This work was supported by The Myositis Association and the Peter Buck Foundation. We would like to thank Dr. Yuanfan Zhang for sharing her expertise and training in the xenograft surgical technique.

All use of research specimens from human subjects was approved by the Johns Hopkins Institutional Review Board (IRB) to protect the rights and welfare of the participants. All animal experiments were approved by the Johns Hopkins University Institutional Animal Care and Use Committee (IACUC) in accordance with the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals. Male NOD-Rag1null IL2rγnull (NRG) host mice (8-12 weeks old) are used to carry out xenograft experiments. These mice are housed in ventilated racks and are given HEPA-filtered, tempered, and humidified air as well as reverse osmosis filtered hyperchlorinated water. Mice are provided water and an irradiated antibiotic diet (Table of Materials) ad libitum, and the facility provides 14 h of light to 10 h of dark as controlled by central timer.
1. Equipment Preparation
<ol>
	<li>Acquire NOD-Rag1null IL2r γnull (NRG) mice, 8-12 weeks of age.</li>
	<li>Autoclave surgical equipment: scissors, forceps, needle holder, surgical stapler (Table of Materials), wound clips, surgical wipes (Table of Materials), and beaker (Figure 1A).</li>
	<li>Prepare 50 mL of muscle media (20% fetal bovine serum, 2% chick embryo extract, 1% antibiotic/antimycotic in Hams F10 Medium). Keep all chemicals/drugs/solutions used for surgery at room temperature unless stated differently in the protocol.</li>
	<li>Prepare a 1 mL syringe with a 26 G needle that is 3/8 inches long containing 2 mg/mL analgesic (Table of Materials), and place on ice. The analgesic can be diluted to the proper concentration using sterile phosphate buffered saline (PBS).</li>
</ol>
2. Surgical Preparation
<ol>
	<li>Obtain a human muscle biopsy under an IRB-approved protocol from patients whose muscles display strength &gt; 4-/5 on the MRC (Medical Research Council) scale12. Place the research specimen in a 100 mm x 15 mm Petri dish containing muscle media. 
	NOTE: The MRC scale is used in clinical practice as an assessment of muscle strength with 0 showing no contraction, 5 showing normal power, and 4 (4- to 4+) showing movement against resistance12. We have found that muscles with mild to moderate weakness (MRC &gt; 4-/5) typically show disease pathology but are not extensively replaced by fatty tissue or fibrosis, both of which impede xenograft regeneration. In the case of autopsy tissue where a recent MRC score is not available, muscle quality can be accessed via gross observation. Muscle biopsies that are pale pink in appearance or have large areas of fatty tissue are not likely to xenograft successfully.</li>
	<li>Remove any remaining fascia or fatty tissue from the specimen with surgical scissors using a stereo microscope and light source to assist visualization.</li>
	<li>Dissect the muscle biopsy into approximately 7 mm x 3 mm x 3 mm pieces with surgical scissors using the stereo microscope and a light source. Ensure fibers are arranged longitudinally within the specimen.</li>
	<li>Place the Petri dish containing dissected muscle on ice. On average, the xenografts are kept in media for 4 hours while surgeries are being performed. However, biopsies have been stored in media for 24 h prior to xenografting, and this delay did not appear to negatively impact transplantation or regeneration.</li>
	<li>Place synthetic, non-absorbable sutures (Table of Materials) in a 100 mm x 15 mm Petri dish containing 70% ethanol.</li>
	<li>Set up a dual procedure anesthesia circuit: arrange the Mapleson E breathing circuit on the stereo microscope and place the induction chamber in a biosafety cabinet (Figure 1A,B).</li>
	<li>Obtain the weight of the NRG mouse by placing in an autoclaved beaker on a scale, and transfer to the induction chamber. Induce anesthesia under 3% isoflurane. Once the appropriate anesthetic depth is achieved—as assessed by observation of respiratory rate, muscle relaxation, and lack of voluntary movement—reduce the vaporizer setting to 1.5% for the remainder of the surgery.</li>
	<li>Transfer the mouse from the induction chamber to the Mapleson E breathing circuit and apply ophthalmic ointment to eyes.</li>
	<li>Remove hair overlying the tibialis anterior (TA) from ankle to knee with a trimmer, followed by a 1 min treatment with hair removal lotion (Table of Materials) (Figure 2A).</li>
	<li>Disinfect the surgical site by swabbing the leg with povidone-iodine solution. Then wash away the remaining povidone-iodine with 70% ethanol.</li>
	<li>Inject the mouse subcutaneously with analgesic, such as carprofen, (Table of Materials) at a dose of 5 mg/kg.</li>
</ol>
3. Xenograft Surgery
<ol>
	<li>Tape down the leg and make a straight incision over the tibilalis anterior (TA) muscle with scissors and iris forceps originating at the distal tendons and terminating below the knee (Figure 2B).</li>
	<li>Separate skin from muscle using blunt dissection with surgical scissors.</li>
	<li>Cut through the epimysium of the TA muscle with scissors starting at the tendon and ending at the knee. 
	NOTE: This is a very superficial cut (less than 0.5 mm; Figure 2B, black dashed line), and the underlying TA should not be damaged in the process as this would make removal more challenging. When performed correctly, the muscle fibers will visibly relax.</li>
	<li>Cut the distal tendon of the TA with scissors, grab the tendon with iris forceps, and pull the TA up toward the knee (Figure 2C).</li>
	<li>Cut the distal tendon of the extensor digitorum longus (EDL) with scissors and pull the EDL up toward the knee (Figure 2D). Once the proximal tendon of the peroneus longus (PL) muscle is visible, remove the EDL with scissors (Figure 2D, green dashed line).</li>
	<li>Remove the TA with scissors (Figure 2D, blue dashed line) and use a surgical wipe wetted with PBS and slight pressure to achieve hemostasis (Figure 2E).</li>
	<li>Thread a suture through proximal peroneus longus (PL) tendon and trim, leaving approximately 1.5 inch of thread on either side of the tendon (Figure 2F).</li>
	<li>Perform the first half of a two-hand surgical square knot, but do not tighten: this will form a circle. Place a xenograft in this circle and tighten the loop to secure the xenograft. Complete the other half of the square knot (Figure 2G,H). This will suture the xenograft to the proximal tendon of the PL.</li>
	<li>Thread suture through distal PL tendon and repeat the square knot technique from step 3.8 to tie the xenograft to the distal tendon (Figure 2H,I). 
	NOTE: The medial tarsal artery and vein can lie close to or on top of the distal tendon of the PL. Do not place sutures through or around these vessels. It is easy to tell if a suture has been improperly placed as vessels will blanch or bleed. If this occurs, remove the suture and place in a different location.</li>
	<li>Pull skin over xenografted muscle, seal with surgical glue, and place 2-3 surgical staples over the incision (Figure 2J).</li>
	<li>Place mouse in a clean cage on a heated pad to recover. Monitor mouse until fully conscious and periodically over the next few days for signs of local systemic infection and to ensure the surgical site is not reopened. 
	NOTE: A single dose of analgesic as described in step 2.11 is typically sufficient for pain relief. However, the mice should also be monitored for persistent pain (e.g. lameness, ruffled coat, hunched posture), and, if necessary, re-dosed with analgesic at 24 h post-operatively.</li>
</ol>
4. Xenograft Collection
NOTE: Xenografts are typically collected between 4 to 6 months post-surgery. However, collections have been performed up to 12 months post-surgery.
<ol>
	<li>Place a covered beaker containing 200 mL of 2-methylbutane in a box containing dry ice for a minimum of 30 min before xenograft collection.</li>
	<li>Induce anesthesia under 3% isoflurane in induction chamber. Once the appropriate anesthetic depth is achieved, reduce the vaporizer setting to 1.5% for the remainder of the surgery.</li>
	<li>Transfer the mouse from the induction chamber to the Mapleson E breathing circuit arranged on a stereo microscope.</li>
	<li>Remove hair overlying the tibialis anterior from ankle to knee with a trimmer and hair removal lotion. The sutures holding the xenograft in place can be seen through the skin (Figure 3A).</li>
	<li>Tape down the leg and use scissors and iris forceps to open skin over the xenograft until both sutures are visible (Figure 3B). Skin overlying the xenograft can be removed as shown to make removal of the xenograft easier.</li>
	<li>Use a scalpel to cut between the xenograft and the tibia (Figure 3B, arrow denotes initial site and direction of incision). This will free one side of the xenograft.</li>
	<li>Use a scalpel to cut between the PL muscle and the gastrocnemius muscle (Figure 3C, incision along epimysium labeled with arrow). The PL will be removed with the xenograft.</li>
	<li>Cut below the distal suture and through the distal tendon of the PL (Figure 3D, cut along dotted line).</li>
	<li>Remove the xenograft and PL by grabbing the suture with iris forceps and deflecting it toward the knee while using scissors to cut it away from the underlying muscle (Figure 3E).</li>
	<li>Cut above the proximal suture with scissors to remove the xenograft and PL (Figure 3F, cut along dotted line in 3E).</li>
	<li>Place the specimen on a small piece of cardboard or plastic, and pin as close to the sutures as possible. While pinning the specimen, gently stretch the muscle to ensure that the fiber orientation is maintained during the snap freezing process. After the pins are securely in place, slide the muscle up the pins so it rests just above the cardboard. 
	NOTE: Alternatively, one end of the xenograft can be mounted in tragacanth on a cork, or it can be submerged entirely in optimal cutting temperature (O.C.T.) compound in a cryomold. With care, muscle conformation can be retained with both methods.</li>
	<li>Snap freeze the xenograft in pre-cooled 2-methylbutane.</li>
	<li>Store xenograft at -80 °C.</li>
	<li>Immediately following xenograft collection, euthanize mice in accordance with American Veterinary Medical Association guidelines:
	<ol>
		<li>Place mice in a sealed chamber with an appropriate waste gas scavenging system. Use isoflurane at a concentration of 3-4% to induce anesthesia.</li>
		<li>Once the appropriate anesthetic depth is achieved—as assessed by observation of respiratory rate, muscle relaxation, and lack of voluntary movement—increase the vaporizer setting to 5% to induce death. Leave the mice in the chamber for an additional 2 min after breathing has ceased. Death is verified by observing that the mice fail to recover within 10 min after overdose of isoflurane.</li>
		<li>Finally, perform cervical dislocation on the mice. 
		NOTE: In the case of bilaterally xenografted mice, the contralateral xenograft can be saved for a later collection. To perform a survival collection, open the skin overlying the xenograft with a single straight cut with surgical scissors, and remove the xenograft as described in steps 4.6 to 4.10. Then close the skin over the empty tibial compartment using surgical glue and staples. Treat the mouse with analgesic as described in step 2.11 and place the mouse in clean cage on heated pad to recover. Monitor the mouse until fully conscious and periodically over the next few days for signs of local systemic infection and to ensure the surgical site is not reopened.</li>
	</ol>
	</li>
</ol>
5. Xenograft Immunohistochemistry
<ol>
	<li>Use a cryostat to cut 10 to 12 μm sections from the collected xenograft onto positively charged slides (Table of Materials).</li>
	<li>Fill staining jar with methanol and pre-cool at -20 °C for 30 min.</li>
	<li>Place slides in ice cold methanol for 10 min to fix and permeabilize the xenograft sections.</li>
	<li>Place slides in staining jar and wash 3x with phosphate buffered saline (PBS) for 5 min.</li>
	<li>Block with anti-mouse IgG (Table of Materials) for 2 h at 4 °C.</li>
	<li>Blot with primary antibodies, such as spectrin, lamin A/C, and embryonic myosin (Table of Materials) in PBS supplemented with 2% goat serum overnight at 4 °C.</li>
	<li>Place slides in a staining jar and wash 3x with phosphate buffered saline (PBS) for 5 min.</li>
	<li>Blot with fluorescent-dye conjugated secondary antibodies (Table of Materials) in PBS supplemented with 2% goat serum for 1 h at room temperature.</li>
	<li>Place slides in staining jar and wash 3x with phosphate buffered saline (PBS) for 5 min.</li>
	<li>Place mounting medium (Table of Materials) over xenografts sections, place coverslip on top, and use nail polish to seal the coverslip.</li>
</ol>
<img alt="Muscle dissection and labeling diagram; EDL and TA muscle identification in rodent limb." class="xfigimg" src="/files/ftp_upload/59966/59966fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Xenograft Surgery. (A) Hair is removed from surgical site. (B) An incision is made over the tibialis anterior (TA). The distal tendons of the TA and extensor digitorum longus (EDL) are marked with arrows. The black dashed line indicates where the epimysium will be cut in step 3.3. (C) The distal tendon of the TA is cut and the muscle is pulled up to the knee. (D) The tendon of the EDL is cut and the EDL is pulled up to the knee. This exposes the proximal tendon of the peroneus longus (PL) marked with an arrow. Dashed lines indicate where to cut with scissors to remove the EDL (green) and PL (blue). (E) The EDL and TA are removed. (F) A suture is placed through the proximal tendon of the PL. (G) The xenograft is placed in the empty tibial compartment and sutured to the proximal PL tendon using a two-hand surgical square knot. (H) A suture is placed through the distal tendon of the PL, marked with an arrow, and another two-hand surgical square knot is used to suture the xenograft to the distal tendon. (I) The xenograft is fully transplanted and sutured to the PL. (J) The skin is closed with surgical glue. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/59966/59966fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Tendon surgical procedure, sequential steps, using forceps, incision, muscle exposure, research method." class="xfigimg" src="/files/ftp_upload/59966/59966fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: 4 Month Xenograft Collection. (A) Hair is removed from surgical site. Sutures are visible under skin. (B) The skin overlying the xenograft is removed. Then the xenograft is grabbed with the iris forceps at the distal suture and gently pulled upward. Starting at the ankle, a scalpel is used to cut along the tibia and free the xenograft. The arrow shows the beginning of the incision along the tibia. (C) By pulling the gastrocnemius muscle to the side, a faint white line of epimysium separating the peroneus longus (PL) muscle and the gastrocnemius (shown by the arrow) becomes visible. Use the scalpel to cut along this line to separate the PL from the other leg muscles. (D) The right side of the xenograft, and the PL are now free from the other muscles in the leg and are ready for removal. The dashed line indicates where to cut with surgical scissors to start removing the xenograft and PL. (E) After cutting below the distal suture, deflect the xenograft toward the knee. The dashed line indicates where to cut with surgical scissors to remove the xenograft and PL from the tibial compartment. (F) The empty tibial compartment with the xenograft and PL successfully removed. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/59966/59966fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors declare that they have no competing financial interests.

Patient-derived xenografts are an innovative way to model muscle disease and carry out preclinical studies. The method described here to create skeletal muscle xenografts is rapid, straightforward, and reproducible. Unilateral surgeries can be performed in 15 to 25 minutes, or bilaterally in 30 to 40 minutes. Bilateral xenografts can provide additional experimental flexibility. For instance, researchers can perform localized treatment of one xenograft, with the other left as a control. The NRG mice are resistant to surgi...

It has been reported that only 13.8% of all drug development programs undergoing clinical trials are successful and lead to approved therapies1. While this success rate is higher than the 10.4% previously reported2, there is still significant room for improvement. One approach to increase the success rate of clinical trials is to improve laboratory models used in preclinical research. The Food and Drug Administration (FDA) requires animal studies to show treatment efficacy and assess toxicity prior to Phase 1 clinical trials. However, there is often limited concordance in treatment outcomes between animal studies and clinical trials3. In addition, the need for preclinical animal studies can be an insurmountable barrier for therapeutic development in diseases that lack an accepted animal model, which is often the case for rare or sporadic diseases.
One way to model human disease is by transplanting human tissue into immunodeficient mice to generate xenografts. There are three key advantages to xenograft models: First, they can recapitulate the complex genetic and epigenetic abnormalities that exist in human disease that may never be reproducible in other animal models. Second, xenografts can be used to model rare or sporadic diseases if patient samples are available. Third, xenografts model the disease within a complete in vivo system. For these reasons, we hypothesize that treatment efficacy results in xenograft models are more likely to translate to trials in patients. Human tumor xenografts have already been successfully utilized to develop treatments for common cancers, including multiple myeloma, as well as personalized therapies for individual patients4,5,6,7.
Recently, xenografts have been used to develop a model of human muscle disease8. In this model, human muscle biopsy specimens are transplanted into the hindlimbs of immunodeficient NRG mice to form xenografts. The transplanted human myofibers die, but human muscle stem cells present in the xenograft subsequently expand and differentiate into new human myofibers which repopulate the engrafted human basal lamina. Therefore, the regenerated myofibers in these xenografts are entirely human and are spontaneously revascularized and innervated by the mouse host. Importantly, fascioscapulohumeral muscular dystrophy (FSHD) patient muscle tissue transplanted into mice recapitulates key features of the human disease, namely expression of the DUX4 transcription factor8. FSHD is caused by overexpression of DUX4, which is epigenetically silenced in normal muscle tissue9,10. In the FSHD xenograft model, treatment with a DUX4-specific morpholino has been shown to successfully repress DUX4 expression and function, and may be a potential therapeutic option for FSHD patients11. These results demonstrate that human muscle xenografts are a new approach to model human muscle disease and test potential therapies in mice. Here, we describe in detail the surgical method for creating human skeletal muscle xenografts in immunodeficient mice.

<table><tbody><tr><td>100 mm x 15 mm Petri dish</td><td>Fisher Scientific</td><td>FB0875712</td><td></td></tr><tr><td>2-Methylbutane</td><td>Fisher</td><td>O3551-4</td><td></td></tr><tr><td>20 mm x 30 mm micro cover glass</td><td>VWR</td><td>48393-151</td><td></td></tr><tr><td>Animal Weighing Scale</td><td>Kent Scientific</td><td>SCL- 1015</td><td></td></tr><tr><td>Antibiotic-Antimycotic Solution</td><td>Corning, Cellgro</td><td>30-004-CI</td><td></td></tr><tr><td>AutoClip System</td><td>F.S.T</td><td>12020-00</td><td></td></tr><tr><td>Castroviejo Needle Holder</td><td>F.S.T</td><td>12565-14</td><td></td></tr><tr><td>Chick embryo extract</td><td>Accurate</td><td>CE650TL</td><td></td></tr><tr><td>CM1860 UV cryostat</td><td>Leica Biosystems</td><td>CM1860UV</td><td></td></tr><tr><td>Coplin staining jar</td><td>Thermo Scientific</td><td>19-4</td><td></td></tr><tr><td>Dissection Pins</td><td>Fisher Scientific</td><td>S13976</td><td></td></tr><tr><td>Dry Ice - pellet</td><td>Fisher Scientific</td><td>NC9584462</td><td></td></tr><tr><td>Embryonic Myosin antibody</td><td>DSHB</td><td>F1.652</td><td>recommended concentration 1:10</td></tr><tr><td>Ethanol</td><td>Fisher Scientific</td><td>459836</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>GE Healthcare Life Sciences</td><td>SH30071.01</td><td></td></tr><tr><td>Fiber-Lite MI-150</td><td>Dolan-Jenner</td><td>Mi-150</td><td></td></tr><tr><td>Forceps</td><td>F.S.T</td><td>11295-20</td><td></td></tr><tr><td>Goat anti-mouse IgG1, Alexa Fluor 488</td><td>Invitrogen</td><td>A-21121</td><td>recommended concentration 1:500</td></tr><tr><td>Goat anti-mouse IgG2b, AlexaFluor 594</td><td>Invitrogen</td><td>A-21145</td><td>recommended concentration 1:500</td></tr><tr><td>Gum tragacanth</td><td>Sigma</td><td>G1128</td><td></td></tr><tr><td>Hams F-10 Medium</td><td>Corning</td><td>10-070-CV</td><td></td></tr><tr><td>Histoacryl Blue Topical Skin Adhesive</td><td>Tissue seal</td><td>TS1050044FP</td><td></td></tr><tr><td>Human specific lamin A/C antibody</td><td>Abcam</td><td>ab40567</td><td>recommended concentration 1:50-1:100</td></tr><tr><td>Human specific spectrin antibody</td><td>Leica Biosystems</td><td>NCLSPEC1</td><td>recommended concentration 1:20-1:100</td></tr><tr><td>Induction Chamber</td><td>VetEquip</td><td>941444</td><td></td></tr><tr><td>Iris Forceps</td><td>F.S.T</td><td>11066-07</td><td></td></tr><tr><td>Irradiated Global 2018 (Uniprim 4100 ppm)</td><td>Envigo</td><td>TD.06596</td><td>Antibiotic rodent diet to protect again respiratory infections</td></tr><tr><td>Isoflurane</td><td>MWI Veterinary Supply</td><td>502017</td><td></td></tr><tr><td>Kimwipes</td><td>Kimberly-Clark</td><td>34155</td><td>surgical wipes</td></tr><tr><td>Mapleson E Breathing Circuit</td><td>VetEquip</td><td>921412</td><td></td></tr><tr><td>Methanol</td><td>Fisher Scientific</td><td>A412</td><td></td></tr><tr><td>Mobile Anesthesia Machine</td><td>VetEquip</td><td>901805</td><td></td></tr><tr><td>Mouse on Mouse Basic Kit</td><td>Vector Laboratories</td><td>BMK-2202</td><td>mouse IgG blocking reagent</td></tr><tr><td>Nail Polish</td><td>Electron Microscopy Sciences</td><td>72180</td><td></td></tr><tr><td>NAIR Hair remover lotion/oil</td><td>Fisher Scientific</td><td>NC0132811</td><td></td></tr><tr><td>NOD-Rag1&lt;sup&gt;null&lt;/sup&gt; IL2rg &lt;sup&gt;null&lt;/sup&gt; (NRG) mice</td><td>The Jackson Laboratory</td><td>007799</td><td>2 to 3 months old</td></tr><tr><td>O.C.T. Compound</td><td>Fisher Scientific</td><td>23-730-571</td><td></td></tr><tr><td>Oxygen</td><td>Airgas</td><td>OX USPEA</td><td></td></tr><tr><td>PBS (phosphate buffered saline) buffer</td><td>Fisher Scientific</td><td>4870500</td><td></td></tr><tr><td>Povidone Iodine Prep Solution</td><td>Dynarex</td><td>1415</td><td></td></tr><tr><td>ProLong&amp;trade; Gold Antifade Mountant</td><td>Fisher Scientific</td><td>P10144 (no DAPI); P36935 (with DAPI)</td><td></td></tr><tr><td>Puralube Ophthalmic Ointment</td><td>Dechra</td><td>17033-211-38</td><td></td></tr><tr><td>Rimadyl (carprofen) injectable</td><td>Patterson Veterinary</td><td>10000319</td><td>surgical analgesic, administered subcutaneously at a dose of 5 mg/kg</td></tr><tr><td>Scalpel Blades - #11</td><td>F.S.T</td><td>10011-00</td><td></td></tr><tr><td>Scalpel Handle - #3</td><td>F.S.T</td><td>10003-12</td><td></td></tr><tr><td>Stereo Microscope</td><td>Accu-scope</td><td>3075</td><td></td></tr><tr><td>Superfrost Plus Microscope Slides</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr><tr><td>Suture, Synthetic, Non-Absorbable, 30 inches long, CV-11 needle</td><td>Covidien</td><td>VP-706-X</td><td></td></tr><tr><td>1ml Syringe (26 gauge, 3/8 inch needle)</td><td>BD Biosciences</td><td>329412</td><td></td></tr><tr><td>Trimmer</td><td>Kent Scientific</td><td>CL9990-KIT</td><td></td></tr><tr><td>Vannas Spring Scissors, 8.0 mm cutting edge</td><td>F.S.T</td><td>15009-08</td><td></td></tr><tr><td>VaporGaurd Activated Charcoal Filter</td><td>VetEquip</td><td>931401</td><td></td></tr><tr><td>Wound clips, 9 mm</td><td>F.S.T</td><td>12022-09</td><td></td></tr></tbody></table>

performing human skeletal muscle xenografts in immunodeficient mice

Treatment effects observed in animal studies often fail to be recapitulated in clinical trials. While this problem is multifaceted, one reason for this failure is the use of inadequate laboratory models. It is challenging to model complex human diseases in traditional laboratory organisms, but this issue can be circumvented through the study of human xenografts. The surgical method we describe here allows for the creation of human skeletal muscle xenografts, which can be used to model muscle disease and to carry out preclinical therapeutic testing. Under an Institutional Review Board (IRB)-approved protocol, skeletal muscle specimens are acquired from patients and then transplanted into NOD-Rag1null IL2r&gamma;null (NRG) host mice. These mice are ideal hosts for transplantation studies due to their inability to make mature lymphocytes and are thus unable to develop cell-mediated and humoral adaptive immune responses. Host mice are anaesthetized with isoflurane, and the mouse tibialis anterior and extensor digitorum longus muscles are removed. A piece of human muscle is then placed in the empty tibial compartment and sutured to the proximal and distal tendons of the peroneus longus muscle. The xenografted muscle is spontaneously vascularized and innervated by the mouse host, resulting in robustly regenerated human muscle that can serve as a model for preclinical studies.

As demonstrated by Yuanfan Zhang et al., this surgical protocol is a straightforward method to produce human skeletal muscle xenografts8. Regenerated xenografts become spontaneously innervated and display functional contractility. In addition, muscle xenografted from FSHD patients recapitulates changes in gene expression observed in FSHD patients8.
In our experience, approximately 7 out of 8 xenografts performed from control patient specimens wil...

Watch this Scientific Journal Video about Performing Human Skeletal Muscle Xenografts in Immunodeficient Mice at JoVE.com

Performing Human Skeletal Muscle Xenografts in Immunodeficient Mice

Complex human diseases can be challenging to model in traditional laboratory model systems. Here, we describe a surgical approach to model human muscle disease through the transplantation of human skeletal muscle biopsies into immunodeficient mice.

Treatment effects observed in animal studies often fail to be recapitulated in clinical trials. While this problem is multifaceted, one reason for this ...

performing-human-skeletal-muscle-xenografts-in-immunodeficient-mice

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

9.8K Views.  Johns Hopkins University School of Medicine. Complex human diseases can be challenging to model in traditional laboratory model systems. Here, we describe a surgical approach to model human muscle disease through the transplantation of human skeletal muscle biopsies into immunodeficient mice.

Video: Performing Human Skeletal Muscle Xenografts in Immunodeficient Mice

Orthotopic Xenografting of Human Luciferase-Tagged Malignant Peripheral Nerve Sheath Tumor Cells for in vivo Testing of Candidate Therapeutic Agents

Although in vitro screens are essential for the initial identification of candidate therapeutic agents, a rigorous assessment of the drug's ability to inhibit tumor growth must be performed in a suitable animal model. The type of animal model that is best for this purpose is a topic of intense discussion. Some evidence indicates that preclinical trials examining drug effects on tumors arising in transgenic mice are more predictive of clinical outcome1and so candidate therapeutic agents are often tested in these models. Unfortunately, transgenic models are not available for many tumor types. Further, transgenic models often have other limitations such as concerns as to how well the mouse tumor models its human counterpart, incomplete penetrance of the tumor phenotype and an inability to predict when tumors will develop.
Consequently, many investigators use xenograft models (human tumor cells grafted into immunodeficient mice) for preclinical trials if appropriate transgenic tumor models are not available. Even if transgenic models are available, they are often partnered with xenograft models as the latter facilitate rapid determination of therapeutic ranges. Further, this partnership allows a comparison of the effectiveness of the agent in transgenic tumors and genuine human tumor cells. Historically, xenografting has often been performed by injecting tumor cells subcutaneously (ectopic xenografts). This technique is rapid and reproducible, relatively inexpensive and allows continuous quantitation of tumor growth during the therapeutic period2. However, the subcutaneous space is not the normal microenvironment for most neoplasms and so results obtained with ectopic xenografting can be misleading due to factors such as an absence of organ-specific expression of host tissue and tumor genes. It has thus been strongly recommended that ectopic grafting studies be replaced or complemented by studies in which human tumor cells are grafted into their tissue of origin (orthotopic xenografting)2. Unfortunately, implementation of this recommendation is often thwarted by the fact that orthotopic xenografting methodologies have not yet been developed for many tumor types.
Malignant peripheral nerve sheath tumors (MPNSTs) are highly aggressive sarcomas that occur sporadically or in association with neurofibromatosis type 13and most commonly arise in the sciatic nerve4. Here we describe a technically straightforward method in which firefly luciferase-tagged human MPNST cells are orthopically xenografted into the sciatic nerve of immunodeficient mice. Our approach to assessing the success of the grafting procedure in individual animals and subsequent non-biased randomization into study groups is also discussed.

Orthotopic Xenografting of Human Luciferase-Tagged Malignant Peripheral Nerve Sheath Tumor Cells for in vivo Testing of Candidate Therapeutic Agents

A method for reliably grafting luciferase-tagged human malignant peripheral nerve sheath tumor cells into the sciatic nerve of immunodeficient mice is described. The use of bioluminescence imaging to demonstrate proper establishment of tumor grafts and criteria for random segregation of animals into study groups are also discussed.

Although in vitro screens are essential for the initial identification of candidate therapeutic agents, a rigorous assessment of the drug's ability to ...

Medicine

Transplantation of Induced Pluripotent Stem Cell-derived Mesoangioblast-like Myogenic Progenitors in Mouse Models of Muscle Regeneration

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells similar to pericyte-derived mesoangioblasts (iPSC-derived mesoangioblast-like stem/progenitor cells: IDEMs) can be established from iPSCs generated from patients affected by different forms of muscular dystrophy. Patient-specific IDEMs can be genetically corrected with different strategies (e.g. lentiviral vectors, human artificial chromosomes) and enhanced in their myogenic differentiation potential upon overexpression of the myogenesis regulator MyoD. This myogenic potential is then assessed in vitro with specific differentiation assays and analyzed by immunofluorescence. The regenerative potential of IDEMs is further evaluated in vivo, upon intramuscular and intra-arterial transplantation in two representative mouse models displaying acute and chronic muscle regeneration. The contribution of IDEMs to the host skeletal muscle is then confirmed by different functional tests in transplanted mice. In particular, the amelioration of the motor capacity of the animals is studied with treadmill tests. Cell engraftment and differentiation are then assessed by a number of histological and immunofluorescence assays on transplanted muscles. Overall, this paper describes the assays and tools currently utilized to evaluate the differentiation capacity of IDEMs, focusing on the transplantation methods and subsequent outcome measures to analyze the efficacy of cell transplantation.

Induced pluripotent stem cell (iPSC)-derived myogenic progenitors are promising candidates for cell therapy strategies to treat muscular dystrophies. This protocol describes transplantation and functional measurements required to evaluate the engraftment and differentiation of iPSC-derived mesoangioblasts (a type of muscle progenitors) in mouse models of acute and chronic muscle regeneration.

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells ...

Bioengineering

Generation of Subcutaneous and Intrahepatic Human Hepatocellular Carcinoma Xenografts in Immunodeficient Mice

In vivo experimental models of hepatocellular carcinoma (HCC) that recapitulate the human disease provide a valuable platform for research into disease pathophysiology and for the preclinical evaluation of novel therapies. We present a variety of methods to generate subcutaneous or orthotopic human HCC xenografts in immunodeficient mice that could be utilized in a variety of research applications. With a focus on the use of primary tumor tissue from patients undergoing surgical resection as a starting point, we describe the preparation of cell suspensions or tumor fragments for xenografting. We describe specific techniques to xenograft these tissues i) subcutaneously; or ii) intrahepatically, either by direct implantation of tumor cells or fragments into the liver, or indirectly by injection of cells into the mouse spleen. We also describe the use of partial resection of the native mouse liver at the time of xenografting as a strategy to induce a state of active liver regeneration in the recipient mouse that may facilitate the intrahepatic engraftment of primary human tumor cells. The expected results of these techniques are illustrated. The protocols described have been validated using primary human HCC samples and xenografts, which typically perform less robustly than the well-established human HCC cell lines that are widely used and frequently cited in the literature. In comparison with cell lines, we discuss factors which may contribute to the relatively low chance of primary HCC engraftment in xenotransplantation models and comment on technical issues that may influence the kinetics of xenograft growth. We also suggest methods that should be applied to ensure that xenografts obtained accurately resemble parent HCC tissues.

Human tumor xenografts in immunodeficient mice are valuable tools to study cancer biology. Specific protocols to generate subcutaneous and intrahepatic xenografts from human hepatocellular carcinoma cells or tumor fragments are described. Liver regeneration induced by partial hepatectomy in recipient mice is presented as a strategy to facilitate intrahepatic engraftment.

In  vivo experimental models of hepatocellular carcinoma (HCC) that recapitulate  the human disease provide a valuable platform for research into disease  ...

Tracking Dynamics of Muscle Engraftment in Small Animals by In Vivo Fluorescent Imaging

Muscular dystrophies are a group of degenerative muscle diseases characterized by progressive loss of contractile muscle cells. Currently, there is no curative treatment available. Recent advances in stem cell biology have generated new hopes for the development of effective cell based therapies to treat these diseases. Transplantation of various types of stem cells labeled with fluorescent proteins into muscles of dystrophic animal models has been used broadly in the field. A non-invasive technique with the capability to track the transplanted cell fate longitudinally can further our ability to evaluate muscle engraftment by transplanted cells more accurately and efficiently.
In the present study, we demonstrate that in vivo fluorescence imaging is a sensitive and reliable method for tracking transplanted GFP (Green Fluorescent Protein)-labeled cells in mouse skeletal muscles. Despite the concern about background due to the use of an external light necessary for excitation of fluorescent protein, we found that by using either nude mouse or eliminating hair with hair removal reagents much of this problem is eliminated. Using a CCD camera, the fluorescent signal can be detected in the tibialis anterior (TA) muscle after injection of 5 x 105 cells from either GFP transgenic mice or eGFP transduced myoblast culture. For more superficial muscles such as the extensor digitorum longus (EDL), injection of fewer cells produces a detectable signal. Signal intensity can be measured and quantified as the number of emitted photons per second in a region of interest (ROI). Since the acquired images show clear boundaries demarcating the engrafted area, the size of the ROI can be measured as well. If the legs are positioned consistently every time, the changes in total number of photons per second per muscle and the size of the ROI reflect the changes in the number of engrafted cells and the size of the engrafted area. Therefore the changes in the same muscle over time are quantifiable. In vivo fluorescent imaging technique has been used primarily to track the growth of tumorogenic cells, our study shows that it is a powerful tool that enables us to track the fate of transplanted stem cells.

Tracking Dynamics of Muscle Engraftment in Small Animals by In Vivo Fluorescent Imaging

We describe an in vivo fluorescence imaging protocol to monitor muscle regeneration by GFP-labeled myoblasts after transplantation into skeletal muscles of both healthy and dystrophic mice. This protocol can be adapted to study muscle regeneration by transplantation of other types of cells and in other muscular conditions as well.

Muscular dystrophies are a group of degenerative muscle diseases characterized by progressive loss of contractile muscle cells. Currently, there is no ...

Biology

Isolation, Culture, Characterization, and Differentiation of Human Muscle Progenitor Cells from the Skeletal Muscle Biopsy Procedure

The use of primary human tissue and cells is ideal for the investigation of biological and physiological processes such as the skeletal muscle regenerative process. There are recognized challenges to working with human primary adult stem cells, particularly human muscle progenitor cells (hMPCs) derived from skeletal muscle biopsies, including low cell yield from collected tissue and a large degree of donor heterogeneity of growth and death parameters among cultures. While incorporating heterogeneity into experimental design requires a larger sample size to detect significant effects, it also allows us to identify mechanisms that underlie variability in hMPC&#160;expansion capacity, and thus allows us to better understand heterogeneity in skeletal muscle regeneration. Novel mechanisms that distinguish the expansion capacity of cultures have the potential to lead to the development of therapies to improve skeletal muscle regeneration.

We present techniques for isolating, culturing, characterizing, and differentiating human primary muscle progenitor cells (hMPCs) obtained from skeletal muscle biopsy tissue. hMPCs obtained and characterized through these methods can be used to subsequently address research questions related to human myogenesis and skeletal muscle regeneration.

The use of primary human tissue and cells is ideal for the investigation of biological and physiological processes such as the skeletal muscle ...

Bioengineering Human Microvascular Networks in Immunodeficient Mice

The future of tissue engineering and cell-based therapies for tissue regeneration will likely rely on our ability to generate functional vascular networks in vivo. In this regard, the search for experimental models to build blood vessel networks in vivo is of utmost importance 1. The feasibility of bioengineering microvascular networks in vivo was first shown using human tissue-derived mature endothelial cells (ECs) 2-4; however, such autologous endothelial cells present problems for wide clinical use, because they are difficult to obtain in sufficient quantities and require harvesting from existing vasculature. These limitations have instigated the search for other sources of ECs. The identification of endothelial colony-forming cells (ECFCs) in blood presented an opportunity to non-invasively obtain ECs 5-7. We and other authors have shown that adult and cord blood-derived ECFCs have the capacity to form functional vascular networks in vivo 7-11. Importantly, these studies have also shown that to obtain stable and durable vascular networks, ECFCs require co-implantation with perivascular cells. The assay we describe here illustrates this concept: we show how human cord blood-derived ECFCs can be combined with bone marrow-derived mesenchymal stem cells (MSCs) as a single cell suspension in a collagen/fibronectin/fibrinogen gel to form a functional human vascular network within 7 days after implantation into an immunodeficient mouse. The presence of human ECFC-lined lumens containing host erythrocytes can be seen throughout the implants indicating not only the formation (de novo) of a vascular network, but also the development of functional anastomoses with the host circulatory system. This murine model of bioengineered human vascular network is ideally suited for studies on the cellular and molecular mechanisms of human vascular network formation and for the development of strategies to vascularize engineered tissues.

Here, we describe a methodology to deliver human cord blood-derived endothelial colony-forming cells (ECFCs) and bone marrow-derived mesenchymal stem cells (MSCs), embedded in a collagen/fibronectin gel, subcutaneously into immunodeficient mice. This cell/gel combination generates a human vascular network that connects with the mouse vasculature.

The future of tissue engineering and cell-based therapies for tissue regeneration will likely rely on our ability to generate functional vascular networks ...

Minimally Invasive Muscle Embedding (MIME) - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

Skeletal muscle possesses regenerative capacity due to tissue-resident, muscle-fiber-generating (myogenic) satellite cells (SCs), which can form new muscle fibers under the right conditions. Although SCs can be harvested from muscle tissue and cultured in vitro, the resulting myoblast cells are not very effective in promoting myogenesis when transplanted into host muscle. Surgically exposing the host muscle and grafting segments of donor muscle tissue, or the isolated muscle fibers with their SCs onto host muscle, promotes better myogenesis compared to myoblast transplantation. We have developed a novel technique that we call Minimally Invasive Muscle Embedding (MIME). MIME involves passing a surgical needle through the host muscle, drawing a piece of donor muscle tissue through the needle track, and then leaving the donor tissue embedded in the host muscle so that it may act as a source of SCs for the host muscle. Here we describe in detail the steps involved in performing MIME in an immunodeficient mouse model that expresses a green fluorescent protein (GFP) in all of its cells. Immunodeficiency in the host mouse reduces the risk of immune rejection of the donor tissue, and GFP expression enables easy identification of the host muscle fibers (GFP+) and donor-cell-derived muscle fibers (GFP-). Our pilot data suggest that MIME can be used to implant an extensor digitorum longus (EDL) muscle from a donor mouse into the tibialis anterior (TA) muscle of a host mouse. Our data also suggest that when a myotoxin (barium chloride, BaCl2) is injected into the host muscle after MIME, there is evidence of donor-cell-derived myogenesis in the host muscle, with approximately 5%, 26%, 26% and 43% of the fibers in a single host TA muscle showing no host contribution, minimal host contribution, moderate host contribution, and maximal host contribution, respectively.

Minimally Invasive Muscle Embedding MIME - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

We describe a novel experimental technique that we call Minimally Invasive Muscle Embedding (MIME), which is based on the evidence that skeletal muscle tissue contains viable myogenic cells that can facilitate donor-cell-mediated myogenesis when implanted into a host muscle.

Skeletal muscle possesses regenerative capacity due to tissue-resident, muscle-fiber-generating (myogenic) satellite cells (SCs), which can form new ...

Xenograft Skin Model to Manipulate Human Immune Responses In Vivo

The human skin xenograft model, in which human donor skin is transplanted onto an immunodeficient mouse host, is an important option for translational research in skin immunology. Murine and human skin differ substantially in anatomy and immune cell composition. Therefore, traditional mouse models have limitations for dermatological research and drug discovery. However, successful xenotransplants are technically challenging and require optimal specimen and mouse graft site preparation for graft and host survival. The present protocol provides an optimized technique for transplanting human skin onto mice and discusses necessary considerations for downstream experimental aims. This report describes the appropriate preparation of a human donor skin sample, assembly of a surgical setup, mouse and surgical site preparation, skin transplantation, and post-surgical monitoring. Adherence to these methods allows for maintenance of xenografts for over 6 weeks post-surgery. The techniques outlined below allow maximum grafting efficiency due to the development of engineering controls, sterile technique, and pre- and post-surgical conditioning. Appropriate performance of the xenograft model results in long-lived human skin graft samples for experimental characterization of human skin and preclinical testing of compounds in vivo.

Xenograft Skin Model to Manipulate Human Immune Responses In Vivo

The present protocol describes how to graft human skin onto non-obese diabetic (NOD)-scid interleukin-2 gamma chain receptor (NSG) mice. A detailed description of the preparation of human skin for transplant, preparation of mice for transplant, transplantation of split-thickness human skin, and post-transplantation recovery procedure are included in the report.

The human skin xenograft model, in which human donor skin is transplanted onto an immunodeficient mouse host, is an important option for translational ...

Immunology and Infection

Injecting Human HSPCs into the Femur of Sub-Lethally Irradiated Mice

Intrafemoral Injection of Human Hematopoietic Stem and Progenitor Cells into Immunocompromised Mice

Hematopoietic stem cells (HSCs) are defined by their lifelong ability to produce all blood cell types. This is operationally tested by transplanting cell populations containing HSCs into syngeneic or immunocompromised mice. The size and multilineage composition of the graft is then measured over time, usually by flow cytometry. Classically, a population containing HSCs is injected into the circulation of the animal, after which the HSCs home to the bone marrow, where they lodge and begin blood production. Alternatively, HSCs and/or progenitor cells (HSPCs) can be placed directly in the bone marrow cavity. 
This paper describes a protocol for intrafemoral injection of human HSPCs into immunodeficient mice. In short, preconditioned mice are anesthetized, and a small hole is drilled through the knee into the femur using a needle. Using a smaller insulin needle, cells are then injected directly into the same conduit created by the first needle. This method of transplantation can be applied in varied experimental designs, using either mouse or human cells as donor cells. It has been most widely used for xenotransplantation, because in this context, it is thought to provide improved engraftment over intravenous injections, therefore improving statistical power and reducing the number of mice to be used.

Author Spotlight: Exploring the Lifespan Dynamics of Healthy Human Hematopoiesis

Intrafemoral injections allow for the engraftment of a small number of hematopoietic stem and progenitor cells (HSPCs), by placing the cells directly in the bone marrow cavity. Here we describe an experimental protocol of intrafemoral injection of human HSPCs into immunodeficient mice.

Hematopoietic stem cells (HSCs) are defined by their lifelong ability to produce all blood cell types. This is operationally tested by transplanting cell ...

A Technique for Human Skin Grafting in a Mouse Model

Source: Moss, M. I. et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/64040">Xenograft Skin Model to Manipulate Human Immune Responses&#160;In Vivo.</a>&#160;J. Vis. Exp. (2022)
This video demonstrates a method for grafting human skin onto an immunodeficient mouse with a suppressed neutrophil response. The method involves applying human split-thickness skin to the mouse&#39;s graft bed, sealing it, and covering it for recovery. Eventually, the human skin adheres to the mouse&#39;s skin, establishing vascularization and circulation, resulting in a successful skin xenograft.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Performing Human Skeletal Muscle Xenografts in Immunodeficient Mice

Summary

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Generation of Subcutaneous and Intrahepatic Human Hepatocellular Carcinoma Xenografts in Immunodeficient Mice

Tracking Dynamics of Muscle Engraftment in Small Animals by In Vivo Fluorescent Imaging

Isolation, Culture, Characterization, and Differentiation of Human Muscle Progenitor Cells from the Skeletal Muscle Biopsy Procedure

Bioengineering Human Microvascular Networks in Immunodeficient Mice

Minimally Invasive Muscle Embedding (MIME) - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

Xenograft Skin Model to Manipulate Human Immune Responses In Vivo

Intrafemoral Injection of Human Hematopoietic Stem and Progenitor Cells into Immunocompromised Mice

A Technique for Human Skin Grafting in a Mouse Model