Name: Isolation of Skeletal Muscle Satellite Cells for In Vitro Myogenesis Studies (Video) | JoVE Journal
Uploaded: 2026-02-24T14:20:00.000+00:00
Duration: 7 min 56 s
Description: Learn an optimized protocol for isolating high-purity satellite cells from mouse skeletal muscle using enzymatic digestion, filtration, and magnetic-activated cell sorting. This method enables reproducible myoblast cultures for muscle regeneration and disease research.

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Confocal imaging was performed at the Laboratory of Imaging Tissue Structure and Function, which serves as an imaging core facility at the Nencki Institute of Experimental Biology and is a part of the infrastructure of the Polish Euro-BioImaging Node. 
This work was supported by National Science Center, Poland (Sonata grant 2022/47/D/NZ3/02737 to Lilya Lehka and Sonata bis grant 2020/38/E/NZ4/00314 to Grzegorz Sumara), and statutory funds from the Ministry of Science and Higher Education to the Nencki Institute. This work was supported by the project financed by the Minister of Education and Science based on contract No 2022/WK/05 (Polish Euro-BioImaging Node &#34;Advanced Light Microscopy Node Poland&#34;).

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</ol>

The authors have no conflicts of interest to declare.

Satellite cells (SCs) constitute the principal myogenic stem and progenitor population responsible for skeletal muscle growth, repair, and regeneration36,37. Their ability to re-enter the cell cycle and differentiate into mature myofibers makes them indispensable for studying skeletal muscle biology. When isolated and cultured under defined in vitro conditions, SCs proliferate as myoblasts that can be induced to differentiate and fuse into multinucleated myotubes, recapitulating key aspects of myogenesis observed in vivo.
Our method enables the recovery of highly viable, homogeneous cell populations by carefully adjusting key parameters, including enzyme concentration, digestion duration, and culture conditions, particularly tailored for neonatal tissues. The combination of enzymatic and mechanical dissociation, followed by multi-step filtration and optional magnetic-activated cell sorting (MACS) together with preplating, ensures high cell yield and purity. Importantly, this protocol enables the generation of cultures that typically exeeds 97% Pax7+ myogenic progenitors, making them suitable for downstream differentiation assays and molecular studies.
The preplating technique relies on the differing adhesion properties of fibroblasts and SCs16,38,39. This step is particularly critical in determining culture purity, and minor deviations in timing or washing can lead to fibroblast contamination or reduced myoblast recovery. The inclusion of a precisely timed preplating step is particularly advantageous, as it efficiently separates myogenic cells from fibroblasts and other non-myogenic populations. This approach can be employed in the absence of magnetic separation equipment or as an additional measure to further increase the purity of myoblast cultures. In our protocol, we optimized this step by using collagen-coated plates and a precisely timed preplating period, which efficiently separates myogenic cells from fibroblasts and other non-myogenic populations.
Another methodological advancement of the current protocol is the use of a defined myoblast proliferation medium (MPM) optimized for both adult and neonatal cells. The combination of high serum concentration, chicken embryo extract (CEE), and basic fibroblast growth factor (bFGF) supports sustained proliferation and maintenance of an undifferentiated phenotype. CEE, in particular, contributes growth-promoting factors that enhance myoblast viability and delay spontaneous fusion, while bFGF maintains SCs identity through activation of canonical signaling pathways29,31. The use of Matrigel-coated surfaces further improves cell adhesion and proliferation efficiency, contributing to the high viability and morphological homogeneity observed within 72 hours post isolation.
Upon transition to differentiation medium, the purified myoblasts readily fuse into multinucleated, contractile myotubes expressing myosin heavy chain (MHC), demonstrating preserved myogenic potential and structural maturation. The resulting cultures are highly suitable for studies of skeletal muscle regeneration, cell signaling, transcriptional regulation, and disease modeling. Confluence is a critical factor, as cultures with very low confluence will fail to progress, while excessively high confluence will induce myoblast fusion and differentiation. Myoblasts should not be maintained in the same culture dish for more than 5 days, regardless of confluence, as prolonged incubation can also promote differentiation.
Despite these advantages, certain steps remain failure-sensitive. Enzymatic digestion must be carefully calibrated to balance tissue dissociation with cell viability. Preplating timing, cell density prior to differentiation, and gentle handling during dissociation all strongly influence yield, purity, and fusion potential. Common issues such as persistent fibroblast contamination, low yield, premature differentiation, poor attachment, or weak myotube formation can often be addressed by minor adjustments in preplating, medium supplementation, or plating technique.
Troubleshooting and practical considerations:
Isolation purity and enzymatic digestion require careful attention, as variations in enzyme type, concentration, incubation time, or tissue mincing can strongly influence both cell yield and culture purity. Under-digestion may leave tissue partially intact, reducing the release of SCs and clogging the strainer in the next step, whereas over-digestion may appear visually similar to a biofilm and can compromise cell viability. Careful optimization is therefore essential for reproducible results.
Filtration and the use of strainers are also critical, since incomplete or inefficient filtration can leave tissue debris or aggregates in the cell suspension, which may interfere with cell attachment and reduce culture purity. Using appropriately sized strainers, gently passing the suspension through them, and pre-wetting the filters can improve debris removal while minimizing mechanical stress on the cells. If larger tissue fragments clog the filter, it should be replaced with a new strainer rather than forcing the suspension through, as cells can become trapped on the filter surface, leading to decreased yield. Repeating filtration with a fresh filter when necessary can further enhance the recovery of viable myoblasts.
Premature or failed differentiation can occur when cultures become overly dense, as this may trigger spontaneous differentiation or premature fusion, thereby compromising experimental timing. Monitoring cell density and adjusting seeding density or medium volume prior to differentiation is therefore critical. In cases of excessive confluence, partial splitting or gentle reseeding can help maintain proliferation without inducing unintended myotube formation. Conversely, if the desired confluency is not reached before the start of differentiation, incubation should be continued until optimal density is achieved to ensure effective differentiation. Maintaining myoblasts beyond the recommended timeframe (&#62;5 days) increases the risk of spontaneous differentiation and reduces proliferative capacity, regardless of initial confluence. Regular assessment of cell morphology, adherence to culture timelines, and timely medium changes are essential to preserve both viability and differentiation potential.
Persistent fibroblast contamination can be minimized through meticulous tissue processing, including careful removal of connective and adipose tissue during muscle dissection. Optimization of the preplating step is particularly important, as rapidly adhering fibroblasts attach to uncoated surfaces while SCs remain in suspension. The use of collagen-coated culture plates in combination with a defined myoblast proliferation medium supplemented with bFGF promotes selective myoblast expansion while limiting fibroblast growth. Additional purification can be achieved through negative selection strategies such as MACS to deplete fibroblast populations. Regular microscopic monitoring is essential to detect early fibroblast overgrowth and enable timely intervention.
In summary, this protocol advances the reproducibility and efficiency of primary myoblast isolation by integrating sequential purification, optimized culture conditions, and adaptable purification options. Importantly, it reliably yields highly enriched Pax7+ cells, which can be further used for controlled studies of SCs biology, myogenic commitment, and differentiation dynamics.
In contrast to protocols optimized for prolonged serial expansion, which can alter cell behavior and reduce myogenic potential and fusion ability, our method is intentionally designed to preserve cells with a robust myogenic identity and high fusion competence. This focus is advantageous for applications that require physiological relevance, efficient myogenic differentiation, and retention of defining SCs characteristics, particularly in experiments centered on studying myogenic differentiation and regeneration.

The capacity of skeletal muscle to regenerate following any insult to its integrity is fundamentally dependent on a specialized population of myogenic precursor cells, known as satellite cells (SCs), named for their satellite position along the myofiber. They localize beneath the muscle fiber lamina, attached to the multinucleated muscle fibers1,2,3. SCs are characterized by the expression of the paired box protein Pax7, which is conserved across multiple species, including human, mouse, monkey, and pig. Pax7, which was identified as the first detectable marker for SCs in both quiescent and activated states, is essential for the development and survival of SCs4. Under normal conditions, SCs are retained in a quiescent state (G0 phase of the cell cycle), characterized by low metabolic activity and being transcriptionally inactive5,6. Upon muscle injury or mechanical stress, they become activated, resume the cell cycle, expand in number, and differentiate to contribute to new myofibers or repair damaged ones7,8,9. During this process, a subset of SCs must undergo self-renewal, returning to a quiescent (Pax7&#8314;) state to maintain the long-term integrity of the stem cell pool. Self-renewal occurs through either asymmetric division, where one daughter cell retains stem-like characteristics while the other becomes a committed myogenic progenitor, or symmetric division, where both daughter cells either maintain a stem-like identity or differentiate, depending on the physiological and niche context10,11,12.
SCs play a vital role in muscle regeneration, making them a key focus in skeletal muscle biology, regenerative medicine, and muscular dystrophy research. Understanding how these cells behave and function not only sheds light on the fundamental processes of muscle repair but also paves the way for developing therapies to treat muscle-wasting conditions and enhance regeneration. 
Isolation of SCs from muscle tissue is a crucial step in in vitro studies, enabling investigation of their molecular characteristics, proliferation potential, differentiation pathways, and responses to various stimuli. However, this process possesses several technical challenges due to the muscle&#39;s complex structure, the relatively low abundance of satellite cells compared to other cell types, and the need to preserve their viability and stemness during isolation.
Currently, three primary techniques are employed for the isolation of SCs: the preplating method, fluorescence-activated cell sorting (FACS), and magnetic-activated cell sorting (MACS). Each method offers distinct advantages and limitations in terms of purity, yield, cell viability, and SCs preservation.
The preplating method exploits the differential adhesive properties of muscle-derived cells, with SCs being among the least adherent. After digesting skeletal muscle enzymatically, the resulting heterogeneous cell suspension is plated onto culture plates typically coated with collagen type I. After 1-24 h (depending on the protocol) incubation at 37&#176;C, non-adherent cells are collected and transferred to a Matrigel-coated plate for further culture13,14,15,16. The resulting population usually includes both SCs and fibroblasts as well as a small number of adipocytes and endothelial cells17,18. Repeating the preplating process every 24-48&#160;h for up to a week for SCs isolated from adult mice and up to 2 weeks for neonates can improve the purity of the cell population. While this method is cost-effective and technically straightforward, it is labor-intensive and often yields cultures with variable composition. The preplating method is suitable for laboratories without FACS or MACS, offering a simple way to enrich satellite cells while preserving viability and stem-like properties. Therefore, it is generally recommended to combine this approach with additional purification methods to enhance the overall purity of the cell population. In the current protocol, we optimized the preplating step by using collagen-coated plates and adjusting the timing depending on the model. This differential adhesion accelerates fibroblast removal, shortening the preplating period, using growth factors, and reducing both handling time and cellular stress during isolation.
FACS is a widely used method for isolating satellite cells based on the expression of specific surface markers (&#945;7-integrin, CD34, CD29, &#946;1-integrin, CXCR4), enabling high purity and specificity19,20. However, it is often challenging due to technical complexity, high cost of antibodies, and the need for specialized equipment. Moreover, the mechanical and biochemical stress imposed during sorting can compromise cell viability. The requirement for generating a single-cell suspension may result in significant cell loss, while reliance on specific fluorescent markers can lead to an incomplete representation of satellite cell heterogeneity. Additionally, enzymatic digestion, which is employed to dissociate cells, has the potential to degrade critical surface antigens, thereby affecting accurate identification and isolation as well, and may also impair myoblast function21,22. FACS is&#160;suitable for isolation highly pure populations within a short period, particularly when precise phenotypic resolution is required. However, this method is less appropriate when maintaining intact, viable cells is critical, due to the mechanical and biochemical stresses imposed during sorting, or for labs lacking specialized equipment.
MACS, on the other hand, is a technique used to isolate satellite cells by binding magnetic beads conjugated to antibodies against specific surface markers of fibroblasts and general immune cells to their surfaces and then separating them from unwanted cell populations using a magnetic field. The elution of the negative selection from the stationary phase is the satellite cell population of interest. MACS enables efficient enrichment of satellite cells with relatively high purity and scalability, while requiring less specialized equipment and technical expertise compared to FACS. Furthermore, its gentle magnetic separation process preserves cell integrity, making it suitable for downstream applications such as cell culture, gene expression analysis, and transplantation studies. However, MACS is limited by lower purity when multiple markers are needed, possible non-specific binding or bead carryover, and a lack of quantitative data on marker expression23,24,25.
Despite the availability of protocols for skeletal muscle cell isolation, obtaining reproducible and compositionally uniform primary cultures remains challenging, particularly due to developmental and tissue-specific variability. SCs isolation from neonatal muscles has been less extensively characterized than from adult muscles, with relatively fewer published protocols, many of which require further optimization depending on technical conditions and experimental purpose26,27,28. Adjustment of parameters, including enzyme concentration, digestion duration, and culture conditions, is often needed for different developmental stages-especially in neonates-to achieve maximal yield, viability, and purity of isolated satellite cells. In this article, we demonstrate an optimized protocol for the isolation of satellite cells from skeletal muscles of adult and neonatal mice (postnatal days 6-14, P6-P14), combining preplating and MACS sorting with adjustments to preplating timing and medium composition. This protocol offers&#160;a reproducible cell population across developmental stages, with isolated cells showing high purity and viability. Immunostaining indicates that the proportion of Pax7+ cells typically exceeds 97%. Moreover, these myoblasts exhibit robust differentiation potential, efficiently forming multinucleated myotubes upon replacement of the growth medium with differentiation medium. Differentiated myotubes can be observed using phase-contrast microscopy or verified through immunostaining for myosin heavy chain (MHC), a marker indicative of mature muscle fibers.

<table><tbody><tr><td>0.25% trypsin-EDTA&amp;nbsp;</td><td>Gibco</td><td>25200-056</td><td></td></tr><tr><td>100 &amp;micro;m cell strainer&amp;nbsp;</td><td>VWR</td><td>732-2759</td><td></td></tr><tr><td>30 &amp;micro;m strainer&amp;nbsp;</td><td>Miltenyi Biotec</td><td>130-041-407</td><td></td></tr><tr><td>40 &amp;micro;m cell strainer&amp;nbsp;</td><td>VWR</td><td>732-2757</td><td></td></tr><tr><td>70 &amp;micro;m cell strainer&amp;nbsp;</td><td>VWR</td><td>130-095-823</td><td></td></tr><tr><td>Acetic acid&amp;nbsp;</td><td>POCH</td><td>568760114</td><td>Dissolved in dH&lt;sub&gt;2&lt;/sub&gt;O to a final concentration of 20 mM and sterilized through a 0.2 &amp;micro;m filter</td></tr><tr><td>Alexa Fluor 488, donkey anti-Rabbit</td><td>Invitrogen&amp;nbsp;</td><td>A21206</td><td>Dissolved in blocking solution (see Section 8.7) at a 1:250 dilution</td></tr><tr><td>Alexa Fluor 488, goat anti-mouse&amp;nbsp;</td><td>Abcam</td><td>ab150113</td><td>Dissolved in blocking solution (see Section 8.7) at a 1:250 dilution</td></tr><tr><td>Ammonium chloride (NH&lt;sub&gt;4&lt;/sub&gt;Cl)&amp;nbsp;</td><td>BioShop</td><td>AMC303.500</td><td>Dissolve in water to a final concentration 50 mM</td></tr><tr><td>Antifade Mounting Media</td><td>Vectashield</td><td>H-2000-10</td><td></td></tr><tr><td>Anti-PAX7 antibody</td><td>Abcam</td><td>ab187339</td><td>Dissolved in blocking solution (see Section 8.7) at a 1:100 dilution</td></tr><tr><td>Basic fibroblast growth factor&amp;nbsp;</td><td>Corning</td><td>354060</td><td>Dissolved in sterile dH&lt;sub&gt;2&lt;/sub&gt;O to a stock concentration of 10 &amp;micro;g/mL&amp;nbsp;</td></tr><tr><td>Betadine, 100 mg/mL</td><td>EGIS Pharmaceuticals</td><td></td><td></td></tr><tr><td>Cell culture plate, 6 well, Cell+</td><td>SARSTEDT</td><td>83.3902.300</td><td></td></tr><tr><td>Chicken embryo extract&amp;nbsp;</td><td>MP Biomedicals</td><td>2850145</td><td></td></tr><tr><td>Collagen I, rat tail</td><td>Gibco</td><td>A10483-01</td><td>Dissolved in 20 mM acetic acid&amp;nbsp;</td></tr><tr><td>Collagenase, type I</td><td>Sigma</td><td>C0130-500MG</td><td>Dissolved in DMEM and sterilized through a 0.2 &amp;micro;m filter</td></tr><tr><td>Dissociation tube (gentleMACS C Tubes)</td><td>Miltenyi Biotec</td><td>130-093-237</td><td></td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium, high glucose</td><td>Gibco</td><td>31966-021</td><td></td></tr><tr><td>EDTA</td><td>Invitrogen</td><td>AM9260G</td><td></td></tr><tr><td>Falcon Conical Centrifuge Tubes, 15 mL</td><td>Corning</td><td>352096</td><td></td></tr><tr><td>Falcon Conical Centrifuge Tubes, 50 mL</td><td>Corning</td><td>352070</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Gibco</td><td>10082147</td><td></td></tr><tr><td>Horse serum&amp;nbsp;</td><td>Gibco</td><td>26050088</td><td></td></tr><tr><td>Magnetic labeling reagent (Satellite Cell isolation Kit)</td><td>Miltenyi Biotec</td><td>130-104-268</td><td></td></tr><tr><td>Magnetic separator (MidiMACS Separator)</td><td>Miltenyi Biotec</td><td>130-042-302</td><td></td></tr><tr><td>Matrigel&amp;nbsp;</td><td>Corning</td><td>CLS356234</td><td>Dissolved in DMEM to 10 % concentration&amp;nbsp;</td></tr><tr><td>Paraformaldehyde, 4% in PBS</td><td>Thermo Fisher Scientific</td><td>J61899.AK</td><td></td></tr><tr><td>Penicillin-Streptomycin&amp;nbsp;</td><td>Gibco</td><td>15140122</td><td></td></tr><tr><td>Phosphate-Buffered Saline&amp;nbsp;</td><td>VWR</td><td>392-0334</td><td></td></tr><tr><td>Separation column (LS Columns)</td><td>Miltenyi Biotec</td><td>130-042-401</td><td></td></tr><tr><td>Skeletal Muscle Myosin Antibody, MHC</td><td>Santa Cruz</td><td>sc-32732</td><td>Dissolved in blocking solution (see Section 8.7) at a 1:50 dilution</td></tr><tr><td>TC-dish, 100 x 20 mm</td><td>SARSTEDT</td><td>83.3902</td><td></td></tr><tr><td>Tissue dissociator (GentleMACS Dissociator)</td><td>Miltenyi Biotec</td><td>130-093-235</td><td></td></tr><tr><td>Triton X-100&amp;nbsp;</td><td>Sigma-Aldrich</td><td>X100-1L</td><td></td></tr></tbody></table>

isolation of skeletal muscle satellite cells for in vitro myogenesis studies

Isolation of Skeletal Muscle Satellite Cells for In Vitro Myogenesis Studies

Skeletal muscle regeneration and growth depend on satellite cells, the muscle tissue&#39;s resident stem cells. Currently, these cells serve as the&#160;in vitro model that most accurately represents natural myogenic processes. Nonetheless, the isolation of satellite cells is technically challenging as they exist in only small numbers within skeletal muscle and the typically heterogeneous nature of the isolated cell populations. In this study, we describe a standardized method for isolating and enriching primary myoblasts from skeletal muscles of both adult and neonatal mice. The procedure outlines muscle dissection, enzymatic dissociation, sequential filtration, preplating steps to limit non-myogenic cells, and defined culture conditions for establishing satellite cell-derived myoblast cultures. The protocol provides practical guidance on timing, handling, and critical steps for establishing a successful culture. Primary myoblasts generated using this protocol can be used for a variety of downstream applications, including studies of myogenesis, functional analysis, genetic manipulation, live-cell imaging, or drug testing. The cells reliably fuse and mature, making them well-suited for studying myoblast fusion, muscle development, regenerative mechanisms to&#160;restore skeletal muscle function, as well as their metabolism.

Successful execution of this protocol yields highly viable, morphologically distinct primary myoblast cultures derived from hindlimb muscles of both adult and neonatal mice. By combining sequential filtration through cell strainers of decreasing pore sizes with optional magnetic-activated cell sorting (MACS), this protocol enables the isolation of a highly purified myoblast population. To further improve the purity of the isolated Pax7+ cells, an optimized preplating step was incorporated. After this step (protocol section 5), most fibroblasts and other non-myogenic cells remain adherent (Figure 3) to the initial collagen-coated surface, whereas myoblasts stay in suspension and subsequently attach to the Matrigel-coated plates (Figure 2B). Building on the previously demonstrated effectiveness of specific components of the myoblast proliferation medium (MPM), including high serum concentration, supplementation with bFGF and chicken embryo extract (CEE), and the use of Matrigel coating-this formulation, optimized by us, provides a distinct proliferative advantage for myoblasts over non-myogenic cells14,30,31,35. Thus, within 72 h of culture in MPM, small, round to spindle-shaped myoblasts with a high nuclear-to-cytoplasmic ratio should predominate (Figure 2C). A minor population of large, flattened fibroblast-like cells may remain, typically representing less than 3% of the culture.
Cell yield after 72 h post isolation using this protocol depends on the age of the donor animals. For myoblasts isolated from one adult mouse using four hindlimb muscles (gastrocnemius, soleus, tibialis anterior, and extensor digitorum longus) from both legs, the typical yield is approximately 2-4 × 10⁶ cells per animal. More than 97% of the cells isolated from adult (**p &lt; 0.01) and neonatal mice (***p &lt; 0.0001) were myogenic progenitors, as indicated by Pax7+ expression (Figure 4A,B), confirming the high enrichment of myogenic progenitors. This Pax7-based purity assessment was performed after completion of the full enrichment workflow, including preplating followed by MACS and/or expansion and not on the early adherent preplate fractions.
Neonatal myoblasts achieved high purity with or without MACS (Figure 4A,B), making this protocol suitable for laboratories with varying levels of technical support. However, when MACS is omitted, additional preplating steps are required to reach comparable purity.
For adult myoblasts, direct statistical comparison of MACS-sorted versus unsorted populations was not performed. However, given lower proliferative capacity and greater commitment to differentiation, MACS is expected to enhance enrichment relative to preplating alone.
Upon switching to differentiation medium (MDM) after reaching 80-90% confluency, myoblasts efficiently fuse to form elongated, multinucleated myotubes within 48-72 h. These myotubes are readily visualized by phase-contrast microscopy (Figure 2D) and can be further confirmed by immunostaining for myosin heavy chain (MHC) (Figure 5A,B). The presence of organized sarcomeric structures in differentiated myotubes (Figure 5B) indicates structural maturation consistent with myogenic differentiation under these culture conditions.
<img alt="Muscle satellite cell isolation process diagram; stages: tissue digestion, cell filtration, MACS." class="xfigimg" src="/files/ftp_upload/70277/70277fig1.jpg" data-altupdatedat="1778558422000" data-alt="Figure 1"> 
Figure 1: Step-by-step protocol for the efficient isolation of muscle satellite cells. <a href="https://www.jove.com/files/ftp_upload/70277/70277fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Muscle cell differentiation stages; neonatal/adult cells; myoblast to myotube; microscopy images." class="xfigimg" src="/files/ftp_upload/70277/70277fig2.jpg" data-altupdatedat="1778558422000" data-alt="Figure 2"> 
Figure 2: Representative phase-contrast microscopy images of the different methodologies discussed in the text. (A) Phase-contrast (20x) images of the isolation of the cells after digestion. (B) Phase-contrast (20x) of the preplating of the cultures with attached fibroblast, together with unattached myoblasts. (C) Phase-contrast (20x) of proliferating myoblasts once attached. (D) Phase-contrast (20x) of differentiated myotubes. Scale bar = 50 µm. <a href="https://www.jove.com/files/ftp_upload/70277/70277fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Neural cell cultures, neonatal and adult, with/without MACS, microscope image, cell density comparison." class="xfigimg" src="/files/ftp_upload/70277/70277fig3.jpg" data-altupdatedat="1778558422000" data-alt="Figure 3"> 
Figure 3: Representative phase-contrast microscopy images (20×) of cells remaining attached following the first preplating step. Scale bar = 50 µm. <a href="https://www.jove.com/files/ftp_upload/70277/70277fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Neonatal and adult cell microscopy with DAPI/Pax7 staining, includes bar graph of nuclear percentages." class="xfigimg" src="/files/ftp_upload/70277/70277fig4.jpg" data-altupdatedat="1778558422000" data-alt="Figure 4"> 
Figure 4: Representative images taken under phase-contrast microscope (20x) and immunocytochemistry (40x). (A) Pax7+ (green) and nuclei stained with DAPI (light blue), and their colocalization in the merged channels. (B) Analysis of the purity of myoblast culture, quantifying the percentage of Pax7+ cells normalized to nuclei per field acquired (one field per well, in at least six wells). Immunostaining was repeated at least three times, with two to three technical replicates per experiment. **p &lt; 0.01, ***p &lt; 0.0001 (Statistical significance was analyzed using Mann Whitney U test). Scale bar = 50 µm. <a href="https://www.jove.com/files/ftp_upload/70277/70277fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Fluorescence microscopy, MHC/DAPI-labeled nascent, mature myotubes; cellular differentiation study." class="xfigimg" src="/files/ftp_upload/70277/70277fig5.jpg" data-altupdatedat="1778558422000" data-alt="Figure 5"> 
Figure 5: Representative immunofluorescence images showing myotubes differentiated from adult mouse myoblasts. (A) Nascent myotubes (3 days in culture) imaged at 20x of MHC (green) and nuclei stained with DAPI (light blue). (B) Mature myotubes (6 days in culture) imaged at 20x and details of the structure of the sarcomeric pattern defined by the myosin staining. Immunostaining was repeated at least three times, with two to three technical replicates per experiment. Scale bar = 50 µm; zoomed-in scale bar = 10 µm. <a href="https://www.jove.com/files/ftp_upload/70277/70277fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

This protocol provides a standardized method for isolating and enriching primary myoblasts from adult and neonatal mouse skeletal muscle through tissue dissociation, sequential filtration, preplating, and defined culture conditions, enabling the establishment and maintenance of satellite cell-derived myoblast cultures for downstream experimental applications.

Skeletal muscle regeneration and growth depend on satellite cells, the muscle tissue&#39;s resident stem cells. Currently, these cells serve as ...

We developed an optimized protocol to isolate viable satellite cells from neonatal and adult mouse skeletal muscle. Existing methods yield variable purity, low neonatal efficiency, and stress-induced cell loss. Our method preserves viability, high purity, as well as reproducibility.To begin, obtain skeletal muscle tissue from mice and wash it first with PBS containing 1%penicillin and streptomycin and a few drops of Betadine. Then rapidly wash the muscles twice in fresh PBS containing penicillin and streptomycin to thoroughly rinse off the Betadine. Next, using autoclave scissors, mince the washed tissue finely into fragments measuring approximately one to two millimeters.Transfer the minced muscles into tissue dissociation tubes containing 0.2%collagenase in DMEM. Incubate the tubes at 37 degrees Celsius for 30 minutes to initiate enzymatic digestion. Using a mechanical tissue dissociator, run the predefined program to facilitate enzymatic digestion while preserving cell viability.Apply the program twice to enhance mechanical disruption of the tissue. Then place the samples back into the incubator at 37 degrees Celsius for an additional 30 minutes to complete the digestion process. Repeat mechanical dissociation once more to ensure thorough dissociation.Confirm that the digestion medium becomes cloudy with few or no visible tissue fragments remaining. Next, place a 100-micrometer cell strainer on top of a 50-milliliter conical tube. Using a pipette, pre-wet the strainer with one milliliter of pre-warmed neutralizing medium.Pour the digested muscle suspension over the pre-wetted 100-micrometer strainer. Then rinse the strainer with an additional one milliliter of neutralizing medium to remove residual debris. After straining the suspension over 70 and 40-micrometer cell strainers, pre-wet a 30-micrometer cell strainer placed on a 15-milliliter conical tube with one milliliter of neutralizing medium.Then pass the filtrate through the strainer and wash again with one milliliter of neutralizing medium to maximize cell recovery. Centrifuge the collected cell suspension at 300 g for 15 minutes at room temperature. Under a sterile biosafety cabinet, carefully remove and discard the supernatant.Confirm the presence of a white cell pellet at the bottom of the tube. To perform magnetic-activated cell sorting, first, suspend the cell pellet in freshly prepared MACS buffer using 80 microliters of buffer per gram of original tissue. Add 20 microliters of magnetic-labeling reagent.Then gently mix and incubate for 15 minutes at two to eight degrees Celsius on ice. Adjust the total volume to 500 microliters with sorting buffer for samples up to five grams of tissue. For larger tissue amounts, divide the suspension equally into multiple tubes.Place a separation column in the magnetic field of a magnetic separator. Wash the column with three milliliters of sorting buffer to equilibrate it. Apply the labeled cell suspension onto the column.Collect the eluant containing the satellite cell-enriched population. Then wash the column twice with one milliliter portions of sorting buffer and combine the collected fractions. Centrifuge the combined suspension at 300 for 10 minutes at room temperature to pellet the isolated satellite cells.Dilute Type I rat collagen in 20 millimolar acetic acid to a final concentration of 50 microgram per milliliter. Add the solution to the culture plates and incubate at room temperature for one hour. Carefully aspirate the collagen solution from the plates.Then rinse the plates three times with sterile PBS using equal volumes to remove residual acid. Next, prepare myoblast proliferating medium for adult mouse myoblasts or neonatal mouse myoblasts. Resuspend the cell pellet in 10 milliliters of myoblast proliferating medium, pre-warmed to 37 degrees Celsius.Seed adult myoblasts onto the collagen-coated plates. Then incubate the plates at 37 degrees Celsius in a carbon dioxide incubator for 24 hours. After the optimized pre-plating step, most fibroblasts and other non-myogenic cells remained adherent to the initial collagen-coated surface, whereas myoblasts stayed in suspension and subsequently attached to the Matrigel-coated plates.Within 72 hours of culture in myoblast proliferation medium, small, round spindle-shaped myoblasts with a high nuclear cytoplasmic ratio predominated. A minor population of large, flattened fibroblast-like cells remained in culture, typically representing less than 3%of the total cells. More than 97%of the cells isolated from adult and neonatal mice were myogenic progenitors, as indicated by PAX7 positive expression.Neonatal myoblasts achieved high purity with or without magnetic-activated cell sorting. After switching to differentiation medium at 80 to 90%confluency, myoblasts fused to form elongated multinucleated myotubes within 48 to 72 hours. Differentiated myotubes were confirmed by immunostaining for myosin heavy chain.The presence of organized sarcomeric structures in differentiated myotubes indicated structural maturation under these culture conditions. This method enables direct observation of satellite cell behavior, fusion parameters, marker expression, and responses to environmental or chemical stimuli. The main challenge is controlling digestion, plating density, and fiberblast removal to ensure healthy, high purity myogenic cells.Isolated cells can support research on muscle regeneration, disease modeling, drug testing, and mechanisms controlling satellite cell differentiation and self-renewal.

isolation-skeletal-muscle-satellite-cells-for-vitro-myogenesis

Research

JoVE Journal

Biology

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Preparation of Primary Myogenic Precursor Cell/Myoblast Cultures from Basal Vertebrate Lineages

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Isolation and Characterization of Satellite Cells from Rat Head Branchiomeric Muscles

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Isolation of Skeletal Muscle Satellite Cells for In Vitro Myogenesis Studies

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