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 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 IL2rγnull (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.
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.
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
2. Surgical Preparation
3. Xenograft Surgery
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.
5. Xenograft Immunohistochemistry
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. Please click here to view a larger version of this figure.
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. Please click here to view a larger version of this figure.
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 will show successful muscle engraftment. A successful xenograft shows robust regeneration of human myofibers as identified with human specific antibodies (Figure 4). Positive embryonic myosin staining within a proportion of myofibers indicates that the regeneration process is still ongoing. In contrast, poor surgical technique or an inadequate specimen may lead to poor regeneration of muscle fibers (Figure 4).
Xenografts performed from a patient diagnosed with an idiopathic inflammatory myopathy (IIM) show moderate numbers of regenerated human myofibers at 4- and 6-month collections, and embryonic myosin staining persists at 6 months (Figure 5A). Inflammatory cells are present in the xenograft as shown by H&E staining (Figure 5A), and have been confirmed with CD3, CD68, and other immunological markers (data not shown). Xenografts are stable within the mouse, and up to 12-month collections have been performed. Individual myofiber size is comparable between the 4- and 6-month IIM xenografts and the original IIM patient biopsy (Figure 5B). Rare fibers showing a cross sectional area (CSA) greater than 3500 μm2 are observed in xenografts but not in the IIM biopsy, indicating that some myofibers in the xenografts can regenerate to a CSA comparable in size to healthy myofibers (Figure 5B).
Figure 1: Surgical Set-up.
A) Standard orientation of stereo microscope, Mapleson E breathing circuit, and surgical tools during xenograft surgery. B) Placement of induction chamber in biosafety cabinet.
Figure 4: Expected Positive and Negative Results.
Xenografts collected 4-months post-surgery showing good or poor regeneration are stained with human-specific lamin A/C (1:50) and human-specific spectrin (1:20) and embryonic myosin (1:10) (Table of Materials). Regions indicated by the white dashed boxes are shown as higher magnification inserts. Scale bar: 200 µm. Please click here to view a larger version of this figure.
Figure 5: Representative Xenograft regeneration.
A) Xenografts (outlined with dashed lines) performed from a patient diagnosed with an idiopathic inflammatory myopathy (IIM) stained with Hematoxylin and Eosin (H&E), human specific Lamin A/C, and human specific spectrin, show myofiber formation within NRG mice at both 4- and 6-month time points. Embryonic myosin staining demonstrates that regeneration is still ongoing at both time points. Scale bar: 200 µm. B) Histograms depicting cross sectional area (CSA) of myofibers from 4- and 6-month xenografts and human biopsies from one patient diagnosed with an idiopathic inflammatory myopathy (IIM) and one healthy control patient. Please click here to view a larger version of this figure.
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 surgical site infection when housed in a pathogen-free facility; in our experience performing more than 200 xenografts, we have never had a mouse acquire a surgical infection. In addition, host mice tolerate the removal of the TA and EDL very well. Within an hour post-surgery, unilaterally and bilaterally xenografted mice will be active and walking around their cage, and even standing up on their hindlimbs. Occasionally we observe some foot drop in host mice, but usually only after a period of inactivity, such as if recently awoken, and within minutes of waking leg use will be normal.
There are several critical steps in the protocol. First, during removal of the EDL and TA, it is very important to not injure the adjacent PL muscle or its tendons. This can be avoided by carefully and correctly identifying the placement of all distal tendons after the initial incision over the TA is performed. In addition, the proximal tendon of the PL should be identified and clearly visible before removal of the EDL (Figure 2D). Second, sutures must be placed through tendons and tightened fully in a proper two-hand surgical square knot. Xenografts regenerate more robustly under tension, and this is only obtainable if the xenograft is tethered to the PL tendons and if the sutures do not loosen post-operatively. Finally, it is important to not damage or sever any major blood vessels supplying the foot. In particular, 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.
This method does have several limitations. It is not amenable to standard functional assays used in mouse models of muscle disease, such as grip strength or treadmill endurance. However, electrophysiological assessments of xenograft function can still be performed. Evoked force measurements can be recorded from xenograft explants, and single enzymatically isolated myofibers from xenografts loaded with ratiometric calcium dyes and electrically stimulated can be used to study calcium dynamics8. Another inherent challenge in this model is that acquiring and working with human tissue can be difficult. Not all laboratories will have easy access to fresh muscle biopsies, but it has been shown that xenografts performed from autopsy tissue approximately 48 hours post mortem can successfully engraft, and this tissue may be easier to obtain for some laboratories8. It is also challenging to manipulate gene expression in human tissue, whereas researchers using standard mouse models of disease can readily use the plethora of mouse genetic tools available.
A strength of this xenograft model is that it allows researchers to study human muscle in vivo. Tissue culture has been used extensively to study the cell and molecular biology of human muscle. Yet, these short-term, ex vivo studies do not always approximate functional muscle in vivo. However, one caveat is that is it challenging to determine how closely xenograft biology and function approximates human muscle due to the contribution of host mouse components during the regenerative process. For instance, human and mouse neuromuscular junctions (NMJs) are morphologically distinct, and there is significant divergence between the synaptic proteome of human and mouse NMJs13. As xenografts are innervated by the mouse host, this may result in biological changes unique to the human xenografts.
In future studies, this skeletal muscle xenograft method could be used to better understand human muscle cell biology and to develop novel models for rare or acquired muscle diseases that currently lack animal models. We anticipate that this will have a significant beneficial impact on therapeutic development for these diseases.
The authors have nothing to disclose.
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.
100 mm x 15 mm Petri dish | Fisher Scientific | FB0875712 | |
2-Methylbutane | Fisher | O3551-4 | |
20 x 30 mm micro cover glass | VWR | 48393-151 | |
Animal Weighing Scale | Kent Scientific | SCL- 1015 | |
Antibiotic-Antimycotic Solution | Corning, Cellgro | 30-004-CI | |
AutoClip System | F.S.T | 12020-00 | |
Castroviejo Needle Holder | F.S.T | 12565-14 | |
Chick embryo extract | Accurate | CE650TL | |
CM1860 UV cryostat | Leica Biosystems | CM1860UV | |
Coplin staining jar | Thermo Scientific | 19-4 | |
Dissection Pins | Fisher Scientific | S13976 | |
Dry Ice – pellet | Fisher Scientific | NC9584462 | |
Embryonic Myosin antibody | DSHB | F1.652 | recommended concentration 1:10 |
Ethanol | Fisher Scientific | 459836 | |
Fetal Bovine Serum | GE Healthcare Life Sciences | SH30071.01 | |
Fiber-Lite MI-150 | Dolan-Jenner | Mi-150 | |
Forceps | F.S.T | 11295-20 | |
Goat anti-mouse IgG1, Alexa Fluor 488 | Invitrogen | A-21121 | recommended concentration 1:500 |
Goat anti-mouse IgG2b, AlexaFluor 594 | Invitrogen | A-21145 | recommended concentration 1:500 |
Gum tragacanth | Sigma | G1128 | |
Hams F-10 Medium | Corning | 10-070-CV | |
Histoacryl Blue Topical Skin Adhesive | Tissue seal | TS1050044FP | |
Human specific lamin A/C antibody | Abcam | ab40567 | recommended concentration 1:50-1:100 |
Human specific spectrin antibody | Leica Biosystems | NCLSPEC1 | recommended concentration 1:20-1:100 |
Induction Chamber | VetEquip | 941444 | |
Iris Forceps | F.S.T | 11066-07 | |
Irradiated Global 2018 (Uniprim 4100 ppm) | Envigo | TD.06596 | Antibiotic rodent diet to protect again respiratory infections |
Isoflurane | MWI Veterinary Supply | 502017 | |
Kimwipes | Kimberly-Clark | 34155 | surgical wipes |
Mapleson E Breathing Circuit | VetEquip | 921412 | |
Methanol | Fisher Scientific | A412 | |
Mobile Anesthesia Machine | VetEquip | 901805 | |
Mouse on Mouse Basic Kit | Vector Laboratories | BMK-2202 | mouse IgG blocking reagent |
Nail Polish | Electron Microscopy Sciences | 72180 | |
NAIR Hair remover lotion/oil | Fisher Scientific | NC0132811 | |
NOD-Rag1null IL2rg null (NRG) mice | The Jackson Laboratory | 007799 | 2 to 3 months old |
O.C.T. Compound | Fisher Scientific | 23-730-571 | |
Oxygen | Airgas | OX USPEA | |
PBS (phosphate buffered saline) buffer | Fisher Scientific | 4870500 | |
Povidone Iodine Prep Solution | Dynarex | 1415 | |
ProLong™ Gold Antifade Mountant | Fisher Scientific | P10144 (no DAPI); P36935 (with DAPI) | |
Puralube Ophthalmic Ointment | Dechra | 17033-211-38 | |
Rimadyl (carprofen) injectable | Patterson Veterinary | 10000319 | surgical analgesic, administered subcutaneously at a dose of 5mg/kg |
Scalpel Blades – #11 | F.S.T | 10011-00 | |
Scalpel Handle – #3 | F.S.T | 10003-12 | |
Stereo Microscope | Accu-scope | 3075 | |
Superfrost Plus Microscope Slides | Fisher Scientific | 12-550-15 | |
Suture, Synthetic, Non-Absorbable, 30 inches long, CV-11 needle | Covidien | VP-706-X | |
1ml Syringe (26 gauge, 3/8 inch needle) | BD Biosciences | 329412 | |
Trimmer | Kent Scientific | CL9990-KIT | |
Vannas Spring Scissors, 8.0 mm cutting edge | F.S.T | 15009-08 | |
VaporGaurd Activated Charcoal Filter | VetEquip | 931401 | |
Wound clips, 9 mm | F.S.T | 12022-09 |