Single Fiber Culture

This method is adapted from the technique used by Prof Michael Rudnicki for the isolation and culture of mouse single muscle fibers. It is useful so as to follow the cell divisions of individual satellite cells, while they remain in the satellite cell niche (between the sarcolemma and the basal lamina). It is also possible to follow a number of important events in the cycle of satellite cells, including activation, proliferation and differentiation. It should be noted that this technique can be adapted to generate pure satellite cell cultures, and can also be used to follow satellite cell migration from the fiber into a matrix. Some very good examples of the power of this technique can be seen in the following publications: Juan et al. 2011, Kuang et al. 2007, Beauchamp et al. 2000 and many more.


  • Filter heat inactivated horse serum (~2ml/animal) through a 0.44um filter and then coat (per animal) 5x60mm culture plates for fiber isolation (fill one plate with 2-3ml, swirl and then transfer to next plate). Allow the plates to dry for 5 mins. The horse serum, will prevent fibers from sticking to the plates.
  • Add 5ml DMEM (25mM glucose + sodium-pyruvate) to each plate, and the place plates in an incubator at 37°C (and 5% CO2)
  • Flame-polish glass Pasteur pipettes. Coat pipettes with heat inactivated horse serum and store at 37°C
  • Prepare growth media (For 50ml: 45ml DMEM (25mM glucose + sodium-pyruvate), 500ul Pen/Strep (100x concentrate), 5ml heat inactivated horse serum, 250ul chick embryo extract) and place in a waterbath at 37°C
  • Prepare collagenase solution (2ml per animal, in a 15ml falcon tube, [make 10ml, good for one week]: 10ml DMEM (25mM glucose + sodium-pyruvate), 100ul Pen/Strep (100x concentrate), 20mg type I collagenase) and incubate in a water bath at 35°C

Fiber isolation

  • Euthanize mouse (as per approved guidelines), spray with EtOH and remove skin from the midline to the ankles (being careful not to transfer any hair onto the muscle)
  • Carefully dissect muscles of interest (EDL and/or soleus) from both hindlimbs. Do not stretch the muscle, and do not touch the muscle directly, handle the muscle via the tendons.
  • Place the muscles immediately into the warmed collagenase solution and incubate at 35°C for 45min. Inverting the tube every 5-10mins (time may vary depending on collagenase, the muscle is ready when fibers can be seen beginning to detach)
  • To stop digestion, transfer the muscles (in solution, use a large bore pipette) to one of the five plates (prepared earlier, one muscle per plate, leaving three empty) containing DMEM
  • One muscle at a time, triturate with the glass pipettes until 20-30 fibers have been released, then transfer the muscle (in solution, use a large bore pipette) to a new plate. Return the freshly isolated fibers to the 37°C incubator

Fiber culture

  • Transfer ‘good’ fibers (should be long and shiny) with a horse serum coated glass Pasteur pipette to Permanox slides (or 24/48 well plates) one at a time (1-3 fibers per well)
  • Wait 2-3mins and then add 350ul growth media to each well
  • Place slide/s at 37°C (and 5% CO2), media should be changed every 48-72hours


  • Wash in 1% Glycine in PBS (2×1 mins)
  • Fix in 2%PFA in PBS (1×10 mins)
  • Wash in PBS (2×2 mins)
  • Wash in 1% Glycine in PBS (1×15 min)
  • Block for 1 hour in blocking buffer (5% heat inactivated horse serum, 1% BSA, 0.5% triton-X 100 in PBS)
  • Incubate overnight at 4°C in 1°ab (diluted in blocking buffer)
  • Wash in PBS (3×5 min)
  • Incubate for 1 hour at RT in 2°ab (diluted in blocking buffer)
  • Wash in PBS (3×5 min)
  • Mount appropriately

Figure – Single Fiber Cultures. Satellite cells from freshly isolated single fibers (T=0) are Pax7 positive but do not yet express MyoD. Following 24hrs in growth media, satellite cells begin to express MyoD.


6 thoughts on “Single Fiber Culture

      1. Y


        thanks for your kind reply. i have one more question: it looks clean under microscope after i wash myofiber three times, but many black debris appears in culture media around suspensioned myofiber in next day. i feel confused where the debris from(it seems from myofiber itself), have you ever see this phenomenon?


      2. Without being able to see the image, I can only guess that it might be something in your growth media (do you filter the media prior to use?) or something is getting transferred with your fiber. You could confirm that it is debris and not contaminating cells by performing a simple dapi stain.

Leave a Reply

Fill in your details below or click an icon to log in: Logo

You are commenting using your account. Log Out /  Change )

Google+ photo

You are commenting using your Google+ account. Log Out /  Change )

Twitter picture

You are commenting using your Twitter account. Log Out /  Change )

Facebook photo

You are commenting using your Facebook account. Log Out /  Change )


Connecting to %s