This protocol outlines the isolation of skeletal and cardiac muscle fibro-adipogenic progenitors (FAPs) from spiny mouse (Acomys) via enzymatic dissociation and fluorescence-activated cell sorting. The FAPs obtained from this protocol can be effectively expanded and differentiated to myofibroblasts and adipocytes.
Due to its exceptional repair program, the spiny mouse is an emerging research model for regenerative medicine. Fibro-adipogenic progenitors are tissue-resident cells that are able to differentiate into adipocytes, fibroblasts, and chondrocytes. Fibro-adipogenic progenitors are fundamental for orchestrating tissue regeneration as they are responsible for extracellular matrix remodeling after injury. This study focuses on investigating the specific role of fibro-adipogenic progenitors in spiny mouse cardiac repair and skeletal muscle regeneration. To this end, a protocol has been optimized for the purification of spiny mouse fibro-adipogenic progenitors by flow cytometry from enzymatically dissociated skeletal and cardiac muscle. The population obtained from this protocol is capable of expanding in vitro, and can be differentiated to myofibroblasts and adipocytes. This protocol offers a valuable tool for researchers to examine the distinctive properties of spiny mouse, and to compare them to the Mus musculus. This will provide insights that could advance the understanding of regenerative mechanisms in this intriguing model.
Initially recognized for its exceptionally fragile skin and remarkable ability to repair skin injuries, spiny mouse has demonstrated superior regenerative capacity in various organ systems, such as musculoskeletal, renal, central nervous system, and cardiovascular, when compared to Mus musculus1,2.
Fibro-adipogenic progenitors (FAPs) are a subset of stromal cells that are resident in various tissues, including skeletal and cardiac muscle. These cells possess a unique capacity to commit to fibrogenic and adipogenic lineages in vivo and in vitro3,4. FAPs play a crucial role in tissue regeneration by modulating the extracellular matrix and supporting the functions of other cell types involved in the repair process. In skeletal muscle, FAPs become activated in response to injury and facilitate muscle stem cell differentiation and myogenesis5,6. In ischemic injury to the heart, FAPs lay down scar tissue to maintain the integrity of the myocardium. In contrast to muscle, cardiac FAPs become chronically activated and contribute to pathological remodeling7,8.
Previous studies have demonstrated that spiny mouse extracellular matrix has different composition, structure, and properties that support regeneration in comparison to Mus musculus9,10. In muscle and heart injuries, FAPs have been noted to contribute to superior healing in spiny mouse11,12,13. Understanding the behavior and regulatory control of spiny mouse FAPs and stroma may shed light on the mechanism behind their regenerative capabilities. While FAPs are extensively studied in other animal models, such as in mus musculus14,15, currently, there are no published protocols for isolating spiny mouse FAPs. Developing such a protocol would fill a significant gap in the field and enable researchers to examine the cellular and molecular mechanisms underlying the spiny mouse's regenerative potential.
This protocol describes a robust and reproducible protocol for isolating, expanding, and differentiating skeletal and cardiac muscle FAPs from spiny mouse. The protocol described here yields high-quality single-cell suspensions that are suitable for fluorescence-activated cell sorting (FACS). By using rh-TGFβ1 or a compatible commercially available media, two methods widely used to differentiate mus musculus FAPs16, the sorted spiny FAPs maintain their capacity to differentiate along the fibrogenic lineage and adipogenic lineage, respectively (Figure 1).
All animal maintenance and experimental procedures were conducted in accordance with the approval of the University of British Columbia Animal Care Committee and the regulations at the University of British Columbia. Animals were housed in an enclosed pathogen-free facility under standard conditions (12:12 light-dark cycle, 21-23 °C, and 40%-60% humidity level) and provided a protein-rich mouse diet and water ad libitum. Adult (4 to 6 months old, 50-60 g) female and male Acomys dimidiatus mice were used for this study. The details of all the reagents and equipment used are listed in the Table of Materials.
1. Solution and buffer preparation
2. Tissue collection
3. Tissue digestion
4. Staining for Fluorescence-activated cell sorting (FACS)
5. Fluorescence-activated cell sorting
6. Tissue culture
7. Fibrogenic differentiation
8. Adipogenic differentiation
9. Immunostaining
The schematic for this protocol to isolate and culture skeletal muscle and cardiac FAPs is summarized in Figure 1. For tissue collection, the liver changing color from dark red to pale yellow is usually indicative of a successful perfusion. With the specified age ranges of spiny mouse, the heart weights are typically around 200 mg, while the quadricep muscle is around 350 mg.
During each digestion buffer change in steps 3.5-3.7, the digestion buffer in the digestion tubes appears opaque as cells are released from the tissue. At the end of the digestion, the solution must be a relatively homogenous slurry with some tissue pieces remaining. Be sure to end the digest before all tissues are completely digested to avoid over-digestion, which will negatively impact the viability of the cells and culture outcomes.
The expected flow plots for FACS are shown in Figure 2. FAPs are expected to represent around 2% of live and LIN- cells. The percentage of PI+ events is an indicator of the quality of the sample preparation and should be less than 10%. In this study, 150k FAPs per heart (ventricles only) and 100k FAPs from two quadricep muscles are typically obtained.
Once in the plate, cells typically attach within 72 h, reach 80% confluency within 5 days, and display typical fibroblast morphology with projections (Figure 5).
Figure 1: Schematic representation of the protocol and approximate time required for each step. Tissue collection: 10 min per mouse. Tissue digestion: 1.5-2 h. Cell staining: 45 min-75 min. Fluorescence-activated cell sorting: 2 h. Fibro-adipogenic progenitors (FAP) expansion: 4 days. FAP differentiation: 48 h for fibrogenic differentiation, 6 days for adipogenic differentiation for skeletal muscle FAPs, and 14 days for cardiac FAPs. Please click here to view a larger version of this figure.
Figure 2: Representative FACS plots for skeletal muscle (top) and cardiac (bottom) FAP isolation. Please click here to view a larger version of this figure.
Figure 3: Spiny mouse skeletal muscle FAP differentiation. (A) Fibrogenic differentiation control. (B) Fibrogenic differentiation. (C) Adipogenic differentiation control. (D) Adipogenic differentiation. Perilipin (green), SMA (magenta), DAPI (blue), scale bar = 100 µm. Please click here to view a larger version of this figure.
Figure 4: Spiny mouse cardiac FAP differentiation. (A) Fibrogenic differentiation control. (B) Fibrogenic differentiation. (C) Adipogenic differentiation control. (D) Adipogenic differentiation. Perilipin (green), SMA (magenta), DAPI (blue), scale bar = 100 µm. Please click here to view a larger version of this figure.
Figure 5: Brightfield images of spiny mouse skeletal muscle (A) and heart (B) FAPs after 5 days of culture, before differentiation. Scale bar = 360 µm. Please click here to view a larger version of this figure.
Controls | CD31 SC | PDGFRα SC | CD31 FMO | PDGFRα FMO | Sample |
CD31 antibody | + | – | – | + | + |
PDGFRα antibody | – | + | + | – | + |
Table 1: Schematic guide for setting up antibody staining cocktails for controls and sample. SC = single color control, FMO = fluorescence minus one control, + = with the corresponding antibody, – = without the corresponding antibody.
Spiny mouse tissues are more sensitive to the stresses in tissue dissociation, and there are several aspects of this protocol aimed at minimizing stress to improve cell viability. Serial enzymatic dissociation technique is employed for sample preparation to lower the concentration of the enzyme required. As enzymatic digestion proceeds, the enzymatic activity decreases. By replacing it with fresh enzymes, consistent enzymatic activity throughout the entire digestion can be better achieved, and high concentrations of the enzyme are avoided. Furthermore, changing the digestion buffer to over-digest the already released cells in the solution is avoided. Secondly, ACK lysis buffer is used to remove red blood cells. Although the heart is exsanguinated and perfused, it still contains red blood cells as it is a highly vascularized organ. By removing contaminating red blood cells, the time required for FACS is minimized to avoid cell degradation. Lastly, it is important to work fast and diligently from step 2.1 to step 6.4, and follow best practices for working with cells, such as gentle pipetting and keeping on ice as much as possible. Some details are difficult to show even in the audiovisual format and may require a certain level of experience.
In cell culture, the initial seeding density is critical for optimal cell expansion. FAPs in the dish communicate in a paracrine fashion and stimulate growth via secreted growth factors. In low seeding density conditions, the cells do not receive sufficient signals for growth and will either fail to expand or require additional culturing time to reach appropriate confluency levels for downstream applications. This is also the rationale for changing only half of the media during each media change in adipogenic differentiation. It is important to leave some conditioned media that contains the growth factors from pre-adipocytes that are critical for their differentiation.
During the immunostaining, it is important to be gentle when adding or removing buffers to minimize cell detachment. In particular, in step 9.1, the cells are the most prone to detach prior to fixation. Manual aspiration, such as with a pipette or a needle and syringe, is recommended, but the use of powerful vacuum aspirators is not advised.
A limitation of this protocol is that PDGFRα is a general marker of the FAPs, and more markers are required to distinguish the unique subtypes of FAPs17. Subtypes such as DPP4+ progenitors like FAPs and CD10+ adipogenic FAPs have been implicated in both disease and homeostatic context18,19,20. FAP subsets likely also exist in spiny mouse. They may exist in different proportions, exhibit different cellular behavior, or have different physiological functions compared to corresponding mus musculus FAP subsets. Further optimization of FAP subtype-specific markers in spiny mouse will be needed to examine the details of spiny mouse FAPs.
In summary, this protocol details a robust and reproducible method of isolating, culturing, and differentiating skeletal muscle and cardiac FAPs from spiny mouse. The spiny mouse is gathering momentum as a new hyper-regenerative animal model. FAPs and the stroma are highly implicated in maintaining muscle and cardiac homeostasis and contribute to organ dysfunction in multiple chronic disease models5,7. Characterizing spiny mouse FAPs may elucidate the mechanism behind their remarkable regenerative capacity and pave the way toward novel strategies in regenerative medicine.
The authors have nothing to disclose.
We would like to acknowledge Andy Johnson and Justin Wong, UBC flow core, for their expertise and help in optimizing the FACS protocol, as well as the UBC Biomedical Research Center animal facility staff for spiny mouse care. Figure 1 has been made using Biorender. Figure 2 has been made using FlowJo software.
0.5M EDTA | Invitrogen | 15575–038 | |
1.7 mL Microcentrifuge tubes | VWR | 87003-294 | |
15 mL centrifuge tube | Falcon | 352096 | |
1x Dulbecco’s Phosphate Buffered Saline (DPBS) | Gibco | 14190-144 | |
20 mL syringe | BD | 309661 | |
4′,6-diamidino-2-phenylindole (DAPI) | Invitrogen | D3571 | |
40 μm cell strainer | Falcon | 352340 | |
48 well flat-bottom tissue culture plate | Falcon | 53078 | |
5 mL polypropylene | Falcon | 352063 | |
5 mL polystyrene round-bottom tube with cell-strainer cap | Falcon | 352235 | |
5 mL syringe | BD | 309646 | |
50 mL centrifuge tube | Falcon | 352070 | |
60 mm Petri dish | Falcon | 353002 | |
96 well V-bottom tissue culture plate | Corning | 3894 | |
Acomys dimidiatus mice (spiny mice) | kindly gifted by Dr. Ashley W. Seifert (University of Kentucky). | ||
Ammonium-chloride-potassium (ACK) lysing buffer | Gibco | A10492-01 | |
Anti-perilipin (1:100) | Sigma | P1873 | |
Anti-SMA (1:100) | Invitrogen | 14-9760-82 | |
APC PDGFRa (1:800) | Abcam | ab270085 | |
BD PrecisionGlide Needle 18 G | BD | 305195 | |
BD PrecisionGlide Needle 23 G | BD | 305145 | |
Bovine serum albumin | Sigma | A7906-100g | |
BV605 CD31 (1:500) | BD biosciences | 744359 | |
CaCl2 | Sigma-Aldrich | C4901 | |
Centrifuge | Eppendorf | 5810R | |
DMEM/F12 | Gibco | 11320033 | |
Donkey anti-mouse Alexa 555 (1:1000) | Invitrogen | A31570 | |
Donkey anti-rabbit Alexa 647 (1:1000) | Invitrogen | A31573 | |
Donkey serum | Sigma | S30-100ML | |
FACS sorter – MoFlo Astrios 5 lasers | Beckman coulter | B52102 | |
Fetal bovine serum | Gemini | 100-500 | |
Fine scissors | FST | 14058-11 | |
Fluoromount-G | SouthernBiotech | 0100-01 | |
Forceps | FST | 11051-10 | |
Hemostat | FST | 91308-12 | |
human FGF-basic recombinant protein (bFGF) | Gibco | 13256029 | |
Human TGF beta 1 recombinant protein (TGFb1) | eBiosciences | 14-8348-62 | |
Incubator – Heracell 160i CO2 | ThermoFisher | 51033557 | |
Inverted microscope – Revolve | ECHO | n/a | |
Liberase | Roche | 5401127001 | |
Mouse MesenCult Adipogenic Differentiation 10x Supplement | STEMCELL technologies | 5507 | |
Mouse on mouse (MOM) blocking reagent | Vector Laboratories | MKB-2213 | |
Paraformaldehyde | Sigma | P1648-500g | |
Penicillin-Streptomycin | Gibco | 15140–122 | |
PicoLab Mouse Diet 20 | LabDiet | 3005750-220 | |
Propidium iodide (PI) | Invitrogen | P3566 | |
Transport vial 5mL tube | Caplugs Evergreen | 222-3005-080 | |
Triton X-100 | Sigma | 9036-19-5 |