Detergent-Free Decellularization Preserves the Mechanical and Biological Integrity of Murine Tendon

Abstract

Tissue decellularization has demonstrated widespread applications across numerous organ systems for tissue engineering and regenerative medicine applications. Decellularized tissues are expected to retain structural and/or compositional features of the natural extracellular matrix (ECM), enabling investigation of biochemical factors and cell-ECM interactions that drive tissue homeostasis, healing, and disease. However, the dense collagenous tendon matrix has limited the efficacy of traditional decellularization strategies without the aid of harsh chemical detergents and/or physical agitation that disrupt tissue integrity and denature proteins involved in regulating cell behavior. In this study, we adapted and established the advantages of a detergent-free decellularization method that relies on latrunculin B actin destabilization, alternating hypertonic-hypotonic salt and water incubations, nuclease-assisted elimination of cellular material, and protease inhibitor supplementation under aseptic conditions. Our method maintained the collagen molecular structure (i.e., minimal extent of denaturation), while adequately removing cells and preserving bulk mechanical properties. Furthermore, we demonstrated that decellularized tendon ECM-derived coatings isolated from different mouse strains, injury states (i.e., naive and acutely injured/“provisional”), and anatomical sites harness distinct biochemical cues and robustly maintain tendon cell viability in vitro. Together, our work provides a simple and scalable decellularization method to facilitate mechanistic studies that will expand our fundamental understanding of tendon ECM and cell biology.

Impact statement

In this study, we present a decellularization method for tendon that does not rely on any detergent or physical processing techniques. We assessed the impact of detergent-free decellularization using tissue, cellular, and molecular level analyses and validated the preservation of gross fiber architecture, collagen molecular structure, and extracellular matrix (ECM)-associated biological cues that are essential for studying physiological cell-ECM interactions. Finally, we demonstrated the applicability of this method for healthy and injured tendon environments, across mouse strains, and for different types of tendons, illustrating the utility of this approach for isolating the contributions of biochemical cues within unique tendon ECM microenvironments.

Introduction

Tendinopathies, or tendon disorders, are debilitating conditions that account for over 30% of musculoskeletal consultations.¹ The tendon extracellular matrix (ECM) is primarily composed of type I collagen fibers that are sparsely populated by resident tendon cells.² Owing to their intrinsically limited healing capacity, ruptured tendons in adults typically heal by forming disorganized and mechanically impaired scar tissue (scar-mediated healing) that leads to long-term deficits in mobility, function, and quality of life.^3,4 A gold standard for the treatment of tendinopathy is surgical reconstruction of the ruptured tendon ends using a donor graft (e.g., xenograft or patient-derived autografts and allografts).

However, the success of these options has been restricted by challenges in the procurement of tendon grafts, postoperative retear rates as high as 94%,^5,6 and their limited capacity in attenuating further tissue degeneration.^2,4,7 Despite the clinical prevalence of tendinopathies and our growing understanding of the hallmarks governing their pathogenesis, the biological mechanisms that ultimately drive effective repair and functional restoration of the tendon remain elusive.^8,9 In particular, the role of local ECM microenvironmental cues in regulating tendon healing remains to be fully elucidated.

Tissue decellularization has emerged as a promising strategy to generate an acellular biologic scaffold to investigate the contributions of naturally derived ECM constituents and topographical features in guiding cell behavior.^10,11 In addition, advances in methods for processing decellularized ECM (dECM) have given rise to several in vitro and in vivo applications, including 2D dECM-coated culture substrates,^12,13 injectable dECM-based biomaterials,^14,15 and 3D dECM-encapsulated hydrogels and electrospun scaffolds^12,15–19 that retain tissue-specific and instructive cues that direct cell morphology, proliferation, and differentiation. However, the majority of tendon decellularization strategies employs harsh chemical detergents such as sodium dodecyl sulfate (SDS) and Triton X-100 and/or a combination of techniques that physically disrupt the ECM (e.g., repeated freeze-thaw cycles, ultrasonication, cryosectioning) that are known to negatively impact protein activity, tissue ultrastructure, and mechanical stability.^10,20–25

Hwang et al reported that even low concentrations of commonly used detergents can denature collagen molecules (i.e., irreversible unfolding of the collagen triple helix) in healthy porcine ligaments.²⁶ Moreover, White et al detected residual detergent fragments in decellularized tissues using time of flight secondary ion mass spectroscopy, which were shown to disrupt the ECM surface ligand landscape and consequently led to adverse effects on reseeded cell behavior in vitro.²⁷ Thus, there is mounting evidence in support of detergent-free decellularization approaches for tendon to more robustly recapitulate the native ECM biological environment.

In this study, we adapted and comprehensively characterized the effects of a detergent-free and nonproteolytic decellularization method that can be used to preserve the ECM integrity of mouse tendons across different genetic backgrounds, injury states, and anatomical sites. We hypothesized that tendons decellularized without detergents will be absent of any detectable molecular collagen denaturation or ECM destruction and therefore maintain their biological niche for cellular interrogation in vitro.

Methods

Animals and acute tendon injury model for tissue collection

All animal procedures were approved by the Cornell University Institutional Animal Care and Use Committee (IACUC). For primary cell isolations, male C57BL/6 (B6) mice at 16 to 18 weeks of age were bred in-house with the original dame and sire purchased from The Jackson Laboratory (No. 000664). For patellar tendon midsubstance punch surgeries, a subset of B6 mice was purchased directly from The Jackson Laboratory and acclimated for at least 1 week. To obtain a tendon biological environment distinct from that of B6 mice during injury, immunodeficient male NOD scid gamma (NSG) mice at 16 to 18 weeks of age were obtained from the Cornell University Progressive Assessment of Therapeutics (PATh) PDX Facility. Mice were housed with up to five animals per cage in an alternating 12-h light/12-h dark cycles with ab libitum access to chow and water.

To generate the injured/“provisional” tendon ECM, a 1-mm diameter, full-thickness biopsy punch (No. 33-31AA-P; Integra LifeSciences, Princeton, NJ) was introduced into the left patellar tendon midsubstance of B6 and NSG mice as described previously.^28,29 Briefly, mice were anesthetized with isoflurane (2% volume, 0.3 L/min) and received preoperative buprenorphine (1 mg/kg body weight, No. 078925235; Patterson Veterinary, Devens, MA) through subcutaneous administration. The fur at the surgical site was removed before making an incision directly above the knee cavity to expose the patellar tendon.

A polyurethane-coated (No. 1867T21; McMaster-Carr, Elmhurst, IL) stainless-steel backing was inserted underneath the tendon before using a biopsy punch to create the injury defect. The backing was removed before closing the skin incision using 6-0 prolene sutures (No. 8695G; Ethicon, Inc., Cincinnati, OH). Mice resumed cage activity. At 1 week postinjury, mice were euthanized by carbon dioxide (CO₂) inhalation followed by cervical dislocation. Hole-punched (provisional ECM) and uninjured contralateral tendons (naive ECM) were immediately harvested, flash-frozen in liquid nitrogen (LN₂), and stored at −80°C until decellularization. To demonstrate the utility of this decellularization method for other tendon tissues, uninjured B6 Achilles tendons were also collected.

For evaluation of bulk tensile mechanical properties, the patella-tendon-tibia complex was harvested from both limbs of uninjured B6 mice, carefully cleaned of the surrounding musculature, flash-frozen in LN₂, and stored at −80°C until decellularization.

Primary mouse tendon cell isolation and culture

Cells were routinely cultured in basal media consisting of low-glucose Dulbecco's modified Eagle's medium (DMEM; No. 10-014; Corning, Corning, NY) supplemented with 10% (v/v) lot-selected fetal bovine serum (FBS; No. 35-015-CV; Corning) and 1% (v/v) penicillin-streptomycin-amphotericin B (PSA; No. 15240062; Thermo Fisher Scientific, Waltham, MA). All experiments were conducted at passage 3 with media replenished every 2 days. All cells lines used in this study were negative for mycoplasma contamination (No. LT07-703; Lonza, Basel, Switzerland).

B6 patellar tendon cells were isolated from uninjured animals as described previously.²⁹ After euthanizing animals, the patellar tendon was harvested from both limbs from a total of five mice, stripped of the sheath and fat pad, and digested with a mixture of collagenase type I (2 mg/mL of digest solution; No. 17100017; Thermo Fisher Scientific) and collagenase type IV (1 mg/mL of digest solution; No. 17104019; Thermo Fisher Scientific) in serum-free DMEM for up to 2 h at 37°C on a rocking shaker. Single-cell suspensions were obtained by passing tissue digests through a 70-μm strainer (No. 352350; Corning) before first seeding cells onto a tissue culture-treated flask (passage 0).

Decellularization of tendon tissue and preparation of ECM-coated substrates

Naive and provisional mouse tendons were decellularized using an adapted detergent-free decellularization method^29,30 for skeletal muscle³¹ based on Latrunculin A (No. 2182-1; Biovision) and alternating hypertonic-hypotonic solutions. Incubation steps were performed with 1 mL of solution (2 mL for mechanical testing samples to ensure that the tissue was fully submerged) in 2.0-mL microcentrifuge tubes (No. MCT-150-C-S; Corning) with agitation at 450 rotations per minute (RPM) using an orbital shaker. All solutions with the exception of Latrunculin B were utilized at room temperature and supplemented with a 1 × protease inhibitor cocktail (No. 78425; Thermo Fisher Scientific) to minimize proteolysis. To limit the risk of contamination, tissue handling and solution changes were done in a biosafety cabinet using aseptic technique. Finally, salt solutions were prepared fresh in sterile deionized (DI) water for each decellularization batch. All 37°C and 65°C oven incubation steps were calibrated precisely using a standard mercury thermometer.

First, frozen tendons were thawed and individually incubated with 50 nM Latrunculin B (No. 2182-1; BioVision, Inc., Milpitas, CA) for 2 h at 37°C to disrupt actin polymerization. Second, tissue samples were then washed once with DI water for 30 min followed by 0.6 M potassium chloride (KCl; No. P3911; Sigma-Aldrich, St. Louis, MO) for 2 h, another DI water wash step for 30 min, and finally 1.0 M potassium iodide (KI; No. BDH9264; VWR International, Radnor, PA) for 2 h to induce osmolysis.

Third, samples were then washed in DI water for 12 h, subjected to the same alternating hypertonic-hypotonic solutions, incubated with 1 × phosphate-buffered saline (PBS; No. 21-040; Corning) supplemented with 1 kU/mL of Pierce Universal Nuclease (No. 88702; Thermo Fisher Scientific) for 2 h to remove cellular debris, and then finally washed with 1 × PBS for 48 h with the solution changed each day. All samples were stored at 20°C until use. All incubation steps were restricted to a tolerance of ±5%, but never any less than the allotted time.

For biochemical analyses, decellularized tendons were lyophilized for at least 72 h, individually weighed to obtain the dry mass, and then incubated with 200 μL of papain digest solution (0.2 mg/mL; No. P4762; Sigma-Aldrich) in 0.5-mL microcentrifuge tubes (No. AM12300; Thermo Fisher Scientific). Next, samples were vortexed to sediment the tissue to the bottom of the tube before digesting for 16 to 18 h at 65°C inside a tube rack on a rocking shaker until completely digested. Papain digests were stored at −20°C until analysis.

To make dECM-coated substrates, lyophilized tendons were first pooled up to 10 mg (dry weight) inside 2.0-mL low-binding tubes with screw caps and O-rings (No. 19-660-1000; Omni, Inc., Kennesaw, GA). Next, samples were mechanically homogenized (Bead Ruptor 24; Omni, Inc.) using five 2.8-mm ceramic beads (No. 19-646; Omni, Inc.) per tube for five pulverization cycles (6.0 m/s velocity for 15 s per cycle) with flash-freezing in LN₂ between each cycle.

Samples were then solubilized (10 mg of dry tissue/mL of solution) with pepsin (1 mg/mL; No. P7012; Sigma-Aldrich) in 0.1 M hydrochloric acid (HCl; No. H1758; Sigma-Aldrich) for exactly 72 h at 4°C at 150 RPM, diluted to a working concentration of 1 mg of dry tissue/mL solution in 0.1 M acetic acid (AcOH; No. 695092; Sigma-Aldrich), and stored at −80°C until use. Coatings were prepared by adding 50 μL of solubilized ECM to flat-bottom 96-well plates overnight (up to 18 h) at 4°C, which were subsequently washed twice with sterile 1 × PBS immediately before seeding cells.

Experiment

Biochemical analysis of dsDNA and sulfated glycosaminoglycan content

Biochemical analyses were performed on non-decellularized and decellularized samples using the papain digests. To determine the efficacy of the decellularization method in removing cellular materials, total double-stranded DNA (dsDNA) content was measured using a commercial Quant-iT PicoGreen kit (P11496; Thermo Fisher Scientific) as per the manufacturer's instructions. Fluorescence was measured at excitation/emission wavelengths of 480/520 on a SpectraMax i3X Multi-Mode Microplate Reader (Molecular Devices, San Jose, CA).

To determine the impact of detergent-free decellularization on the preservation of noncollagenous ECM constituents, sulfated glycosaminoglycan (sGAG) content was measured using the dimethylmethylene blue (DMMB; No. 341088; Sigma-Aldrich) assay.³² Standard curves were generated using chondroitin A (No. C9819; Sigma-Aldrich) for a range of sGAG concentrations. After the addition of the DMMB solution to the samples, absorbances were immediately measured at 540 and 595 nm to calculate the sample concentration by subtracting the absorbance at 595 nm from the absorbance at 540 nm, following a blank offset of DI water only.

Histology

Non-decellularized and decellularized B6 patellar tendons were embedded in optimal cutting temperature (No. 23-730-571; Fisher Scientific, Hampton, NH) compound and cryosectioned sagittally at a thickness of 6 μm. To determine the effect of decellularization on tissue structure and removal of cellular materials, cryosections were stained with hematoxylin and eosin (H&E) and 4′,6-diamidino-2-phenylindole (DAPI, No. D1306; Thermo Fisher Scientific), respectively. Images were acquired at 40 × magnification on an inverted brightfield microscope (H&E) or Axio Observer Z1 epifluorescence microscope (DAPI).

Fluorescent assessment of collagen denaturation with carboxyfluorescein-labeled collagen hybridizing peptide

To determine the extent of collagen denaturation after detergent-free decellularization, non-decellularized and decellularized cryosections were stained with 20 μM of carboxyfluorescein-labeled collagen hybridizing peptide^33–35 (CF-CHP; No. FLU300; Echelon Biosciences, Salt Lake City, UT) in 1 × PBS as per the manufacturer's instructions. Positive controls for CF-CHP staining were prepared by heating uninjured B6 patellar tendons in 1 × PBS at 65°C for 1 min and cryosectioning as described above. Trimeric CF-CHP solution was heated at 80°C for 5 min to obtain thermally dissociated monomeric strands. Monomeric CF-CHP solution was incubated at −80°C for 15 s and then quenched to room temperature. Cryosections were stained with 25 μL of CF-CHP, incubated at 4°C for 12 h in a humidified chamber, and carefully washed thrice in 1 × PBS for 5 min per wash to remove unbound CF-CHP.

Finally, stained cryosections were dehydrated using a graded alcohol series and mounted in EUKITT^® medium (No. 15322-10; Electron Microscopy Sciences, Hatfield, PA). Images were acquired on a Zeiss LSM 710 confocal microscope (Zeiss) at 10 × magnification and 488 nm laser excitation. Identical imaging parameters were used for all samples. Integrated density measurements of CF-CHP staining indicative of denatured collagen were quantified in ImageJ by a blinded user using a custom MATLAB script to exclude sectioning artifacts from analysis. A total of two to four stained sections (spaced at least 100–150 μm apart from each other) taken throughout the full thickness of the tendon were averaged for each sample to account for any variation in the penetration of decellularization reagents.

Bulk tensile testing

Non-decellularized and decellularized B6 patella-tendon-tibia complexes were secured using custom grips in a 1 × PBS bath at room temperature. A nominal 0.15 N preload was applied and then followed by preconditioning for 15 cycles at 1% strain and a frequency of 1 Hz. Stress relaxation was assessed at 5% strain (5% strain/s) with a recovery time of 300 s. After stress relaxation testing, tendons were held for an additional 300 s in the absence of any applied load. Pre-onditioning was then reapplied as described above before pulling samples uniaxially to failure at a rate of 0.1% strain/s to record the ultimate load and stiffness. Tendon cross-sectional area was calculated from digital images taken using a monochrome industrial camera (DMK 33UX250) during the 0.15 N preload before mechanical testing.

Assessment of cytocompatibility using live-dead cell staining

To assess the cytocompatibility of dECM-derived biochemical cues, B6 patellar tendon cells were seeded (7500 cells/well) in basal media onto B6-naive or provisional ECM-coated 96-well glass-bottom plates (No. 655892; Greiner Bio-One, Monroe, NC). Noncoated wells served as controls. After adhering for 4 h, cells were rinsed once with plain DMEM and then cultured in FluoroBrite DMEM (No. A1896701; Thermo Fisher Scientific) supplemented with 1% lot-selected FBS and 1% PSA for 3 days.

Live/dead cell staining was performed by incubating cells with 5 μM calcein AM (No. C3100MP; Thermo Fisher Scientific) and 3 μM propidium iodide (No. P1304MP; Invitrogen, Waltham, MA) for 30 min. Images were then immediately acquired on a Zeiss Axio Observer Z1 epifluorescence microscope at 2.5 × magnification. Identical imaging parameters were used for all samples. Cell viability was calculated as the percentage of calcein-positive cells divided by the total number of cells. Three technical replicates were averaged for each biological replicate.

Analysis of single-cell morphology in vitro

To functionally evaluate if the altered provisional ECM composition of NSG tendons due to their suppressed immune response is preserved with decellularization, B6 patellar tendon cells were seeded (2000 cells/well) in basal media onto NSG-naive or provisional ECM-coated 96-well glass-bottom plates for 18 h. Cells were then fixed with 4% paraformaldehyde (No. 15170; Electron Microscopy Sciences) for 30 min, permeabilized with 0.1% Triton X-100 for 15 min, blocked with 2.5% normal horse serum (No. S-2012; Vector Laboratories, Newark, CA) for 30 min, and concurrently stained with Alexa Fluor 488 Phalloidin (1:400 dilution; No. A12379; Thermo Fisher Scientific) and DAPI (1:1000 dilution) in 2.5% normal horse serum for 1 h, while covered from light.

All immunostaining steps were performed at room temperature. Cells were then rinsed twice and mounted with 1 × PBS before images were acquired on a Zeiss Axio Observer Z1 epifluorescence microscope at 20 × magnification. Single-cell morphological parameters, including cell spreading area, cell perimeter, and circularity (calculated as 4 × π × cell spreading area/perimeter²) were quantified in ImageJ by a blinded user.

Statistical analysis

Biochemical and mechanical analyses for non-decellularized and decellularized samples were compared using an unpaired Student's t-test. CF-CHP-integrated density and cell viability between uncoated control, naive, and provisional ECM-coated substrates were compared using a one-way analysis of variance with post hoc Tukey. Single-cell morphology between naive and provisional ECM-coated substrates was compared using an unpaired two-tailed Welch's t-test to account for unequal variance between our population means. All experiments were conducted with a minimum of three biological replicates with each biological replicate representing a unique tendon sample or cell line isolated from a different batch of mice.

Experimental Results

Detergent-free decellularization preserved tissue structure and removed cells

Supporting our hypothesis, qualitative histological examination of H&E-stained naive B6 patellar tendons revealed the preservation of the general collagen fiber architecture characteristic of native tendon structure following decellularization (Fig. 1A; top panel). Both H&E and DAPI staining (Fig. 1A) illustrated the removal of resident cells from naive tendons after decellularization. Corroborating these visual observations, dsDNA content in B6-naive and provisional tendons was significantly reduced from 1039.14 ± 184.1 and 2957.37 ± 298.1 ng/mg, respectively, to 47.64 ± 5.36 (p < 0.0001) ng/mg and 184.6 ± 24.32 ng/mg (p < 0.0001) after decellularization (Fig. 1B).

FIG. 1.

Histological and biochemical evaluation of non-decellularized and decellularized B6 patellar tendons. (A) Representative H&E (top panel; scale bar, 100 μm) and DAPI staining (bottom panel; scale bar, 25 μm) demonstrated preservation of collagen fiber alignment, while removing cellular material. White arrows indicate cells. N = 6 samples per group. (B) Total dsDNA and (C) sGAG content in B6-naive and provisional tendons were reduced before and after decellularization as measured by the Quant-iT PicoGreen and DMMB assays, respectively. N = 7–8 samples per group. Dashed black line indicates 50 ng/mg threshold for dsDNA content. Data are presented as mean ± standard deviation. **p < 0.01, ***p < 0.001, ****p < 0.0001. DAPI, 4′,6-diamidino-2-phenylindole; DMMB, dimethylmethylene blue; dsDNA, double-stranded DNA; H&E, hematoxylin and eosin; sGAG, sulfated glycosaminoglycan. Color images are available online.

Similarly, dsDNA content in B6-naive Achilles tendons was significantly reduced from 425.50 ± 5.17 to 26.61 ± 0.55 ng/mg after decellularization (data not shown), establishing the efficacy of this decellularization method for different tendon tissues. As expected, the effect of decellularization on reducing overall dsDNA content was significantly greater (p = 0.0041) in naive (95.42% ± 0.52% reduction) compared to provisional tendons (93.76% ± 0.82% reduction), which is presumably attributed to the nearly threefold increase in cellularity and formation of dense granular tissue (i.e., reduced penetration of decellularization reagents) during acute tendon injury.

The sGAG content in B6-naive and provisional tendons was reduced from 5.29 ± 1.08 and 13.53 ± 0.79 μg/mg, respectively, to 3.00 ± 0.48 μg/mg (p = 0.0002) and 12.17 ± 1.66 μg/mg (p = 0.075) after decellularization (Fig. 1C). Interestingly, the overall reduction in sGAG content was significantly lower (p < 0.0001) in provisional (10.01% ± 12.25%) compared to naive (43.38% ± 9.03%). These results indicate that the sGAG-rich provisional ECM composition produced during the early proliferative phase of wound healing^36,37 is largely maintained following detergent-free decellularization.

Detergent-free decellularized tendons showed no sign of collagen denaturation

Collagen denaturation was not visually detected in non-decellularized and decellularized B6-naive patellar tendons as evidenced by a lack of CF-CHP staining compared to the heat-denatured control group (Fig. 2A). Furthermore, there was no difference in CF-CHP staining intensity between superficial and deeper cryosections of the tendon. Supporting our hypothesis, CF-CHP-integrated density of non-decellularized and decellularized samples was comparable and were both significantly lower (p < 0.0001 for both) compared with the heat-denatured control group (Fig. 2B), confirming that detergent-free decellularization combined with protease inhibitors limited proteolytic degradation of the tendon ECM environment.

FIG. 2.

Molecular assessment of collagen denaturation using CF-CHP staining. (A) Representative images of CF-CHP (green indicative of denatured collagen)-stained cryosections from B6-naive patellar tendon for non-decellularized, decellularized, and heat-denatured conditions. Dashed white lines and saturated images (bottom panel) indicate the tissue outline/representative tissue section, respectively. Scale bar, 200 μm. N = 6 samples per group. (B) Quantification of CF-CHP integrated density illustrated minimal collagen denaturation in non-decellularized and decellularized tendons. Data are presented as mean ± standard deviation. ****p < 0.0001. CF-CHP, carboxyfluorescein-labeled collagen hybridizing peptide. Color images are available online.

Bulk tissue mechanical properties were not affected by detergent-free decellularization

All tested samples failed at the tendon midsubstance. Supporting our hypothesis, there was no difference in ultimate load, stiffness, or stress relaxation between non-decellularized and decellularized B6-naive patellar tendons (Supplementary Fig. S1 and Fig. 3A–C). Taken together with our histological assessment, our visual observations of minimally disrupted collagen organization with decellularization are supported by these unchanged tissue-level mechanical properties. Surprisingly, while sGAGs are known to be hydrophilic macromolecules that bind water and therefore impart the viscoelastic behavior of tendon,^38,39 the percentage stress relaxation remain unchanged, despite a 43.3% reduction in sGAG content.

FIG. 3.

Bulk tensile testing of non-decellularized and decellularized B6-naive patellar tendons. (A) Ultimate load, (B) stiffness, and (C) stress relaxation of B6-naive patellar tendons were not affected by detergent-free decellularization. N = 6–8 per group. Data are presented as mean ± standard deviation.

Tendon ECM-derived coatings are cytocompatible and promote cell survival

After 3 days under 1% FBS/serum-deprived conditions, B6 patellar tendon cells cultured on B6-naive (p = 0.028) and provisional (p = 0.045) ECM-coated substrates exhibited significantly greater viability compared to the uncoated control (Fig. 4A, B). There was no difference in the total cell number between each experimental group (data not shown), indicating that tissue-specific biochemical cues harnessed by the decelluarized tendon ECM environment promote cell survival.

FIG. 4.

Cytocompatibility assessment of tendon cells on decellularized B6-naive and provisional ECM-coated substrates. (A) Representative images of live-dead (green indicates live and magenta indicates dead) cell staining of B6 patellar tendon cells cultured on uncoated, naive ECM-coated, and provisional ECM-coating substrates for 3 days under serum-deprived conditions. Scale bar, 200 μm. (B) Quantification of live-dead cell staining showed significantly increased cell viability on tendon ECM-coated substrates compared to the uncoated control. N = 3 per group. Data are presented as mean ± standard deviation. *p < 0.05. ECM, extracellular matrix. Color images are available online.

Single-cell morphology assessment in vitro

To further validate the utility of our detergent-free decellularization method in preserving the unique tendon ECM composition of other mouse strains, we analyzed the morphology of B6 patellar tendon cells cultured on naive and provisional ECM-coated substrates derived from tendons of immunodeficient NSG mice (Fig. 5A).^40,41 As expected, B6 tendon cells showed significantly lower spreading area (p = 0.017) on NSG provisional ECM-coated substrates (3879.24 ± 2134.61 μm²) compared to naive ECM-coated substrates (5052.16 ± 3532.0569 μm²) (Fig. 5B). There was no difference in cell perimeter or circularity for B6 tendon cells cultured on naive and provisional ECM-coated substrates. These data indicate the capacity of this detergent-free decellularization approach to evaluate how altered tendon ECM-associated compositions, such as with diminished proinflammatory cytokine signaling during NSG tendon healing, differentially modulate tendon cell behavior.

FIG. 5.

Single-cell morphological analysis of B6 patellar tendon cells cultured on NSG-naive and provisional ECM-coated substrates. (A) Representative cytoskeletal outlines of 10 representative cells cultured on NSG-naive and provisional ECM-coated substrates. Scale bar, 200 μm. (B) Quantification of single-cell morphology revealed that B6 tendon cells exhibited significantly lower spreading area when cultured on NSG provisional ECM-coated substrates compared to naive ECM-coated substrates. N = 72 cells per group from three independent experiments (each biological replicate represented by a different color). Data are presented as mean ± standard deviation. *p < 0.05. NSG, NOD scid gamma. Color images are available online.

Discussion

To date, the primary challenge in maximizing the potential of decellularized tissues and organs for clinical and mechanistic studies has been balancing the removal of cellular materials (i.e., as to reduce immunogenicity) and adequately preserving the complex structural and biochemical milieu of the ECM environment. The negative consequences of traditional detergent-based decellularization methods such as SDS and Triton X-100 on tendon structure and composition have been well established.

Mechanical overload of rat tail tendon fascicles has also been shown to lead to unfolding of collagen molecules as detected by CF-CHP staining,^42,43 suggesting that irreversible ECM damage achieved by either mechanical or proteolytic mechanisms may detrimentally alter cell-ECM interactions. Indeed, Veres et al reported decreased accumulation of macrophage-like cells cultured in vitro at sites of mechanically induced plastic deformation in decellularized collagen fibrils and an activated phenotype associated with increased cell spreading and upregulated catabolic activity.⁴⁴ Collectively, there is mounting evidence that tissues decellularized with detergents pose a risk of compromising biological functionality and stimulating an aberrant cellular response.

In this study, we successfully developed a detergent-free decellularization method for murine tendons that circumvents these concerns associated with traditional approaches. A limitation of this study is that the structural assessment of decellularized tendons was limited to qualitative visual observations using H&E staining. Bulk tissue mechanics remained unchanged following decellularization, which suggests that the hierarchical tendon ultrastructure was preserved, but quantitative analyses (e.g., Picrosirius red staining and/or second-harmonic generation imaging) are necessary to definitively ascertain any effect on collagen fiber alignment.

Nevertheless, we have previously applied our decellularization method to investigate the cellular and molecular processes underlying regenerative tendon healing by isolating provisional tendon ECM^29,30 from the super-healer Murphy Roths Large (MRL/MpJ) mouse strain.^29,45–48 Complementing our minimal collagen denaturation data in this study, we have previously shown that innate differences in the ECM-sequestered growth factor signaling of B6- and MRL/MpJ-naive and provisional tendons are retained after decellularization.²⁹ We also previously demonstrated that B6 tendon cells cultured on decellularized MRL/MpJ dECM-coated substrates exhibited enhanced cell proliferation, elongation, and the formation of cellular protrusions that are characteristic of MRL/MpJ tendon cell behavior.²⁹

Although the decellularized B6 provisional ECM exceeded the 50 ng dsDNA/per mg of dry weight threshold proposed by Crapo et al,¹⁰ we have previously implanted MRL/MpJ dECM-derived hydrogels into injured B6 patellar tendons in vivo and improved the tissue structure and mechanical properties without observing a deleterious inflammatory reaction indicative of an adverse immunogenic response.³⁰ Excitingly, these findings validate the versatility of our detergent-free decellularization method and its untapped capacity to mechanistically interrogate tendon ECM biology using other transgenic mouse strains, sex-based comparisons, enzyme-mediated ECM depletion (e.g., elastase for elastin or plasmin for fibrin), and other models of tendon injury such as chronic overuse through strenuous exercise or fatigue loading.⁴⁹

This study determined that our detergent-free decellularization method is capable of preserving ECM-derived biological cues across different tendon environments for cellular assessment under 2D culture conditions. Our recent work has shown that B6 and MRL/MpJ tendon cells cultured on commercial 3D nanofiber scaffolds elicit distinct cellular responses to anisotropic and isotropic surface topographies associated with homeostasis and wound healing, respectively.⁵⁰ To gain further insight into how the interplay between the biochemical and structural properties of the ECM regulate cell activity, future studies could augment these biomaterial-based approaches by applying the dECM as a surface coating.

Future experiments should also probe cell behavior under 3D culture systems using established methods for fabricating dECM-derived hydrogels⁵¹ or electrospun scaffolds¹⁹ that better recapitulate the complexities of the physiological environment. Finally, the methods presented in this study open up the avenue for future studies to explore whole-tissue recellularization strategies through surgical repair (e.g., anterior cruciate ligament reconstruction), in vivo transplantation,⁵² or ex vivo bioreactor and organ culture systems to assess the crosstalk between ECM and mechanical regulation.

Footnotes

Acknowledgment

The authors thank Claudia Fischbach-Teschl for providing training and technical assistance with their epifluorescence microscope.

Authors' Contributions

J.C.M.: conceptualization (Lead), data collection and formal analysis (Lead), writing—original draft (Lead), and review and editing (Equal). A.M.: data collection and formal analysis (Supporting), software (Lead), writing—original draft (Supporting), and review and editing (Supporting). J.P.: data collection and formal analysis (Supporting), writing—original draft (Supporting), and review and editing (Supporting). B.V.: data collection and formal analysis (Supporting), writing—original draft (Supporting), and review and editing (Supporting). N.A.-P.: conceptualization (Supporting), writing—original draft (Supporting), and review and editing (Equal).

Disclosure Statement

No competing financial interests exist.

Funding Information

This work was supported by the National Institutes of Health (NIH) under the following award numbers: R01AR608301 (to N.A.-P.), R01AR052743 (to N.A.-P.), and S10RR025502 (to Cornell Biotechnology Resource Center). The authors also acknowledge support from the National Science Foundation (NSF) Graduate Research Fellowship Program (GRFP) DGE-1650441 (to J.C.M.).

Supplementary Material

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