This study was funded by a Mosaic grant (017.009.009) from The Netherlands Organization for Scientific Research (NWO) and a Start grant (S-13-167C) for young investigators from the AO Foundation. Z.Y.R is supported by the National Institutes of Health (grant # AG021566, AG035377, NS090051).

All experiments described herein were approved by the local Board for Animal Experiments from the Radboud University Nijmegen in accordance with Dutch laws and regulations (RU-DEC 2013-205).
1. Extracellular Matrix Gel Spots
<ol>
	<li>Perform the following steps one day before the isolation:
	<ol>
		<li>Thaw an aliquot extracellular matrix gel (100 &#181;l) at 4 &#176;C for at least 1.5 hr. Dilute 1:10 in Dulbecco's modified Eagle's medium; with 4,500 mg/L glucose, 4 mM L-glutamine, and 110 mg/ml sodium pyruvate (DMEM). Keep the extracellular matrix gel at 4 &#176;C at all times. Note: Abrupt temperature changes will result in uneven coating and crystal formation.</li>
		<li>Keep the diluted extracellular matrix gel solution on ice for 15 min.</li>
		<li>Pre-chill a 20 &#181;l micropipette for 10 min.</li>
		<li>Put 8-well chamber slides into a 100 mm Petri dish and transfer the dish onto a cold surface (e.g. a freezer pack) for 10 min.</li>
		<li>Use the pre-chilled micropipette to put a drop of 10 &#181;l extracellular matrix gel in each well. Keep the Petri dish on the cold surface for at least another 7 min (Figure 1A).</li>
		<li>Completely remove the remaining extracellular matrix gel (Figure 1B), and dry the wells at 37 &#176;C overnight.</li>
	</ol>
	</li>
</ol>
2. Dissection of Head Muscles (Masseter, Digastric, and Levator Veli Palatini)
<ol>
	<li>Before dissection, prepare 50 ml of phosphate-buffered saline (PBS) supplemented with 2% Penicillin-Streptomycin (P/S). Keep on ice.</li>
	<li>After euthanasia of one young adult rat (9 weeks) with CO2/O2, decapitate the head and remove the skin from the head. Transfer the head to ice-cold PBS supplemented with 2% P/S in a 50 ml tube.</li>
	<li>Masseter muscle (derived from the 1st branchial arch)
	<ol>
		<li>Place the head with one side up on a silicone pad and fix with hypodermic needles (Figure 2A).</li>
		<li>Identify the parotid gland and the facial nerve (Figure 2A). Expose the deep fascia covering the gland. Cut the fascia and remove the gland using dissection scissors. Identify the external auditory canal. Trace the facial nerve from the stylomastoid foramen and carefully remove the temporal, zygomatic, and buccal branches with a scalpel blade No. 15.</li>
		<li>Free the superficial head of the masseter muscle by removing the fascia. Identify both superficial and deep heads of the masseter muscle. Trace the superficial head until its thick tendinous aponeurosis inserted in the zygomatic process of the maxilla.</li>
		<li>Separate the tendon from its origin at the zygomatic process with a straight forceps. Cut it with a scalpel blade No. 15 or dissection scissors and carefully life it (Figure 2B).</li>
		<li>Dissect the superficial head of the masseter until its insertion at the angle and inferior half of the lateral surface of the ramus of the mandible with a scalpel blade No. 15 (Figure 2C). Now, completely remove the muscle.</li>
	</ol>
	</li>
	<li>Posterior belly of the digastric muscle (derived from the 2nd branchial arch)
	<ol>
		<li>Place the head in a supine position on the silicone pad and fix with hypodermic needles (Figure 3A).</li>
		<li>Remove the subcutaneous fat overlying both sublingual and submandibular glands. Next, remove the superficial fascia and glands using dissection scissors. Expose the digastric muscle (anterior and posterior belly).</li>
		<li>Hold the anterior tendon of the posterior belly with a straight forceps, cut it, and dissect it carefully until its origin in the tympanic bulla (Figure 3B). Do the same at the contralateral side.</li>
	</ol>
	</li>
	<li>Levator veli palatini muscle (derived from the 4th branchial arch)
	<ol>
		<li>After dissection of the posterior belly of the digastric muscle, localize the stylohyoid muscle, pull it laterally, and carefully remove it (Figure 4A).</li>
		<li>Localize the tendon of the levator veli palatini that inserts at the tympanic bulla (Figure 4A). Dissect it carefully and cut it on both sides.</li>
		<li>Look for the trachea and the esophagus that runs behind it. Lift the esophagus, and expose the pharynx and the larynx.</li>
		<li>Localize and dissect the area of the superior pharyngeal constrictor muscle. Identify the levator veli palatini and cut it at both sides (Figure 4B). 
Note: Directly after dissection, carefully remove tendon and connective tissue from each muscle under the stereo microscope. Submerge all specimens quickly in ethanol 70%, and transfer them to ice-cold PBS supplemented 2% P/S in a 15 ml tube.
</li></ol>
</li>
</ol>
3. Isolation of Satellite Cells
<ol>
	<li>Perform the following preparation steps for SC isolation from 3 groups of muscles:
	<ol>
		<li>Prepare 7.5 ml of 0.1% pronase in DMEM. Filter the solution through a 0.22 &#181;m filter. Pre-warm the solution at 37 &#176;C in a water bath for 10 min before isolation.</li>
		<li>Prepare 35 ml of DMEM supplemented with 10% Horse Serum (HS) and 1% P/S. Also pre-warm at 37 &#176;C in a water bath.</li>
		<li>Prepare 15 ml culture medium which consists of DMEM supplemented with 20% fetal bovine serum (FBS), 10% HS, 1% P/S and 1% chicken embryo extract (CEE). Pre-warm at 37 &#176;C in a water bath.</li>
		<li>Pre-coat six plastic pipettes (10 ml) with HS and dry for at least 10 min before use.</li>
	</ol>
	</li>
	<li>In the culture hood, transfer each muscle into a well of a 6-well plate. Using the dissection scissors, cut the muscle in small pieces of about 2 mm. Be careful not to mince the tissue too much.</li>
	<li>Carefully add 2.5 ml of 0.1% pronase solution to each well and incubate at 37 &#176;C for 60 min. Gently shake the plate after 20, 40, and 60 min. Note: The exact duration of the incubation depends on factors like age and strain of the animals.</li>
	<li>Monitor under the microscope. Check the muscle fragments and stop the enzymatic digestion when the fibers bundles get a loosened appearance (Figure 5).</li>
	<li>Add 2.5 ml of DMEM supplemented with 10% HS and 1% P/S. Transfer to a 15 ml tube and centrifuge the tubes at 400 x g for 5 min. Discard the supernatant by decantation.</li>
	<li>Add 5 ml DMEM supplemented with 10% HS and 1% P/S. Pipette the solution up and down with a 10 ml plastic pipette (trituration) for at least 20 times to homogenize the tissue.</li>
	<li>Centrifuge the tubes at 200 x g for 4 min. Collect the supernatant and transfer into a 15 ml tube.</li>
	<li>Add 5 ml DMEM supplemented with 10% HS and 1% P/S. Pipette again with a 10 ml plastic pipette until the tissue fragments passes easily through the pipette.</li>
	<li>Centrifuge the tubes at 200 x g for 4 min and collect the supernatant in a 15 ml tube.</li>
	<li>Put a cell strainer (40 &#181;m) onto a 50 ml tube and transfer the supernatant containing the dissociated cells onto the filter. Wash with 1 ml DMEM for maximal cell recovery.</li>
	<li>Centrifuge the tubes at 1,000 x g for 10 min and discard the supernatant with a pipet.</li>
	<li>Resuspend the pellet in 300 &#181;l culture medium and count the cells in a hemocytometer.</li>
</ol>
4. Differentiation of Satellite Cells on Extracellular Matrix Gel Spots
<ol>
	<li>Dilute the cell suspension to obtain 1.5 x 103 cells in 10 &#181;l of culture medium.</li>
	<li>Secure the covers of the chambers slides with tape and mark the spots with a black marker on the bottom side of the object glass.</li>
	<li>Using a micropipette, put a drop of 10 &#181;l cell suspension onto the extracellular matrix gel spot. Check under the microscope whether the drop of cell suspension has been placed correctly on the spot. Incubate for six hours at 37 &#176;C.</li>
	<li>Carefully add 400 &#181;l of culture medium (DMEM supplemented with 20% FBS, 10% HS, 1% P/S and 1% CEE) and incubate for three days at 37 &#176;C. 
	Note: At this point, freshly isolated SC are subjected to massive trauma (enzymatic digest and harsh trituration) and they need to recover. Do not disturb the cells during the first three days 37. Next, the culture medium can be changed depending on the type of experiment. 
	The extracellular matrix gel spots can be seeded with a high cell density (1.5-2.5 x 103/20 &#181;l) for differentiation assay. Culture medium (DMEM supplemented with 20% FBS, 10% HS, 1% P/S and 1% chicken embryo extract) can be replaced every third day.</li>
	<li>Alternatively, if expansion and passing is desired follow the next steps:
	<ol>
		<li>Thaw an aliquot extracellular matrix gel (500 &#181;l) at 4 &#176;C for at least 1.5 hr. Dilute 1:10 in DMEM and follow the recommendations in point 1.1.1.</li>
		<li>Pre-chill a 10 ml pipette for 10 min at 4 &#176;C.</li>
		<li>Transfer three T75 flasks onto a cold surface (e.g. a freezer pack) for 10 min.</li>
		<li>Use the pre-chilled pipette to put 1 ml extracellular matrix gel into each flask. Check that the surface is covered completely. Keep the flasks on the cold surface for at least another 7 min (Figure 1A).</li>
		<li>Completely remove the remaining extracellular matrix gel with a 10 ml pipette, and dry the wells at 37 &#176;C for 1 hr.</li>
		<li>After counting, resuspend the freshly isolated SCs in 10 ml of culture medium (DMEM supplemented with 20% FBS, 10% HS, 1% P/S and 1% chicken embryo extract) and seed in the pre-coated T75 flasks.</li>
		<li>After three days, change the medium (and every third day) until 80% confluence is reached. For passaging, wash the T75 flasks three times with PBS. Next add 1 ml 0.25% trypsin solution and incubate for three min at 37 &#176;C. Resuspend in 9 ml of culture medium (DMEM supplemented with 20% FBS, 10% HS, 1% P/S and 1% chicken embryo extract) and centrifuge at 200 x g for 5 min. Discard the supernatant. After counting, resuspend 1 x 106 cells in 1,000 &#181;l of culture medium and freeze the cells.</li>
	</ol>
	</li>
</ol>

The authors have no conflicting interests to disclose.

SCs from different branchiomeric head muscles were isolated from one 9-week-old Wistar rat and cultured directly on extracellular matrix gel spots without prior expansion and passaging. After isolation, the cells were counted and seeded at the same cell density. For the parallel isolation of three different muscles, this method takes about 4 hr. To avoid culture contamination, a critical step is the rapid washing in alcohol 70% after dissection of the muscles.
During SC isolation it is important to cut the muscle tissue into small pieces (about 2 mm) but avoid too much mincing as this will result in a small cell yield because of cell damage. Also, the duration of the enzymatic digestion must be checked carefully under the microscope to avoid further damage. The aim of the digestion is to dissociate the myofibers. Since more than 90% of the isolated cells express Pax7, no further purification is required (Figures 6-8). This avoids extra purification steps in other methods such as pre-plating on uncoated dishes14,37,38, fractionation on Percoll39,40, or fluorescent- or magnetic cell sorting41,43. For trituration it is essential to induce shear between the tissue fragments and the opening of the pipette tip as this allows the mechanical release of the SCs. If the trituration with a 10 ml pipette (inside diameter tip: 1 mm) is difficult, a 5 ml (inside diameter tip: 2 mm) pipette can be used first. Alternatively, glass Pasteur pipettes can be cut at the desired diameter and be used. This method is simple, efficient and allows the simultaneous isolation of SC from different muscle samples.
The culture plates for SCs can also be coated with gelatin or collagen, but our previous studies show that extracellular matrix gel is far better for the maintenance of the myogenic potential than collagen38. The extracellular matrix gel spots of millimeter size (10 &#181;l/&#216; 2 mm or 20 &#181;l/&#216; 4 mm) allows the study of proliferation and differentiation of SCs with limited numbers of cells. For the differentiation assay about 8 to 20 times fewer cells are required compared to a 24-well plate (&#216; 15.6 mm), and about 80 to 200 times fewer compared to 35 mm Petri dishes (&#216; 35 mm)14,38.
Since extracellular matrix gel is expensive, this method is also more cost-efficient. In addition, the chamber slides can be replaced by plastic cover slips to further reduce the costs. For the preparation of the extracellular matrix gel spots overnight drying of the chamber slides is essential. As the extracellular matrix gel spots are transparent, it is necessary to mark the spots at the bottom side using back lighting. The chambers slides are fixed in a Petri dish for easy manipulation. Further cell culture expansion is not necessary, which offers the possibility to study the SCs of smaller muscles or small muscle samples. Alternatively, e.g. for PCR or muscle constructs if more cells are needed, the freshly isolated SCs can first be expanded in T75 flasks as indicated above.
SCs isolated using this protocol are not suitable for further purification with flow cytometry immediately after isolation. The digestion with pronase causes extensive digestion of the surface antigens14. Horse serum and fetal bovine serum that are used for cell culture must first be properly characterized before isolation, as different lot numbers differentially affected myoblasts proliferation and differentiation.
In recent years, there is a growing interest in the muscles derived from the branchial arches and the head mesoderm (e.g. the extraocular muscles)24. It has been clearly demonstrated that head and limb muscles possess highly different properties. Masseter muscle from old animals seems to retain their regenerative capacity in comparison with limb muscles25,26. SCs from the extraocular muscles possess a robust proliferation and differentiation capacity comparable to SCs from head muscles, and show a larger engraftment potential than limb muscle SCs24.
The fiber type distribution and myosin composition varies among muscle groups and also between species. Muscles originating from the first branchial arch in humans contain both slow and fast fibers (subtypes IIA and IIX), neonatal myosins and myosins typical for developing cardiac muscle. In rodents these muscles contain about 95% fast fibers myosin IIA and IIb)44-46. Studies on avian muscles show that SCs from different muscle fiber types vary in differentiation capacity. SCs from fast fibers only differentiate into fast muscle fibers, while SCs from slow fibers can differentiate into both fiber types47. In addition, the percentage of SCs in fast muscle fibers is lower than in slow muscle fibers48,49. This indicates that the fiber type distribution must be taken into account for studies on muscles in the craniofacial area. Similar to cleft palate muscles, the LVP in rodents contains almost exclusively fast fibers50. For that reason, SCs from the LVP are suitable for pre-clinical studies in the field of cleft palate.
This protocol offers new possibilities to study SCs derived from branchiomeric head muscles or other smaller muscles or smaller muscles samples. This will facilitate the development of new therapies to improve the regeneration of muscles in the maxillofacial area in conditions such as cleft palate but also in other conditions affecting smaller muscles.

An erratum was issue for <a href="http://www.jove.com/video/52802">Isolation and Characterization of Satellite Cells from Rat Head Branchiomeric Muscles</a>. The fourth figure was updated to explain the isolation of the LVP better.

Erratum: Isolation and Characterization of Satellite Cells from Rat Head Branchiomeric Muscles

About 1:500 to 1:1,000 newborns exhibit a cleft involving the lip and/or palate (CLP); thus this is the most common congenital malformation in humans1. The muscles of the soft palate are critical for the functioning of the soft palate during speech, swallowing, and sucking. If a cleft of the soft palate is present, these muscles are abnormally inserted into the posterior end of the palatal bone.
The soft palate moves up and down during speech, preventing air to escape through the nose. Children with a cleft in the palate do not have this control function resulting in a phenomenon known as velopharyngeal dysfunction2,3. Although the treatment protocols are variable, surgical repair of the soft palate takes place in early childhood (6-36 months of age)4. The abnormally inserted muscles of the soft palate can be surgically corrected5-7, however, velopharyngeal dysfunction persists in 7% to 30% of the patients2,3,8-10.
The ability of skeletal muscle to regenerate through the action of satellite cells (SCs) is well established11,12. Upon muscle injury, SCs are activated and migrate to the site of injury. They then proliferate, differentiate, and fuse to form new myofibers or repair damaged ones13. Quiescent SCs express the transcription factor Pax714,15, while their progeny, the proliferating myoblasts, additionally express the myogenic determination factor 1 (MyoD)16. Differentiating myoblasts start to express myogenin (MyoG)17. The terminal differentiation of myoblasts is marked by the formation of myofibers, and the expression of muscle-specific proteins such as myosin heavy chain (MyHC)16,18.
Recently, several strategies have been used in regenerative medicine to improve muscle regeneration of limb muscles19-23. Specific studies on branchiomeric head muscles are also important because it was recently demonstrated that they differ from other muscles in several aspects24. In contrast with limb muscles, it has been suggested that branchiomeric head muscles contain less SCs25, regenerate slower, and more fibrous connective tissue is formed after injury26 In addition, proliferating SCs from branchiomeric head muscles also express other transcription factors. For instance, Tcf21, a transcription factor for craniofacial muscle formation is strongly expressed in regenerating head muscles but hardly in regenerating limb muscles25. The muscles in the soft palate of CLP patients are usually smaller and less well-organized compared to normal palatal muscles27,28. Slow and fast fibers are both present in the soft palate muscles but the slow fibers are more abundant. In contrast, cleft muscles contain a higher proportion of fast fibers and also a reduced capillary supply compared with normal soft palate muscles29-31. Fast fibers are more prone to contraction-induced injury31-33. The accompanying poor capillary supply may also promote fibrosis34,35. All these aspects may contribute to the poor regeneration of soft palate muscles after surgical cleft closure36. In view of this, a protocol for the isolation and characterization of branchiomeric head muscle SCs is crucial. This provides the possibility to study SC biology of branchiomeric head muscles. In addition, new therapies based on tissue engineering can be developed to promote muscle regeneration after surgery in CLP and other conditions compromising the craniofacial area.
In general, SCs can be obtained after dissociation of muscle tissue14. Mincing, enzymatic digestion, and trituration are generally required to release SCs from their niche. SCs can be purified by pre-plating on uncoated dishes14,37,38, fractionation on Percoll39,40, or fluorescent- or magnetic cell sorting41-43. Here we present a new economic and rapid protocol for the isolation of satellite cells from branchiomeric head muscles of young adult rats. This protocol is based on a previous manuscript14 and specifically adapted for small tissue samples. The isolation of SCs from representative muscles originating from the 1st, 2nd, and 4th branchial arches are described. After isolation, low numbers of satellite cells are cultured on extracellular matrix gel spots of millimeter size to study their differentiation. This approach avoids the requirement for the expansion and passaging of SCs.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Hypodermic Needle 25G 0,5x25m</td>
			<td>BD Microlance</td>
			<td>300400</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting scissors</td>
			<td>Braun</td>
			<td>BC154R</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro forceps straight</td>
			<td>Braun</td>
			<td>BD330R</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Scalpel Blade No.15</td>
			<td>Swann-Morton</td>
			<td>0205</td>
			<td></td>
		</tr>
		<tr>
			<td>Alcohol 70%</td>
			<td>Denteck</td>
			<td>2,010,005</td>
			<td></td>
		</tr>
		<tr>
			<td>Permanox Slide, 8 Chamber</td>
			<td>Thermo Scientific</td>
			<td>177445</td>
			<td></td>
		</tr>
		<tr>
			<td>6 well cell culture plate</td>
			<td>Greiner bio-one</td>
			<td>657160</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Culture Dishes (100 x 20 mm)</td>
			<td>Greiner bio-one</td>
			<td>664160</td>
			<td></td>
		</tr>
		<tr>
			<td>15 ml sterile conical centrifuge tube</td>
			<td>BD Biosciences</td>
			<td>352097</td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml sterile conical centrifuge tube</td>
			<td>BD Biosciences</td>
			<td>352098</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell strainer (40 &#956;m)</td>
			<td>Gibco</td>
			<td>431750</td>
			<td></td>
		</tr>
		<tr>
			<td>10 mL serological pipette</td>
			<td>Greiner bio-one</td>
			<td>607180</td>
			<td></td>
		</tr>
		<tr>
			<td>20&#181;L FT20</td>
			<td>Greiner bio-one</td>
			<td>774288</td>
			<td></td>
		</tr>
		<tr>
			<td>Matrigel, Phenol-Red Free</td>
			<td>BD Biosciences</td>
			<td>356237</td>
			<td>10 mL</td>
		</tr>
		<tr>
			<td>Pronase</td>
			<td>Calbiochem</td>
			<td>53702</td>
			<td>10KU</td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline</td>
			<td>Gibco</td>
			<td>14190-144</td>
			<td>500 mL</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium, high glucose, GlutaMAX Supplement, pyruvate&#160;</td>
			<td>Gibco</td>
			<td>10569-010</td>
			<td>500 mL</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum&#160;&#160;</td>
			<td>Fisher Scientific&#160;</td>
			<td>3600511</td>
			<td>500 mL</td>
		</tr>
		<tr>
			<td>Horse Serum</td>
			<td>Gibco</td>
			<td>26050088</td>
			<td>500 mL</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (10,000 U/mL)</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td>100 mL</td>
		</tr>
		<tr>
			<td>Chicken Embryo Extract</td>
			<td>MP Biomedicals</td>
			<td>2850145</td>
			<td>20 mL</td>
		</tr>
	</tbody>
</table>

isolation and characterization of satellite cells from rat head branchiomeric muscles

Fibrosis and defective muscle regeneration can hamper the functional recovery of the soft palate muscles after cleft palate repair. This causes persistent problems in speech, swallowing, and sucking. In vitro culture systems that allow the study of satellite cells (myogenic stem cells) from head muscles are crucial to develop new therapies based on tissue engineering to promote muscle regeneration after surgery. These systems will offer new perspectives for the treatment of cleft palate patients. A protocol for the isolation, culture and differentiation of satellite cells from head muscles is presented. The isolation is based on enzymatic digestion and trituration to release the satellite cells. In addition, this protocol comprises an innovative method using extracellular matrix gel coatings of millimeter size, which requires only low numbers of satellite cells for differentiation assays.

Using this protocol, the masseter muscle (one side) yields 0.8-1 x 106 cells, the digastric muscle (posterior belly) yields 1.5-2 x 105 cells, and levator veli palatini muscle yields 1-1.5 x 105 cells. Cell yields depend on the muscle type, strain, and age of the animal. For comparison between the three muscle groups, freshly isolated SCs were seeded at the same cell density (1.5 x 103/10 &#181;l). Directly after isolation, more than 90% of the freshly isolated cells express Pax7 (Figure 6).
Day 4, 7 and 10 cultures were stained with antibodies against Pax7, MyoD, MyoG and MyHC immunostaining. Five arbitrary fields were counted per culture using a 20X objective. At day 4 Pax 7 and Myo D is expressed in all muscle groups (Figures 6 and 7 and 8), however the progeny of SatCs from the masseter and digastric muscles start expressing myogenin earlier than the levator veli palatini muscle (Figure 9). At day 10, the expression of MyoG is strongly reduced in all groups (Figure 9). A few days after seeding on the extracellular matrix gel spots, the proliferating cells begin to fuse and form multi-nucleated myotubes, which express myosin heavy chain. Small myotubes are clearly visible at day 7 (Figure 10). At day 10, twitching of the myotubes can be observed (Video 1).
<img alt="Figure 1" src="/files/ftp_upload/52802/52802fig1.jpg" /> 
Figure 1: Extracellular matrix gel spots in a chamber slide. (A) For easy manipulation, place the 8-well chamber slide into a 100 mm Petri dish. Pipet 10 &#181;l extracellular matrix gel in each chamber and put it on a cold surface (7 min). (B) Chamber slide after the excess extracellular matrix gel is removed.
<img alt="Figure 2" src="/files/ftp_upload/52802/52802fig2.jpg" /> 
Figure 2: Dissection of the masseter muscle. (A) Head of the animal in a lateral view. Ear (E), Parotid gland (P) and facial nerve (VII). (B) Tendinous aponeurosis (Te) of the superficial head of the masseter muscle (Ms) and temporal muscle (T). Separate the tendon from its insertion with a forceps. (C) Carefully dissect the muscle until its insertion at the ramus of the mandible. E: ear, P: parotid gland, VII: facial nerve, T: Temporal muscle, Ms: superficial head of the masseter muscle, Te: tendon, Mp: deep head of the masseter muscle.
<img alt="Figure 3" src="/files/ftp_upload/52802/52802fig3.jpg" /> 
Figure 3: Dissection of the posterior belly of the digastric muscle. (A) Head of the animal in a supine position. Localize the submandibular gland (Sg), masseter muscle (M), facial nerve (VII) and sternocleidomastoid muscle (SCM). Remove the submandibular gland. (B) Localize the digastric muscle anterior (AD) and posterior belly (PD). With a straight forceps, take the anterior tendon of the posterior belly, cut it and dissect it carefully until its origin in the tympanic bulla (ty). E: ear, Sg: submandibular gland, VII: facial nerve, M: masseter muscle, SMC: sternocleidomastoid muscle, AD: anterior belly digastric muscle, PD: posterior belly digastric muscle, Ty: Tympanic bulla.
<img src="/files/ftp_upload/52802/52802fig4new.jpg" align="Figure 4" /> Figure 4: Dissection of the levator veli palatini muscle. (A) General view after dissection of the digastric muscle (posterior belly). Stylohyoid muscle (St) and tendon of the levator veli palatini can be localized. Note the trachea (T) and esophagus (Es) running behind it. (B) After lifting the trachea and the esophagus the pharynx (P) is exposed. The levator veli palatini that runs laterally towards the soft palate is now visible. The arrow indicates the dissected superior pharyngeal constrictor muscle; note the levator veli palatini muscles at both sides. E: ear, St: stylohyoid muscle, VII: facial nerve, M: masseter muscle, AD: anterior belly digastric muscle, PD: posterior belly digastric muscle, T: trachea, Es: esophagus, P: Pharynx, *levator veli palatini muscle.
<img alt="Figure 5" src="/files/ftp_upload/52802/52802fig5.jpg" /> 
Figure 5: Appearance of the muscle tissue (A) before and (B) after enzymatic digestion with pronase. Note that muscle bundles appear to be loosened after enzymatic digestion.
<img alt="Figure 6" src="/files/ftp_upload/52802/52802fig6highres.jpg" width="350" /> 
Figure 6: Pax 7 immunostaining. Freshly isolated SCs, applied to extracellular matrix gel at the end of isolation (about 6 hours after initial tissue digestion). Five arbitrary fields were counted using a 10X objective with an average of 210 cells per field. Approximately 90% of the cells are Pax 7 positive. DAPI: blue, Pax7: red. Scale bar, 100 &#181;m.
<img alt="Figure 7" src="/files/ftp_upload/52802/52802fig7highres.jpg" width="700" /> 
Figure 7: Pax 7, MyoD immunostaining. Day 4, 7 and 10 cultures were stained with antibodies against Pax7, and MyoD immunostaining. (A&#8211;C) and (D&#8211;F) Representative photomicrographs of day 4 and 7 cultures from the masseter muscle. (G and H) The number of Pax7+ and MyoD+ nuclei per microscopic field was counted and expressed as a percentage of the total number of nuclei (DAPI). DAPI: blue, Pax7: red, and MyoD: green. Scales bar, 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/52802/52802fig7highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" src="/files/ftp_upload/52802/52802fig8highres.jpg" width="700" /> 
Figure 8: Distribution of Pax7&#177;/MyoD&#177; in cultures from mononucleated cells in cultures from masseter, digastric and levator veli palatine muscle. (A&#8211;C) Day 4, 7 and 10 cultures were stained with antibodies against Pax7, and MyoD immunostaining. The total number of cells is based on of the total number of nuclei (DAPI). (D) Data quantification of Pax7&#177;/MyoD&#177; cells. <a href="https://www.jove.com/files/ftp_upload/52802/52802fig8highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" src="/files/ftp_upload/52802/52802fig9highres.jpg" width="700" /> 
Figure 9: Myogenin immunostaining. Day 4, 7 and 10 cultures were stained with antibodies against Myogenin. (A&#8211;D) Representative photomicrographs of day 4 and 7 cultures from the levator veli palatine muscle. (E) The number of MyoG+ nuclei per microscopic field was counted and expressed as a percentage of the total number of nuclei (DAPI). (F) Data quantification of MyoG+ cells. DAPI: blue, Myogenin: green. Scales bar, 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/52802/52802fig9highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" src="/files/ftp_upload/52802/52802fig10.jpg" /> 
Figure 10: Myosin Heavy Chain immunostaining. Day 4, 7 and 10 cultures were stained with antibodies against myosin heavy chain (MyHC). Representative photomicrographs of day 4, 7 and 10 cultures from the digastric (DIG) muscle. At day 7, small myotubes are present while at day 10 long and well-organized myotubes are evident. Scales bar, 200 &#181;m. <a href="https://www.jove.com/files/ftp_upload/52802/52802fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Myotube twitching. Examples of two representative fields with twitching myotubes are shown for day 10 cultures from digastric muscle. <a href="https://www.jove.com/files/ftp_upload/52802/Video1.mpg" target="_blank">Please click here to view this video.</a>

Watch this Scientific Journal Video about Isolation and Characterization of Satellite Cells from Rat Head Branchiomeric Muscles at JoVE.com

Isolation and Characterization of Satellite Cells from Rat Head Branchiomeric Muscles

This protocol describes the isolation of satellite cells from branchiomeric head muscles of a 9 week-old rat. The muscles originate from different branchial arches. Subsequently, the satellite cells are cultured on a spot coating of millimeter size to study their differentiation. This approach avoids the expansion and passaging of satellite cells.

Fibrosis and defective muscle regeneration can hamper the functional recovery of the soft palate muscles after cleft palate repair. This causes persistent ...

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Developmental Biology

10.8K Views.  Radboud University Medical Center. This protocol describes the isolation of satellite cells from branchiomeric head muscles of a 9 week-old rat. The muscles originate from different branchial arches. Subsequently, the satellite cells are cultured on a spot coating of millimeter size to study their differentiation. This approach avoids the expansion and passaging of satellite cells.

Video: Isolation and Characterization of Satellite Cells from Rat Head Branchiomeric Muscles

Isolation and Quantitative Immunocytochemical Characterization of Primary Myogenic Cells and Fibroblasts from Human Skeletal Muscle

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated from human muscle biopsy samples using enzymatic digestion and their myogenic properties studied in culture. Quantitatively, the two main adherent cell types obtained from enzymatic digestion are: (i) the satellite cells (termed myogenic cells or muscle precursor cells), identified initially as CD56+ and later as CD56+/desmin+ cells and (ii) muscle-derived fibroblasts, identified as CD56&#8211; and TE-7+. Fibroblasts proliferate very efficiently in culture and in mixed cell populations these cells may overrun myogenic cells to dominate the culture. The isolation and purification of different cell types from human muscle is thus an important methodological consideration when trying to investigate the innate behavior of either cell type in culture. Here we describe a system of sorting based on the gentle enzymatic digestion of cells using collagenase and dispase followed by magnetic activated cell sorting (MACS) which gives both a high purity (&#62;95% myogenic cells) and good yield (~2.8 x 106 &#177; 8.87 x 105 cells/g tissue after 7 days in vitro) for experiments in culture. This approach is based on incubating the mixed muscle-derived cell population with magnetic microbeads beads conjugated to an antibody against CD56 and then passing cells though a magnetic field. CD56+ cells bound to microbeads are retained by the field whereas CD56&#8211; cells pass unimpeded through the column. Cell suspensions from any stage of the sorting process can be plated and cultured. Following a given intervention, cell morphology, and the expression and localization of proteins including nuclear transcription factors can be quantified using immunofluorescent labeling with specific antibodies and an image processing and analysis package.

The main adherent cell types derived from human muscle are myogenic cells and fibroblasts. Here, cell populations are enriched using magnetic-activated cell sorting based on the CD56 antigen. Subsequent immunolabelling with specific antibodies and use of image analysis techniques allows quantification of cytoplasmic and nuclear characteristics in individual cells.

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated ...

Isolation of Neural Stem/Progenitor Cells from the Periventricular Region of the Adult Rat and Human Spinal Cord

Adult rat and human spinal cord neural stem/progenitor cells (NSPCs) cultured in growth factor-enriched medium allows for the proliferation of multipotent, self-renewing, and expandable neural stem cells. In serum conditions, these multipotent NSPCs will differentiate, generating neurons, astrocytes, and oligodendrocytes. The harvested tissue is enzymatically dissociated in a papain-EDTA solution and then mechanically dissociated and separated through a discontinuous density gradient to yield a single cell suspension which is plated in neurobasal medium supplemented with epidermal growth factor (EGF), basic fibroblast growth factor (bFGF), and heparin. Adult rat spinal cord NSPCs are cultured as free-floating neurospheres and adult human spinal cord NSPCs are grown as adherent cultures. Under these conditions, adult spinal cord NSPCs proliferate, express markers of precursor cells, and can be continuously expanded upon passage. These cells can be studied in vitro in response to various stimuli, and exogenous factors may be used to promote lineage restriction to examine neural stem cell differentiation. Multipotent NSPCs or their progeny can also be transplanted into various animal models to assess regenerative repair.

The adult mammalian spinal cord contains neural stem/progenitor cells (NSPCs) that can be isolated and expanded in culture. This protocol describes the harvesting, isolation, culture, and passaging of NSPCs generated from the periventricular region of the adult spinal cord from the rat and from human organ transplant donors.

Adult rat and human spinal cord neural stem/progenitor cells (NSPCs) cultured in growth factor-enriched medium allows for the proliferation of ...

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Isolation and Characterization of Mesenchymal Stromal Cells from Human Umbilical Cord and Fetal Placenta

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly gained attention because they do not pose any ethical or moral concerns. Current methods to isolate MSCs from UC yield low amounts of cells with variable proliferation potentials. Since UC is an anatomically-complex organ, differences in MSC properties may be due to the differences in the anatomical regions of their isolation. In this study, we first dissected the cord/placenta samples into three discrete anatomical regions: UC, cord-placenta junction (CPJ), and fetal placenta (FP). Second, two distinct zones, cord lining (CL) and Wharton&#39;s jelly (WJ), were separated. The explant culture technique was then used to isolate cells from the four sources. The time required for the primary culture of cells from the explants varied depending on the source of the tissue. Outgrowth of the cells occurred within 3 - 4 days of the CPJ explants, whereas growth was observed after 7 - 10 days and 11 - 14 days from CL/WJ and FP explants, respectively. The isolated cells were adherent to plastic and displayed fibroblastoid morphology and surface markers, such as CD29, CD44, CD73, CD90, and CD105, similarly to bone marrow (BM)-derived MSCs. However, the colony-forming efficiency of the cells varied, with CPJ-MSCs and WJ-MSCs showing higher efficiency than BM-MSCs. MSCs from all four sources differentiated into adipogenic, chondrogenic, and osteogenic lineages, indicating that they were multipotent. CPJ-MSCs differentiated more efficiently in comparison to other MSC sources. These results suggest that the CPJ is the most potent anatomical region and yields a higher number of cells, with greater proliferation and self-renewal capacities in vitro. In conclusion, the comparative analysis of the MSCs from the four sources indicated that CPJ is a more promising source of MSCs for cell therapy, regenerative medicine, and tissue engineering.

Here, we present a protocol for the dissection of human umbilical cord (UC) and fetal placenta sample into cord lining (CL), Wharton&#39;s jelly (WJ), cord-placenta junction (CPJ), and fetal placenta (FP) for the isolation and characterization of mesenchymal stromal cells (MSCs) using the explant culture technique.

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly ...

Isolation of Type I and Type II Pericytes from Mouse Skeletal Muscles

Pericytes are perivascular multipotent cells that show heterogeneity in different organs or even within the same tissue. In skeletal muscles, there are at least two pericyte subpopulations (called type I and type II), which express different molecular markers and have distinct differentiation capabilities. Using NG2-DsRed and Nestin-GFP double-transgenic mice, type I (NG2-DsRed+Nestin-GFP-) and type II (NG2-DsRed+Nestin-GFP+) pericytes have been successfully isolated. However, the availability of these double-transgenic mice prevents the widespread use of this purification method. This work describes an alternative protocol that allows for the easy and simultaneous isolation of type I and type II pericytes from skeletal muscles. This protocol utilizes the fluorescence-activated cell sorting (FACS) technique and targets PDGFR&#946;, rather than NG2, together with the Nestin-GFP signal. Following isolation, type I and type II pericytes show distinct morphologies. In addition, type I and type II pericytes isolated with this new method, like those isolated from the double-transgenic mice, are adipogenic and myogenic, respectively. These results suggest that this protocol can be used to isolate pericyte subpopulations from skeletal muscles and possibly from other tissues.

This work describes a FACS-based protocol that allows for easy and simultaneous isolation of type I and type II pericytes from skeletal muscles.

Pericytes are perivascular multipotent cells that show heterogeneity in different organs or even within the same tissue. In skeletal muscles, there are at ...

Isolation and Characterization of Primary Rat Valve Interstitial Cells: A New Model to Study Aortic Valve Calcification

Calcific aortic valve disease (CAVD) is characterized by the progressive thickening of the aortic valve leaflets. It is a condition frequently found in the elderly and end-stage renal disease (ESRD) patients, who commonly suffer from hyperphosphatemia and hypercalcemia. At present, there are no medication therapies that can stop its progression. The mechanisms that underlie this pathological process remain unclear. The aortic valve leaflet is composed of a thin layer of valve endothelial cells (VECs) on the outer surfaces of the aortic cusps, with valve interstitial cells (VICs) sandwiched between the VECs. The use of a rat model enables the in vitro study of ectopic calcification based on the in vivo physiopathological serum phosphate (Pi) and calcium (Ca) levels of patients who suffer from hyperphosphatemia and hypercalcemia. The described protocol details the isolation of a pure rat VIC population as shown by the expression of VIC markers: alpha-smooth muscle actin (&#945;-SMA) vimentin and tissue growth factor beta (TGF&#946;) 1 and 2, and the absence of cluster of differentiation (CD) 31, a VEC marker. By expanding these VICs, biochemical, genetic, and imaging studies can be performed to study and unravel the key mediators underpinning CAVD.

This protocol describes the isolation, culture, and calcification of rat-derived valve interstitial cells, a highly physiological in vitro model of calcific aortic valve disease (CAVD). Exploitation of this rat model facilitates CAVD research in exploring the cell and molecular mechanisms that underlie this complex pathological process.

Calcific aortic valve disease (CAVD) is characterized by the progressive thickening of the aortic valve leaflets. It is a condition frequently found in ...

Identification of Skeletal Muscle Satellite Cells by Immunofluorescence with Pax7 and Laminin Antibodies

Immunofluorescence is an effective method that helps to identify different cell types on tissue sections. In order to study the desired cell population, antibodies for specific cell markers are applied on tissue sections. In adult skeletal muscle, satellite cells (SCs) are stem cells that contribute to muscle repair and regeneration. Therefore, it is important to visualize and trace the satellite cell population under different physiological conditions. In resting skeletal muscle, SCs reside between the basal lamina and myofiber plasma membrane. A commonly used marker for identifying SCs on the myofibers or in cell culture is the paired box protein Pax7. In this article, an optimized Pax7 immunofluorescence protocol on skeletal muscle sections is presented that minimizes non-specific staining and background. Another antibody that recognizes a protein (laminin) of the basal lamina was also added to help identify SCs. Similar protocols can also be used to perform double or triple labeling with Pax7 and antibodies for additional proteins of interest.

The precise identification of satellite cells is essential for studying their functions under various physiological and pathological conditions. This article presents a protocol to identify satellite cells on adult skeletal muscle sections by immunofluorescence-based staining.

Immunofluorescence is an effective method that helps to identify different cell types on tissue sections. In order to study the desired cell population, ...

Isolation and Characterization of Cardiac Mesenchymal Stromal Cells from Endomyocardial Bioptic Samples of Arrhythmogenic Cardiomyopathy Patients

A normal adult heart is composed of several different cell types, among which cardiac mesenchymal stromal cells represent an abundant population. The isolation of these cells offers the possibility of studying their involvement in cardiac diseases, and, in addition, provides a useful primary cell model to investigate biological mechanisms.
Here, the method for the isolation of C-MSC from arrhythmogenic cardiomyopathy patients&#39; bioptic samples is described. The endomyocardial biopsy sampling is guided in the right ventricular areas adjacent to the scar visualized by electro-anatomical mapping. The digestion of the biopsies in collagenase and their plating on a plastic dish in culture medium to allow C-MSC growth is described. The isolated cells can be expanded in culture for several passages. To confirm their mesenchymal phenotype, the description of immuno-phenotypical characterization is provided. C-MSC are able to differentiate into several cell types like adipocytes, chondrocytes, and osteoblasts: in the context of ACM, characterized by adipocyte deposits in patients&#39; hearts, the protocols for the adipogenic differentiation of C-MSC and the characterization of lipid droplet accumulation are described.

In this article, a method to isolate cardiac mesenchymal stromal cells from endomyocardial bioptic samples of arrhythmogenic cardiomyopathy patients is provided. Their characterization and the protocol to boost their adipogenic differentiation are described.

A normal adult heart is composed of several different cell types, among which cardiac mesenchymal stromal cells represent an abundant population. The ...

Isolation and Characterization of Human Umbilical Cord-derived Mesenchymal Stem Cells from Preterm and Term Infants

Mesenchymal stem cells (MSCs) have considerable therapeutic potential and attract increasing interest in the biomedical field. MSCs are originally isolated and characterized from bone marrow (BM), then acquired from tissues including adipose tissue, synovium, skin, dental pulp, and fetal appendages such as placenta, umbilical cord blood (UCB), and umbilical cord (UC). MSCs are a heterogeneous cell population with the capacity for (1) adherence to plastic in standard culture conditions, (2) surface marker expression of CD73+/CD90+/CD105+/CD45-/CD34-/CD14-/CD19-/HLA-DR- phenotypes, and (3) trilineage differentiation into adipocytes, osteocytes, and chondrocytes, as currently defined by the International Society for Cellular Therapy (ISCT). Although BM is the most widely used source of MSCs, the invasive nature of BM aspiration ethically limits its accessibility. Proliferation and differentiation capacity of MSCs obtained from BM generally decline with the age of the donor. In contrast, fetal MSCs obtained from UC have advantages such as vigorous proliferation and differentiation capacity. There is no ethical concern for UC sampling, as it is typically regarded as medical waste. Human UC starts to develop with continuing growth of the amniotic cavity at 4-8 weeks of gestation and keeps growing until reaching 50-60 cm in length, and it can be isolated during the whole newborn delivery period. To gain insight into the pathophysiology of intractable diseases, we have used UC-derived MSCs (UC-MSCs) from infants delivered at various gestational ages. In this protocol, we describe the isolation and characterization of UC-MSCs from fetuses/infants at 19-40 weeks of gestation.

Human umbilical cord (UC) can be obtained during the perinatal period as a result of preterm, term, and postterm delivery. In this protocol, we describe the isolation and characterization of UC-derived mesenchymal stem cells (UC-MSCs) from fetuses/infants at 19-40 weeks of gestation.

Mesenchymal stem cells (MSCs) have considerable therapeutic potential and attract increasing interest in the biomedical field. MSCs are originally ...

In Vitro Generation of Somite Derivatives from Human Induced Pluripotent Stem Cells

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give rise to multiple cell types, including the myotome (MYO), sclerotome (SCL), dermatome (D), and syndetome (SYN), which in turn develop into skeletal muscle, axial skeleton, dorsal dermis, and axial tendon/ligament, respectively. Therefore, the generation of SMs and their derivatives from human induced pluripotent stem cells (iPSCs) is critical to obtain pluripotent stem cells (PSCs) for application in regenerative medicine and for disease research in the field of orthopedic surgery. Although the induction protocols for MYO and SCL from PSCs have been previously reported by several researchers, no study has yet demonstrated the induction of SYN and D from iPSCs. Therefore, efficient induction of fully competent SMs remains a major challenge. Here, we recapitulate human SM patterning with human iPSCs in vitro by mimicking the signaling environment during chick/mouse SM development, and report on methods of systematic induction of SM derivatives (MYO, SCL, D, and SYN) from human iPSCs under chemically defined conditions through the presomitic mesoderm (PSM) and SM states. Knowledge regarding chick/mouse&#160;SM development was successfully applied to the induction of SMs with human iPSCs. This method could be a novel tool for studying human somitogenesis and patterning without the use of embryos and for cell-based therapy and disease modeling.

We present here a protocol for the differentiation of human induced pluripotent stem cells into each somite derivative (myotome, sclerotome, dermatome, and syndetome) in chemically defined conditions, which has applications in future disease modeling and cell-based therapies in orthopedic surgery.

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give ...