Introduction

Engraftment of skeletal muscle cells is commonly used to study muscle wasting, regeneration, and differentiation in the context of muscle injury and muscular dystrophies. Muscle cell transplantation has also been proposed as a therapeutic treatment for volumetric muscle loss, traumatic muscle contusion, muscular dystrophies and other myopathies (Partridge et al. 1989; Vilquin et al. 2005; Quenneville and Tremblay 2006; Halum et al. 2008; Ambrosio et al. 2010; Morosetti et al. 2011; Lavasani et al. 2014; Negroni et al. 2015; Kim et al. 2016; Barthelemy and Wein 2018). The term, “ xenograft”, is used to describe the transplantation of tissue or cells from a donor of one species into a host of a different species. Most commonly, and throughout this review, “xenograft” describes human muscle tissue or mononuclear muscle precursor cells engrafted into a rodent host.

Myoblast transfer therapy was pioneered in the 1990s with the goal of treating Duchenne muscular dystrophy (DMD) by delivering healthy, dystrophin-expressing myoblasts to dystrophic muscle groups. Though these initial trials showed little clinical benefit (reviewed in Smythe et al. 2000; Mouly et al. 2005; Skuk and Tremblay 2015), these studies provided much insight into the conditions necessary for muscle cell engraftment, as well as the factors that limit its success. Pivotal studies by Riederer et al. identified three major hurdles to successful xenotransplantation, namely early progenitor cell death, limited cell proliferation, and poor migration within engrafted muscle (Riederer et al. 2012). They showed that muscle cell differentiation within the first 3 days after engraftment prevents proliferation and migration of muscle progenitor cells and that methods to increase proliferation and inhibit early differentiation resulted in a greater number of muscle fibers formed by the engrafted cells (Riederer et al. 2012). Other groups have investigated the use of other cell types, including satellite cells, induced pluripotent stem (iPS) cells, and mesangioblasts in engraftment.

Recent advances in methods for xenografting have focused on identifying new immunodeficient mice and myogenic cell types, adapting methods to injure the host muscle prior to engraftment, and intervening to encourage proliferation of the engrafted cells and reduce proliferation of host cells after engraftment. Currently, xenografts continue to be used to study muscle differentiation and regeneration, stem cell and satellite cell biology, revascularization and reinnervation of muscle, and the control of fibrosis and liposis. For clinical purposes, xenografts are also being used to study corrected-cell and gene therapies for muscular dystrophies, including DMD, as models for Facioscapulohumeral muscular dystrophy (FSHD), and to examine potential small molecule and virally-based therapies for these and other muscle diseases (Vilquin et al. 2005; Morosetti et al. 2011; Sakellariou et al. 2016; Chen et al. 2016; Barthelemy and Wein 2018; Mueller et al. 2019). Together, these studies employ a vast array of methods. We present this comprehensive review of these methods to help researchers design and combine approaches to improve the quality of their xenografts to study muscle regeneration, muscle diseases, and potential cell therapies.

Skeletal muscle cell transplantation methods

General schema

The techniques used to xenograft muscle cells and tissue vary widely and are largely dependent on their application and the goals of the specific study. Some variables include cell type, mechanism under investigation, and limitations in time and resources that affect throughput. Generally, xenografts are generated in immunodeficient rodent hosts that have undergone some form of muscle preinjury to create a niche for muscle-derived cells or tissue to engraft (Fig. 1). The graft is typically initiated by direct intramuscular injection or by surgical implantation of myogenic cells or tissue, respectively, but systemic cell delivery is also being explored. Engraftment can be followed for several days, weeks, or months, often with interventions to enhance muscle fiber development (Fig. 1).

Fig. 1
figure 1

General schematic of xenograft transplantation method including common protocol variations

Full size image

The success of engraftment is determined by the proportion of the engrafted muscle that is of donor origin. Brimah et al. suggested that the best way to identify human regions of mosaic fibers is with human-specific antibodies to spectrin (h-spectrin) and to the nuclear envelope protein, lamin A/C (h-lamin A/C) (Brimah et al. 2004). Unlike immunolabeling for human-β-galactosidase (produced by cells transformed to express it) and, perhaps, human dystrophin, h-spectrin does not spread in host muscle and is likely to be a better indicator of myonuclear domain size (Brimah et al. 2004). Recently, however, it was reported that the h-spectrin antibody can detect false-positives in regenerating muscles of several immunodeficient mouse strains, possibly due to cross-reaction with utrophin (Rozkalne et al. 2014). Several human specific antibodies should therefore be used to ensure accurate quantification. Antibodies to human lamin A/C, spectrin, and dystrophin are most commonly used for this purpose.

Additional considerations in evaluating the success of engraftment are the ability of the engrafted human cells or tissue to repopulate the satellite cell niche, to be innervated by murine motor neurons, and to acquire an adequate blood supply. These factors will affect survival of the graft, and, most importantly for many studies, its ability to improve function. The most significant factor, however, is the requirement that the engrafted cells or tissue not be rejected by the host immune system.

Immunodeficient rodent hosts

In order for cross-species transplantation (xenografting) to be successful, the host’s immune system must not reject tissue or cells from the donor. Several immunodeficient rodent strains are available that permit human cell engraftment, including nude mice and rats, scid mice, Rag, and NOD mouse variations. The Jackson Laboratory provides a comprehensive list of immunodeficient mouse strains and their properties (https://www.jax.org/jax-mice-and-services/find-and-order-jax-mice/most-popular-jax-mice-strains/immunodeficient-mouse-and-xenograft-host-comparisons).

Some of the earliest muscle xenografting studies utilized severe combined immunodeficient (scid) mice which largely lack T and B lymphocytes (Huard et al. 1994; Skuk et al. 1999) yet still possess a partially functional innate immune system (Lovik 1995). Scid mice have been used for studies of nerve regeneration by multipotent muscle progenitor cells (Lavasani et al. 2014) and for studies of a potential cell therapy for FSHD using mesangioblasts (Morosetti et al. 2011). Recent studies of DMD have utilized the dystrophin-deficient mdx mouse crossed into the scid, nude, and other immunodeficient genetic backgrounds (Morrison et al. 2000; Farini et al. 2007; Walsh et al. 2011; Chirieleison et al. 2012; Martinez-Sarra et al. 2017; Benedetti et al. 2018; Meng et al. 2015) (reviewed by McGreevy et al. 2015). Nude mice and rats, while lacking mature T cells, are otherwise immune-competent and have limited applications for xenografting. Scid mice are preferred in some instances, although their typically short lifespan and “leaky” lymphocyte phenotype limit their usefulness for long-term studies, which require the complete absence of adaptive immunity (The Jackson Laboratory) (Cooper et al. 2001).

Borrowing from other xenografting fields, Cooper and colleagues demonstrated that the recombinase-activating gene 2 and the common cytokine receptor γ chain gene (Rag2−/γc−) double mutant mouse is superior to the scid mouse for studies of myoblast engraftment because it is completely devoid of B and T lymphocytes and produces defective natural killer cells, allowing muscle xenografts to persist long enough to reach maturity (Cooper et al. 2001). This mouse strain, as well as an alymphoid triple mutant variation lacking the C5 complement gene (C5−/Rag2−/γc−), were used by several groups to perform many of the seminal studies that led to the development of myoblast transfer therapy for DMD (Goldman et al. 1998; Cooper et al. 2001; Guigal et al. 2002; Morgan et al. 2002; Cooper et al. 2003; Brimah et al. 2004; Chicha et al. 2005; Silva-Barbosa et al. 2005; Mamchaoui et al. 2011; Riederer et al. 2012; Bencze et al. 2012; Vallese et al. 2013; Meng et al. 2015). Additional variant strains that are immunodeficient but also lack specific proteins linked to muscular dystrophies, such as α- (Sgca-null/scid/beige) or β-sarcoglycan (sgcb−/Rag2−/γc−), have also been developed and used for xenografting (Tedesco et al. 2012; Martinez-Sarra et al. 2017; Lorant et al. 2018; Torrente et al. 2004; Meng et al. 2011).

More recently the NOD scid gamma (NSG) mouse has become the leading standard in xenografting cancer and stem cells because it is able to engraft the widest range of cell types and has a longer lifespan than scid mice (The Jackson Laboratory). These mice lack T and B lymphocytes and natural killer cells and have defective dendritic cells and macrophages (The Jackson Laboratory). The NSG mice and the dystrophic NSG-mdx have been widely used for studies of satellite cell regeneration and corrected-cell therapies for DMD and other myopathies (Xu et al. 2015; Benedetti et al. 2018; Arpke et al. 2013; Young et al. 2016).

Finally, a variation of the NOD mouse, the NOD Rag gamma (NRG) mouse, was first examined in xenografting studies by Silva-Barbosa et al. (2005). This strain better tolerates the use of DNA-damaging agents and irradiation, making it the best option for studies requiring the elimination of host muscle fibers and stem cells by these methods. We have confirmed that the NOD-Rag mouse can withstand local X-irradiation of the hindlimb up to 25 Gy, which completely inhibits regeneration of murine muscle, with little or no consequences to health or life span (Sakellariou et al. 2016; Mueller et al. 2019).

A recent study of cell therapy for volumetric muscle defect repair used nude rats to engraft human myogenic precursor cells and showed functional improvements in the injured rat muscle (Kim et al. 2016). Immunocompromised Rag2-knockout rats are commercially available but to our knowledge have not yet been used in muscle grafting experiments. Rats may be particularly useful hosts when larger samples of engrafted tissue are required, but the greater availability and lower costs for purchase, breeding and housing immunodeficient mice provide substantial advantages.

Preinjury

Generating robust xenografts of muscle cells generally requires one or more forms of preinjury in order to create a niche for muscle cells to engraft (Fig. 1). Preinjury methods include ionizing radiation, mechanical disruption, and the use of myotoxic pharmacological agents. Typically, X-irradiation is used to disrupt the host satellite cell niche for studies which aim to avoid host-derived muscle regeneration, while myotoxins or mechanical injury is used to destroy host muscle fibers.

Irradiation

High dose X-irradiation is used to limit the formation of host-donor mosaic fibers during regeneration and to enhance the formation of xenografts that are highly enriched in fibers of donor origin. Host satellite cell proliferation is inhibited by 16-18 Gy X-irradiation (Morgan et al. 1990). Host satellite cells are more effectively eliminated by 18 Gy X-irradiation than by myotoxins such as BaCl2, notexin or cardiotoxin, and as a result, the irradiated muscles regenerate significantly less than the intoxicated ones (Boldrin et al. 2012). A subset of mouse host satellite cells are irradiation-resistant, however, and become activated after large scale injury (Brimah et al. 2004; Heslop et al. 2000). Higher doses are therefore needed to suppress the contributions of myogenesis of host cells to the xenografts. Slower rates of irradiation of the mouse hindlimbs significantly increased the number of fibers that formed from the donor cells (Gross et al. 1999). More recently, utilizing the irradiation-resistant NRG mouse strain (Pearson et al. 2008), our group delivered 25 Gy X-irradiation to mouse hindlimbs, shielding the body with lead to prevent damage to other tissues, and showed that the human fibers (labeled for h-spectrin) with centrally located nuclei of murine origin (not labeled for human lamin A/C) constituted less than 1.5% of all centrally nucleated myofibers (CNFs) in the xenografts (Sakellariou et al. 2016). Irradiation is also reported to increase laminin deposition in areas of damage, aiding myoblast engraftment (Silva-Barbosa et al. 2008).

Mechanical disruption

Cryoinjury has been widely used to induce muscle damage to create a niche for engrafting muscle progenitor cells. Typically, cryolesions are created by surgically exposing the muscle of interest and inducing two or more cycles of freeze-thawing with a metal rod first cooled in liquid nitrogen and then applied to the muscle for several seconds (Cooper et al. 2001; Brimah et al. 2004; Silva-Barbosa et al. 2005, 2008; Mamchaoui et al. 2011; Riederer et al. 2012; Hardy et al. 2016; Lorant et al. 2018). In a comparative study, more myogenic cells were found to be engrafted in cryoinjured TAs than in cardiotoxin-injured TAs after 4 weeks (Silva-Barbosa et al. 2005). A more physiologically relevant form of mechanical injury is performed by dropping a small steel ball directly onto the muscle of interest. This produces a high-energy blunt injury, or contusion, and has been used to study muscle allografting and the role of inflammation in muscle regeneration in mice (Ambrosio et al. 2010; Liu et al. 2017, 2018, 2019; Xiao et al. 2016). Contusion and cryoinjury induce local muscle fiber damage, while largely preserving the original basal lamina surrounding each muscle fiber. They are therefore most effective for studies of xenografting muscle precursor cells into discreet areas to study regeneration and their incorporation into regenerating muscle.

Surgical removal of large sections of muscle has also been used to create a niche for engraftment, especially prior to xenografting bundles of muscle tissue, to model repair by progenitor cells after volumetric muscle loss, and to test potential cell therapies for muscle wasting (Zhang et al. 2014; Kim et al. 2016; Kuhn et al. 2017; Baker et al. 2017). Conversely, surgical excision of the entire tibialis anterior (TA) has been used to functionally overload a synergistic muscle, the extensor digitorum longus (EDL), which induces degeneration and regeneration in the EDL (Ambrosio et al. 2009). Likewise, muscle atrophy and denervation have also been used to induce muscle wasting prior to xenografting (Lavasani et al. 2014). Each of these methods of mechanical disruption induce different forms of injury, to varying degrees of physiological relevance and, more importantly, usefulness in addressing particular questions.

Although not a mechanical disruption per se, exercise (e.g., intensive swimming) has been used prior to initiating a graft in dystrophic mice, to promote greater muscle turnover and create a niche more receptive to engraftment (Torrente et al. 2001, 2004; Benchaouir et al. 2007).

Myotoxins and chemical injuries

Several commercial myotoxins and chemicals have been developed and compared to injure muscle, leading to the death of myofibers and subsequent regeneration. Boldrin et al. (2012) showed that many more satellite cells (initially introduced exogenously) were present in muscles 3 days and 4 weeks after intoxication with BaCl2, notexin, or cardiotoxin than after X-irradiation. Notably, the three toxins were indistinguishable in this respect. A more recent study compared cardiotoxin, notexin, and BaCl2 to cryoinjury in immune-competent mice (Hardy et al. 2016). They assayed necrosis, markers of inflammation, satellite cell proliferation, fibrosis and vascularization, and showed that each method had distinct regeneration profiles. Interestingly, cardiotoxin was the only method in which cytokines were eventually restored to normal levels, whereas BaCl2, notexin and freeze injury caused inflammatory cytokines to remain elevated over the duration of the study (Hardy et al. 2016). In another report, however, levels of inflammatory cells in cardiotoxin-treated TAs in immune-compromised murine hosts were higher than in cryoinjured TAs (Silva-Barbosa et al. 2005). For cardiotoxin and notexin, there was batch to batch variation in some of the measured parameters (Hardy et al. 2016) which we have also seen in our unpublished studies (Llach, Mueller et al., in preparation). Several questions still remain, including the relative effects of these injuries in immunodeficient mice, where inflammation may be suppressed. The majority of the studies on muscle injury methods have been performed on immune-competent muscle, which may respond differently. The effects of the immune system during muscle injury have a major impact on regeneration (Tidball 2005, 2011; Yang and Hu 2018), which means immune-competent muscle may respond differently to injury than immunodeficient muscle. Therefore, a comprehensive analysis of these preinjury methods in immunodeficient mice should be investigated.

There are also several other sources of myotoxins that have not been tested in comparative studies, including cardiotoxins from different snake species and different methods for their partial or complete purification (Hodges et al. 1987; Chien et al. 1994; Mir et al. 2008). One of the most widely used forms of cardiotoxin, previously supplied by Sigma-Aldrich, is no longer produced, which has led us and other investigators in search of the closest replacement. In our experience, none of the other commercially available toxins are comparable to the Sigma-Aldrich cardiotoxin in their ability to kill murine muscle fibers in immunodeficient mice and subsequently to support engraftment. Indeed, in our studies, BaCl2 seems to be superior to commercial toxins in damaging mouse muscle fibers and for subsequent engraftment of donor cells (Llach, Mueller et al., in preparation). As dosing is limited by the fact that high concentrations of BaCl2 is cardiotoxic and can kill the treated mice, we hope that other investigators will continue to test potential myotoxins with the goal of identifying the optimal reagent for xenografting.

Cell types

The consideration of which cells are best suited for engineering of skeletal muscle tissue has been reviewed (Fishman et al. 2013). Protocols for xenografting skeletal muscle have utilized a number of different cell types for studies of regeneration after injury or atrophy, disease modeling, and cell therapy strategies. Early xenografting studies identified muscle precursor cells based on desmin immunolabeling, following primary cell isolation from muscle tissue (Cooper et al. 2001; Brimah et al. 2004; Riederer et al. 2012). Engraftment of primary cells into adult host muscle is generally difficult, however, unless the regenerative capacity of the endogenous satellite cell population has been reduced (e.g., by X-irradiation). Recently, however, methods to isolate muscle precursor cells from adult biopsy, autopsy, and fetal tissue have expanded and several discreet lineages with varying ability to form fibers and to differentiate into satellite cells in muscle xenografts have been identified (Lorant et al. 2018; Negroni et al. 2015).

Lorant et al. composed an excellent list of features of adult stem cells, distinct from satellite cells, that have been identified for potential use in xenografting, and specifically for DMD cell therapies (See Table S1 of Lorant et al. 2018). Briefly, these include myoblasts, CD133+ cells, hematopoietic stem cells, pericytes, mesangioblasts, muscle-derived stem cells, mesenchymal stem cells, myo-endothelial cells, and “side population” cells derived from interstitial tissue (Lorant et al. 2018). The properties of each cell type, including their stability, proliferation and differentiation ability, ability to integrate into host muscle and to home to muscle through the bloodstream, are also listed in Table S1 of Lorant et al. (2018), which readers should consult for details and relevant references.

In some cases, the same cell types vary in their ability to generate muscle, perhaps due to factors associated with the murine strain used and the mode of delivery (Dellavalle et al. 2007; Meng et al. 2011). Other human sources include fetal and neonatal muscle-derived stem cells and myogenic precursor cells (Brimah et al. 2004; Riederer et al. 2012; Silva-Barbosa et al. 2005, 2008), immortalized cells (Mamchaoui et al. 2011; Sakellariou et al. 2016; Mueller et al. 2019), iPS cells (reviewed in Danisovic et al. 2018), dental pulp pluripotent-like stem cells (Martinez-Sarra et al. 2017), and satellite cells (Hall et al. 2017). Muscle-derived stem/progenitor cells isolated from adult skeletal muscle have even been used to study nerve regeneration and dysphagia (Lavasani et al. 2014; Walz et al. 2008; Lavasani et al. 2013; Kuhn et al. 2017).

Immortalized cells

As noted above, successful xenografting of myogenic cells is a function of the ability of these cells to survive, proliferate, migrate throughout the muscle compartment, and eventually differentiate into myofibers in the host environment (Riederer et al. 2012). A key determinant of engraftment ability is the maintenance of myogenic precursor cells in a proliferative state. Reversible cell immortalization overcomes the issues of limited proliferation and cell cycle senescence in culture. The advantages of studying immortalized cells include the relative ease of preparation (transfection of primary cultures, followed by derivation of individual clones, if desired), the cell’s retention of the properties of the parent primary cells from which they were derived (e.g., Mueller et al. 2019), the ease of culture, and the available of essentially unlimited numbers of cells for engraftment.

Several methods of immortalization have been developed, which usually include transduction of human telomerase reverse transcriptase (hTERT) and another anti-senescence gene. hTERT maintains telomere ends so that otherwise post-mitotic cells can undergo continuous population doublings in culture. As other inhibitors of proliferation remain, however, complimentary transduction with another anti-senescence gene is necessary. Currently the two primary choices as partners of hTERT are Bmi-1 and CDK4 (Cudre-Mauroux et al. 2003; Zhu et al. 2007). Bmi-1 negatively regulates cell cycle-repressing genes p16 and p19Arf, allowing cells to avoid senescence (Jacobs et al. 1999). CDK4 directly regulates the cell cycle, allowing cells to progress through G1 phase. Both methods overcome problems with previous immortalization methods that resulted in major chromosomal abnormalities and phenotypic changes (Cudre-Mauroux et al. 2003; Zhu et al. 2007). DMD cells immortalized with Bmi-1 and hTERT are currently being used to develop corrected-cell therapies for DMD, while cells immortalized with CDK4 and hTERT are being used to model FSHD and to study other complex muscular dystrophies (Benedetti et al. 2018; Stadler et al. 2011; Sakellariou et al. 2016; Mamchaoui et al. 2011; Arandel et al. 2017; Mueller et al. 2019).

Satellite cells

Recent studies have made significant advances in the methods for transplanting satellite cells and activated muscle stem cells (Sacco et al. 2008; Boldrin et al. 2012; Boldrin and Morgan 2012; Charville et al. 2015; Hall et al. 2017). Several have also begun to clarify the similarities and differences in Pax7 expression, cell-surface marker expression, and the regenerative potential of human compared to murine satellite cells (Reimann et al. 2004; Boldrin and Morgan 2012; Xu et al. 2015; O’Connor et al. 2009; Decary et al. 1997). To assess their characteristics in vivo, Xu et al. isolated satellite cells from biopsies of several human muscles and engrafted them into the mouse TA (Xu et al. 2015). They showed that xenografted CD56+/CD29+ human myoblasts reconstituted the satellite cell niche and formed several clusters of human fibers within the graft. Upon reinjury, they saw robust expansion of the engrafted human muscle fibers, indicating that the regenerative capacity of these human satellite cells is maintained. Importantly, they saw no major differences in the relative quantity or ability of the human satellite cells to engraft between the biopsied muscles groups, suggesting that large numbers of satellite cells can be isolated from multiple muscle groups from a single human donor (Xu et al. 2015). Ishii et al. (2018) found similar results with satellite cells that were isolated and maintained in the presence of laminin E8 to promote their “stemness”. Upon engraftment, these cells repopulated the satellite cell niche and promoted muscle regeneration (Ishii et al. 2018).

The ability of myogenic cells to repopulate the satellite cell niche is necessary for studies of the possible role of satellite cells in disease and regeneration between healthy and diseased human muscle. Certain cell types are able to generate both muscle fibers and to become Pax7+ satellite cells upon transplantation. Among the cell types capable or repopulating the satellite cell niche are blood- and muscle-derived CD133+ cells (Torrente et al. 2004; Benchaouir et al. 2007); human mesoangioblast/pericyte-derived cells (Dellavalle et al. 2011), side-populations, interstitial cells (Asakura et al. 2002) and normal or immortalized human muscle precursor cells (Ehrhardt et al. 2007; Mueller et al. 2019).

iPS cells

Several laboratories have recently begun to explore the use of iPS cells in skeletal muscle grafts. Pluripotent stem cells can be selected for their myogenic capacity, or programmed to become myogenic by transformation to express Pax3 and/or Pax7, or MyoD, or by the addition of factors to the culture medium (Mizuno et al. 2010; Darabi et al. 2011; Swartz et al. 2016; Torihashi et al. 2015). These cells have several advantages, including the ease of preparing them from individuals with a wide range of muscle disorders without requiring biopsies, and, like immortalized human muscle precursor cells, their availability in unlimited numbers in culture. They are already being examined in the context of different diseases of muscle (Jiwlawat et al. 2019; Nakajima et al. 2019; Steele-Stallard et al. 2018; Salani et al. 2012; Kim et al. 2019); (reviewed in Piga et al. 2019; Xia et al. 2018; Danisovic et al. 2018; Pourquie et al. 2018). In xenografting studies, iPS cells transfected to express Pax3/Pax7 engraft better than fetal myoblasts and reseed the satellite cell niche in the engrafted muscle (Incitti et al. 2019). Similarly, myogenic cells created by expressing Pax3 alone in human iPS cells developed into small bundles in tissue culture that remained functional upon engraftment into immunocompromised mice (Rao et al. 2018). Zhao et al. reprogrammed fibroblasts from dystrophic mdx mice with site-specific recombinases to create dystrophin-positive myogenic precursors that successfully engrafted into mdx muscles, albeit in relatively low numbers (Zhao et al. 2014). Similar studies were reported by Cai et al. (2016), Goudenege et al. (2012) and Beck et al. (2011), although with different cell populations and transformation methods. Tedesco et al. (2012) derived mesangioblast-like cells from iPS cells isolated from α-sarcoglycan-null mice, corrected the genetic deficit, and introduced them into sarcoglycan-null immunodeficient mice. They found limited regions of the muscles that were successfully engrafted, as indicated by the presence of α-sarcoglycan.

Although iPS cells require considerable effort to prepare and characterize prior to use in transplantation studies, they may well define the future of xenografting to study human muscle development, examine the pathogenic mechanisms of different myopathies and muscular dystrophies, and test a range of therapeutics in preparation for clinical trials. Their use in treating diseases of muscle is also under consideration (Maffioletti et al. 2014; Hagan et al. 2018; Pareja-Galeano et al. 2016; Loperfido et al. 2015).

Cellular properties and culture conditions affecting engraftment

Importantly, the age of donor cells, measured by age of the donor, mean population doublings prior to engraftment, and telomere length, have all been reported to affect regenerative capacity (Decary et al. 1997; Cooper et al. 2003; Brimah et al. 2004; Schafer et al. 2006). The greater the number of population doublings and the shorter the telomere length, the lower the ability to engraft, as measured by the number of donor fibers formed in xenografts of muscle precursor cells (Decary et al. 1997; Cooper et al. 2003; Brimah et al. 2004). The severity of the donor’s myopathy is also inversely correlated with xenografting capacity, perhaps due to progenitor cell exhaustion in affected muscles (Schafer et al. 2006; Morosetti et al. 2007). Culture conditions can also impact the myogenicity of donor cells. Our group and others have observed that culturing muscle precursor cells under conditions that increase proliferation, while suppressing differentiation, increases the fiber number within xenografts formed by those cells. Some of these conditions include the use of high serum concentrations, culture on laminin, heat shock treatment, and co-culture with IGF-1, bFGF and others (Silva-Barbosa et al. 2008; Sakellariou et al. 2016; Brimah et al. 2004; Danisovic et al. 2018; Riederer et al. 2008).

Cell delivery

The method of cell delivery is also an important consideration in xenograft studies. While most studies have relied on single intra-muscular injection, progressive cell delivery (multiple injections) and systemic delivery have been explored. Kim et al. (2016) showed that progressive intra-muscular injections of cells over 4 weeks restored greater muscle force, increased muscle mass and differentiation, reduced fibrosis and increased vascularization in a volumetric muscle defect repair model, as compared to a single injection. Several studies suggest CD133+ muscle progenitor cells isolated from muscle or blood could be introduced systemically to deliver gene therapies for DMD, although engraftment of patient-derived CD133+ cells, in particular, has only been variably successful (Torrente et al. 2004, 2007; Gavina et al. 2006; Benchaouir et al. 2007; Negroni et al. 2009; Meng et al. 2014, 2018). Mesangioblasts and other pericytes may also be systemically delivered, and have been characterized in xenografts as a potential cell therapy approach for FSHD (Morosetti et al. 2007, 2011). Pericytes delivered intra-arterially in three bouts were able to colonize and generate new muscle fibers in hindlimb muscles of scid/mdx mice (Dellavalle et al. 2007), but, as noted above, another study using intra-arterial injection reported that muscle-derived cells did not contribute to regeneration in all hosts (Meng et al. 2011). The variability in the success of xenografting following systemic introduction of donor cells suggests that critical factors remain to be discovered and refined before this method can be more widely adopted.

Interventions to enhance engraftment

Once myogenic precursor cells are delivered to muscle, several methods of intervention can be used to enhance engraftment (Fig. 1). Due to the inverse relationship between the proliferation of myogenic cells and their differentiation into myotubes and myofibers, interventions aimed at increasing differentiation should start at least 3 days after engraftment (Riederer et al. 2012). Interventions can largely be divided into two main categories: exercise-mimicking and molecular.

Interventions that mimic exercise can include voluntary exercise, such as housing mice in cages with running wheels or performing bilateral hindlimb engraftment which encourages the mouse to use both legs in order to ambulate (Llach, Mueller et al., in preparation). Functional overloading of the EDL by surgical removal of the TA, treadmill exercise, and swimming protocols have also been used to improve engraftment of muscle derived stem cells (Ambrosio et al. 2009, 2010; Fakhfakh et al. 2012a, b). Intermittent neuromuscular electrical stimulation, performed while the mouse is under anesthesia, also increases engraftment by increasing fiber number, size, and maturity (Sakellariou et al. 2016; Mueller et al. 2019).

Molecular interventions to improve tissue and cell engraftment include local or systemic injection of small molecules, synthetic matrices, growth factors, and myostatin and TGF-β inhibitors (Brimah et al. 2004; Fakhfakh et al. 2012a, b; Gerard et al. 2012). Of these, the best studied are interventions which modulate the TGF-beta signaling pathway, known to play an important role in muscle regeneration. In myopathies that involve dysregulated muscle repair, myostatin, a TGF-β superfamily myokine, and TGF-β overexpression at the site of muscle injury inhibit cell proliferation, delay differentiation, and promote the formation of fibrosis in place of degenerating myofibers (reviewed in Burks and Cohn 2011; Kim and Lee 2017). Blocking myostatin signaling by either overexpressing follistatin, an endogenous myostatin antagonist, or a dominant-negative form of the TGF-β superfamily receptor, ActRIIB, improves the transplantation of human myoblasts in Rag/mdx TAs (Benabdallah et al. 2008; Fakhfakh et al. 2011). These experiments were followed by methods that could be applied to humans such as systemic delivery of ActRIIB/Fc and oral treatment with Losartan, a small molecule that decreases TGF-β expression. Combined with forced swimming to encourage muscle turnover, ActRIIB/Fc treatment led to increased body weight and increased hypertrophy of dystrophin-positive fibers in the TAs of Rag-null/mdx mice engrafted with human myoblasts (Fakhfakh et al. 2012a, b). Losartan treatment increases myoblast proliferation and fusion, while decreasing negative regulators of muscle repair (Fakhfakh et al. 2012a, b). Combined, these studies show that modulating the activity of the TGF-β superfamily can be used to increase myogenicity in xenografting protocols.

Co-injection of fibrin gel with human myoblasts significantly increases cell survival after implantation and the number of dystrophin-positive fibers formed at 3 weeks post-engraftment in Rag/mdx mice (Gerard et al. 2012). A pro-survival cocktail, containing Matrigel, IGF-I and several other factors, provides the same benefit (Laflamme et al. 2007; Gerard et al. 2012). Keratin hydrogels, biodegradable polymers, decellularized tissue scaffolds, and growth-factor releasing matrices have also been suggested as potential tools for muscle repair (Saxena et al. 1999; Lee et al. 2000; Richardson et al. 2001; Levenberg et al. 2005; Baker et al. 2017; Urciuolo and De Coppi 2018), although comprehensive studies of their utility in xenografting have not yet been performed.

Recent muscle xenograft applications

FSHD xenograft models and cell therapy

FSHD is a debilitating muscle wasting disease caused by the pathogenic misexpression of the transcription factor, DUX4. DUX4 is a primate-lineage gene located within 3.3 kb repeated units in the distal end of chromosome 4 (reviewed in Tassin et al. 2013). The downstream gene targets of the mouse Dux paralog are unlike the targets of DUX4 (Sharma et al. 2013; Eidahl et al. 2016), and the epigenetic regulation of DUX4 expression is likely dependent on other human genes (reviewed in Daxinger et al. 2015; DeSimone et al. 2017; Hamel and Tawil 2018; Himeda et al. 2018). Thus, models of FSHD that reproduce the genetic and epigenetic features of mature human muscle cannot be readily generated by standard transgenic or knockout methods. Recently, two novel models were developed by engrafting immortalized human muscle cells or muscle fibers into the mouse hindlimb to allow for studies of endogenous human gene expression in a living tissue (Chen et al. 2016; Sakellariou et al. 2016; Mueller et al. 2019).

Chen et al. used a xenograft model of FSHD to test the efficacy of an antisense phosphorodiamidate morpholino oligonucleotide (PMO) to suppress DUX4 expression (Chen et al. 2016). They surgically removed the TA and EDL muscles from NRG mice and engrafted 8 × 3 × 1 mm bundles of biopsy or autopsy human donor muscle collected from FSHD patients and control relatives into the anterior compartment of the mouse hindlimb (Zhang et al. 2014; Chen et al. 2016). After 4–6 months they electroporated the engrafted muscle with the PMO and collected the tissue 2 weeks later. They saw a reduction in DUX4 and DUX4-target expression in FSHD patient-derived muscle xenografts treated with the PMO, providing evidence that antisense PMOs may be effective in treating FSHD (Chen et al. 2016). This method, while effective for testing proof-of-principle FSHD therapies, is limited by the difficulty of obtaining muscle biopsies, the time it takes to achieve sufficient engraftment, the variability in the quality of the grafts it produces, and the relatively small number of xenografts that can be generated by each biopsy.

Our laboratory has developed a xenograft model of FSHD which avoids most of these issues. Most importantly, it is scalable, to allow for larger studies of FSHD pathophysiology and in vivo testing of potential FSHD therapeutics. Our method uses immortalized human muscle precursor cells (hMPCs) from FSHD patients and control relatives engrafted into the TA of NRG mice, which has been irradiated and intoxicated by myotoxins to create a niche for new human-derived muscle to develop (Sakellariou et al. 2016). We see robust engraftment by 4 weeks and the genetic and epigenetic profiles of our xenografts replicate those of FSHD patients (Sakellariou et al. 2016; Mueller et al. 2019). An advantage of our method is that it occasionally produces a TA muscle in the mouse that is completely human in origin. The disadvantage is that this is infrequent, as the xenografts typically contain only 400–900 human fibers. However, because we are using immortalized FSHD cell lines, rather than primary cells or bundles of muscle fibers, we are able to generate an almost infinite number of genetically identical grafts for studies of both FSHD pathophysiology and drug screening. We are currently testing several potential therapeutic compounds and gene therapies to treat FSHD.

The potential for cell therapy to treat FSHD has also been studied using xenografts. FSHD myoblasts (Vilquin et al. 2005) or mesangioblasts (Morosetti et al. 2011) isolated from unaffected muscles of patients contribute to the regeneration of murine muscle fibers after myotoxin or cryoinjury, respectively. One report described a defective homing mechanism in FSHD mesangioblasts derived from severely affected muscles, which should be considered in designing autologous cell therapies (Morosetti et al. 2011). Combined, the FSHD xenograft models and FSHD cell transfer studies suggest that xenografting will be a pivotal technique going forward to elucidate the mechanism of muscle wasting in FSHD and to design and test therapeutic strategies to treat this debilitating disease.

Xenografts in studies of Duchenne muscular dystrophy

DMD is an X-linked disease caused by mutations in the dystrophin gene, which is required for sarcomeric and sarcolemmal integrity (Mokri and Engel 1975; Hoffman et al. 1987; Porter et al. 1992). Several xenograft strategies to study and treat DMD have been reported, including those that involve gene therapy, cell therapy, or a combination of the two, which aim to replenish the muscle with dystrophin. These have been extensively reviewed (Shimizu-Motohashi et al. 2016; Barthelemy and Wein 2018; Shimizu-Motohashi et al. 2019; Danisovic et al. 2018; Lorant et al. 2018) and suggest that novel methods of engraftment have the potential to lead to new approaches to treating DMD.

Pericytes, but not satellite cells, isolated from DMD biopsies and corrected with human mini-dystrophin, were delivered to muscle tissue by intra-arterial injection. These cells were able to traffic to and engraft the TA of scid/mdx mice to form dystrophin-positive muscle fibers (Dellavalle et al. 2007). Similarly, in another study using intra-arterial delivery, exon-skipped DMD-CD 133+ cells colonized the TA and were able to improve muscle morphology, function, and dystrophin expression in scid/mdx mice (Benchaouir et al. 2007). However, a more recent study concluded that while DMD-CD133+ cells may regenerate muscle fibers in muscle xenografts, they do not repopulate the satellite cell niche like healthy donor-derived CD133+ cells (Meng et al. 2018). This lack of myogenic potential may be due to exhaustion of the muscle stem cells (Heslop et al. 2000), leading to an increase in CD133+ expression in non-myogenic cells (Meng et al. 2018). In another study, CRISPR/Cas9 reframed human iPS-derived skeletal muscle cells engrafted into the TAs of NSG-mdx mice and led to the expression of dystrophin and β-dystroglycan, restoring the dystrophin-glycoprotein complex (Young et al. 2016). Human adipose-derived mesenchymal stem cells delivered either locally or systemically have also shown great promise for xenotransplantation, expressing dystrophin in muscles of mdx mice and golden retriever muscular dystrophy dogs (Rodriguez et al. 2005; Vieira et al. 2012).

In one of the most promising series of studies to date, Tedesco et al. corrected mdx-derived mesangioblasts using a human artificial chromosome (HAC) encoding human dystrophin and engrafted those cells into mdx mice. This yielded dystrophin-positive fibers and satellite cells that persisted in the host for up to 8 months (Tedesco et al. 2011). This study led to further studies, which showed that HACs encoding dystrophin can be introduced into immortalized DMD-patient-derived myoblasts or mesangioblasts and then engrafted into the TAs of scid/mdx mice to form dystrophin-corrected muscle fibers (Benedetti et al. 2018). These authors also designed a synthetic HAC which expresses all of the necessary components for this strategy to be used as a novel gene therapy: genes for controlled levels of dystrophin protein expression, reversible immortalization, inducible differentiation, and regulated cell death (Benedetti et al. 2018). These and other xenograft studies of potential cell therapies including TALEN and CRISPR-corrected DMD-patient-derived iPS cells are ongoing (Li et al. 2015). Future publications will reveal if these strategies are successful.

Discussion

Xenografts of human muscles in mice provide an important tool for studies of human muscle physiology and pathophysiology, as well as unique tissues with which to test the specificity and efficacy of drugs targeted to human diseases in general and to diseases of muscle in particular. Ideally, the xenografts should have a large number of myofibers of human origin in a defined portion of the muscle, that can be isolated and studied without significant contamination by murine host tissue. Most studies have not had this as an aim, however. Rather, they have aimed to learn if engraftment methods can be used to restore normal gene expression and suppress pathophysiology in murine models of diseases such as DMD. These studies have produced promising results, but their applicability to human health is so far limited by the difficulties in introducing human myogenic precursor cells into patients by systemic injection, such that they access all the skeletal muscles of the human body affected by dystrophinopathy, while avoiding attack by the immune system.

Xenografting human myogenic cells into murine muscle to achieve muscles that are exclusively human in origin to model muscular dystrophies is also technically challenging, although noteworthy strides in this direction have recently been made. The major difficulty so far is eliminating all of the host muscle fibers within a particular compartment, and then ensuring that the hMPCs that are introduced to replace them distribute widely enough and are robust enough to repopulate the muscle compartment with human fibers. The fact that intoxicated muscles tend to degenerate only partially seems to be the major stumbling block here, likely because the myotoxins typically used in these experiments do not access all of the sub-compartments of the muscle. The reason for this is unclear, although it may be due to the persistence of diffusion barriers created by connective tissue. Multiple injections into widely spaced sites might solve this problem, but the toxicity to mice of some of the reagents, such as BaCl2, limits the usefulness of this approach. We were only able to generate intact TA muscles with almost exclusively human myofibers using the cardiotoxin preparation from Sigma, which was never fully defined chemically and which is no longer commercially available. In its continued absence, other approaches, such as freezing and thawing the host muscle, may be the better alternative.

Limiting the ability of the endogenous murine tissue to regenerate is also a significant issue. X-irradiation is so far the most widely used method to do so, but most immunodeficient murine strains are damaged by the doses of X-rays needed to entirely eliminate the contribution of murine satellite cells to the xenograft, and even NRG mice, which are more radiation-resistant, show occasional skin lesions following irradiation of their hindlimbs. Other side effects of X-irradiation, such as inflammation, may also occur. Alternative radiation sources, such as gamma radiation, may prove to be both gentler and more efficient (Caiozzo et al. 2010). To date, gamma irradiation has not yet been tested in muscle for its usefulness in xenografting experiments.

The literature is clear about the best sources of donor cells for engraftment, however, at least when numerous grafts containing large numbers of fibers of human origin are needed. Immortalized myogenic precursor cells, cultured with a minimal number of population doublings, have now been widely used and have in most cases given excellent results. They are easy to grow in large numbers and provide an almost unlimited source of identical donor material. This provides a significant advantage over transplantation of primary cells or tissues, including intact human muscle fibers or fiber bundles, which are by definition limited to clinical samples obtained through biopsies, which is neither always possible nor without risk. Furthermore, immortalized hMPCs can also be transfected to express particular proteins or other macromolecules, allowing very specific tests of the roles of these molecules in disease or disease prevention. The major disadvantage of the immortalized cells is that immortalization by hTERT and either Bmi-1 or CDK4 makes the cells potentially tumorigenic and thus unsuitable for any clinical studies unless they are remortalized first. Remortalization is possible if the transgenes are introduced between sites susceptible to Cre or other excision sequences, but this would have to be very efficient, indeed, to eliminate the possibility of tumor formation following clinical use. Tumorigenicity of hMPCs in mice has not been a common problem in xenografting studies, however, making them nearly ideal for studies of pathogenic mechanisms and of therapies for muscle diseases. As the field progresses, hMPCs may ultimately be replaced by iPS cells, which have the advantage that they can be more easily derived from patients with different genetic variations, but methods for xenografting of iPS cells are still not well developed.

Although the discussion above summarizes the many advantages of xenografting, the approach has several disadvantages. First, success in generating large, robust xenografts requires the use of immune-compromised mice, which means that the contribution of the immune system to diseases and disease progression cannot be addressed with these methods (Tidball 2011; Yang and Hu 2018). As a cautionary tale, allografts of bone marrow mesenchymal stem cells in immune-competent mice resulted in increased inflammation, oxidative stress, fibrosis, and impaired regeneration (Liu et al. 2019). This is also a serious drawback for studies of the inflammatory diseases of muscle, such as polymyositis and dermatomyositis, as well as of autoimmune diseases of muscle, such as myasthenia gravis. Another drawback is that, unless introduction of the donor cells is systemic, xenografting is typically limited to a particular muscle or muscle group, and thus cannot recapitulate the rostro-caudal or dorso-ventral position of the muscles affected by disease in the human body. Although they can reach maturity, the muscles in the grafts may not fully report on the responses of muscles in aging individuals or in individuals with diseases that have progressed significantly. Finally, the methods are quite labor intensive and are thus much more difficult to generate and maintain than spontaneous, transgenic or knock-out murine models of human muscle diseases.

Human-to-mouse muscle xenografts have one obvious advantage over other murine models, however: they are comprised of human tissue and thus provide the best source of mature human muscle, outside of the clinic, to study the specificity and efficacy of drugs designed to treat human diseases. One example of particular interest to us is the observation that overexpression of DUX4, the pathogenic agent in FSHD, causes many genetic changes in murine muscle that are not seen in human FSHD muscle. Because of their ability to report specifically on the biochemical and molecular responses of human muscle, even if they may not yet reliably report on physiological changes, we believe that xenografts should be used by academic laboratories and pharmaceutical companies routinely as part of preclinical drug trials.

The range of options open to researchers interested in using xenografts in their studies is large but it should not be daunting. Some methods are clearly adapted to study particular questions related to muscle (e.g., satellite cell biology, the satellite cell niche, muscle regeneration, etc.) whereas others are more suited to examine disease mechanisms or to test potential therapeutics. There are nevertheless several key questions in muscle biology that may be addressable with xenografting methods that have not yet been tried, such as those related to the formation and extent of myonuclear domains, and the dosage of particular genes required to keep fibers healthy or to cause disease. Our aim has been to summarize the field and to encourage investigators not only to use established methods, but also to test new approaches that may be applicable to these and other compelling questions in muscle biology.