrogen storage (vapor phase).

Twenty-four hours after seeding primary cultures, observe the cultures using phase contrast microscopy. You should be able to see some cells attached to the surface of the flask although there will be a significant number of floating cells and debris. Carefully change the medium so that you do not dislodge lightly adherent cells.

Problem	Cause	Solution
Cells attach poorly	Tissue stored too long/improperly	Use tissue within 24 hours of harvest for best results. Store tissue in culture medium at 4ºC until use.
	Culture surfaces not coated properly	Check the method used for coating flasks or adding coating matrix to the cell inoculum.
	Improper enzymatic treatment	Check the concentration of collagenase. Make sure the collagenase solution is completely removed after centrifugation of the cells.
Cells grow slowly	Medium and/or supplement stored incorrectly, beyond expiration date	Check the expiration date on the label of the products and do not use the product after the expiration date.   Check the storage conditions as described in the product manual. Confirm that the products were stored properly.
	Supplemented medium stored too long or improperly	Store supplemented medium in the dark at 4ºC for up to 1 month from the time the basal medium is supplemented with dFAS
Cells become contaminated with microorganisms	Improper tissue storage	Store tissue in medium containing antibiotic/antimycotic. Store tissue at 4ºC. Wash tissue thoroughly in medium containing antibiotic/antimycotic at the start of the procedure (see Preparing Tissue, Step 7).
	Expired or incorrect concentration of antibiotic/anti-mycotic solution used	Check the expiration date on the product and do not use after the expiration date. Check for the proper dilution of the product in the supplemented medium and correct if necessary.