Porcine Lung-Derived Extracellular Matrix Hydrogel Properties Are Dependent on Pepsin Digestion Time

Abstract

Hydrogels derived from decellularized lungs are promising materials for tissue engineering in the development of clinical therapies and for modeling the lung extracellular matrix (ECM) in vitro. Characterizing and controlling the resulting physical, biochemical, mechanical, and biologic properties of decellularized ECM (dECM) after enzymatic solubilization and gelation are thus of key interest. As the role of enzymatic pepsin digestion in effecting these properties has been understudied, we investigated the digestion time-dependency on key parameters of the resulting ECM hydrogel. Using resolubilized, homogenized decellularized pig lung dECM as a model system, significant time-dependent changes in protein concentration, turbidity, and gelation potential were found to occur between the 4 and 24 h digestion time points, and plateauing with longer digestion times. These results correlated with qualitative scanning electron microscopy images and quantitative analysis of hydrogel interconnectivity and average fiber diameter. Interestingly, the time-dependent changes in the storage modulus tracked with the hydrogel interconnectivity results, while the Young's modulus values were more closely related to average fiber size at each time point. The structural and biochemical alterations correlated with significant changes in metabolic activity of several representative lung cells seeded onto the hydrogels with progressive decreases in cell viability and alterations in morphology observed in cells cultured on hydrogels produced with dECM digested for >12 and up to 72 h of digestion. These studies demonstrate that 12 h pepsin digest of pig lung dECM provides an optimal balance between desirable physical ECM hydrogel properties and effects on lung cell behaviors.

Impact statement

Extracellular matrix (ECM) hydrogels, which offer unique material advantages, have been developed to model the ECM in vitro, to improve tissue-engineered and biomanufactured scaffolds, and directly adapted as clinical therapies in vivo. Standardized enzymatic digestion protocols have supported the rapid adoption of ECM hydrogels, however, contribution of specific digestion parameters has been overlooked. Using pig lung decellularized ECM as a model system, this research demonstrates digestion time-dependent effects on the physical, mechanical, and biological properties of the resulting hydrogels. Importantly, this highlights the possibility that similar evaluations could yield significant and impactful performance benefits if applied for individual tissues and applications.

Introduction

Treatments for lung diseases such as chronic obstructive pulmonary disease, idiopathic pulmonary fibrosis, and cystic fibrosis are limited, and lung transplantation is often the only option for patients in later stages. As such, there is a critical need for new treatment strategies and for improved models for studying lung disease mechanisms. One promising approach, increasingly adopted for respiratory bioengineering, is the application of tissue-derived scaffolds prepared from native lung tissue using decellularization techniques.

Acellular lung scaffolds, initially developed for whole-organ engineering^1–6 and subsequently adapted to study lung biology in vitro,^7–10 retain much of complexity of the native extracellular matrix (ECM) composition and bioactivity. As a model of the lung microenvironment, acellular lung scaffolds have been shown to induce relevant effects on cells seeded in vitro, including the formation of organized structures, directed differentiation of lung progenitors, and differential response of cells seeded into scaffolds derived from diseased lungs.^2,6,11,12 As a result, there is increasing investigation and use of decellularized scaffolds for isolating tissue-specific ECM (decellularized ECM [dECM]) to generate new materials for tissue engineering, as clinical therapies, and to improve models for studying the ECM in vitro.^13–17

To isolate tissue-specific dECM, lung or otherwise, decellularized tissues are lyophilized, milled, and then stored as a microparticle dECM powder.¹⁸ As a stand-alone material, dECM powder has been directly used in tissue engineering applications,^19,20 as a therapeutic agent,²¹ and for cell delivery in vivo.^19,22 However, further processing is required to prepare dECM for most biomaterial formats. One strategy, in which dECM powder is suspended in chemical solvents, has been used to prepare lung dECM for generation of electrosprayed nanoparticles,²³ electrospun airway scaffolds,²⁴ and thin ECM “tissue papers”.²⁵ However, for other applications, including formation of hydrogels, the dECM structure must be fully dissembled to allow soluble ECM to reassemble as a gel.

The most common strategy used to solubilize lyophilized lung dECM is enzymatic pepsin digestion in acidic buffer. In this process, dECM powder is solubilized into a monomeric suspension at room temperature (RT) and subsequently pH neutralized to prepare a thermosensitive pregel solution which polymerizes into an ECM hydrogel when incubated at 37°C.^26–32 Previously, we had developed and characterized the first lung-derived ECM hydrogel, using 48 h enzymatic digestion, and demonstrated its capacity to support encapsulated cells as a potential in vivo therapeutic delivery vehicle.^26,27 More recently, lung ECM hydrogels have been further developed as a model to study ECM derived from healthy and diseased lungs,^31,32 as a bioink for three-dimensional (3D) additive manufacturing processes^33,34 and as a protective therapy for radiation-induced lung injury in vivo.³⁰

This rapid adoption highlights the potential for lung ECM hydrogels to make an impact in several areas. However, there are challenges to address. Notably, most ECM hydrogels are produced using a general strategy, in which the decellularized tissue is lyophilized, milled, and then enzymatically digested, most frequently with pepsin, in an acidic medium for 24–72 h to achieve solubility.^35,36 Under these standardized protocols tissues successfully used to prepare ECM hydrogels in addition to lung include small-intestinal submucosa,³⁷ urinary bladder,^35,38–40 adipose,⁴¹ heart,^42–46 cornea,⁴⁷ dermis,⁴⁸ liver,^49–52 esophagus,⁵³ demineralized bone,⁵⁴ cartilage,^55–57 central nervous system,⁵⁸ umbilical cord,⁵⁹ and muscle.⁶⁰

One limitation is that ECM hydrogels prepared by enzymatic digestion are not as mechanically robust as their source tissue, a key factor in directing biologically relevant cell responses.^61,62 Previous studies addressing this limitation have demonstrated the ability to control the mechanical properties of ECM hydrogels by (1) manipulating protein concentration, salt buffer composition, and/or pH of the pregel solution,^27,38,48,63 (2) treating the resulting hydrogels with chemical crosslinkers,^{27,42,64–67} and (3) addition of synthetic functional groups or polymers to the solublized dECM.^68–71 While each of the strategies described can be implemented to tune the properties of ECM hydrogels, they all rely on postdigestion sample manipulation to achieve this success.

Interestingly, this highlights the role of the enzymatic pepsin digestion on ECM hydrogel properties as an area that has previously been understudied. The enzymatic activity of pepsin is determined by the pH and temperature of the digestion buffer, with peak activity at 37°C and pH ∼2.0.^72–74 Also, the rate and progress of the digestion are dependent on process control parameters, including agitation speed, pepsin-to-substrate ratio, and digestion time.⁷⁵ Pepsin exhibits cleavage-site specificity, for example, the enzyme selectively removes exposed telopeptides of collagen 1 at a faster rate than it disrupts the triple helix structure with nonspecific cleavages.^76,77 This is important, as collagen 1 is a significant component of lung-derived dECM.

Studies investigating the preparation of polymerizable pepsin-soluble collagen 1, have demonstrated that digestion time and temperature effect the gelation behavior, physical structure, and mechanical properties of the resulting collagen 1 hydrogels.⁷⁸ Notably, studies investigating the effect of pepsin digestion on decellularized cardiac scaffolds found that the composition of the soluble ECM fraction was digestion time-dependent and that cardiomyocytes proliferated faster on tissue culture plastic coated with ECM digested for shorter times.⁷⁹ However, there is no comparable available information with respect to the effects of pepsin digestion time on subsequent properties of decellularized lung hydrogels. In this context, the objective of this study was to investigate the effect of pepsin digestion time on the physical, mechanical, and bioactive properties of ECM hydrogels, using decellularized pig lungs as a model system.

Materials and Methods

Materials preparation

Porcine lung tissue decellularization and processing

Lungs from domestic pigs (Sus scrofa), 4–6 months old, were obtained from local slaughterhouses. Pig lungs were decellularized in a dedicated sterile biosafety cabinet according to a previously established protocol.²⁶ In brief, porcine lungs were rinsed externally with 1 × phosphate-buffered saline (PBS) to remove excess blood, and the right lung was carefully dissected to preserve the main bronchus and pulmonary vascular portals. The lung was then sequentially treated with and incubated in a series of reagents, including 0.1% v/v Triton X-100 (24 h, 4°C), 2% w/v sodium deoxycholate (24 h, 4°C), 1 M NaCl (1 h, RT), and DNase (1 h, RT). Between each treatment step, the lung was washed with ultrapure DI H₂O (MilliQ) to remove the previous reagent and cellular debris.

For treatment and rinse steps, the solutions were perfused through the airway and then vasculature, three times each. After the last step, the lung was washed five times with sterile PBS to remove any remaining detergents or debris. Following decellularization, the large cartilaginous airways were dissected and discarded, and the remaining tissue was sectioned into smaller pieces (∼1–2 cm³), loosely packed in sterile 50 mL tubes, frozen at −80°C, and then dried using a Cryozone 2.5 L lyophilizer (Labconco). The dried acellular tissue was milled using a liquid nitrogen freezer mill (SPEX 6700) for 5 min at the highest speed to produce a fine particle dECM powder. Dry milled dECM powder was stored at −80°C until use.

Enzymatic digestion of pig lung decellularized extracellular matrix

Pig lung dECM was enzymatically digested as previously described.^26,27 In brief, milled dECM powder was suspended at 10 mg/mL in a 0.01 M HCl solution containing 1 mg/mL pepsin from porcine gastric mucosa (3450 U/mg, P68875G; Sigma). Enzymatic digestion was performed at RT under constant stir bar agitation, and samples were collected at 4, 12, 24, 48, and 72 h for the following characterization experiments. The digestion solution isolated at each time point was prepared as a total ECM sample, while a smaller volume was used to prepare a fractionated sample. In this case, the digest solution was centrifuged at 20,000 g for 5 min to pellet insoluble protein; the supernatant collected is the soluble ECM sample. During sample collection, the pH of the digest buffer was monitored using low-range test strips (pH range: 1.0–6.0; Hydrion Cat # 152), and values were recorded for each time point.

Preparation of pig lung-derived extracellular matrix hydrogels

To prepare the thermosensitive ECM pregel solutions for hydrogel experiments, the total and soluble ECM samples were treated to balance pH and salt ions using a standard neutralizing solution of 1 M NaOH (FisherSci) and 10 × PBS (Gibco) at a 9:10 ratio. Before neutralization, both the digestion sample and neutralization solution were chilled on ice for 5 min. The neutralization was conducted by adding 23% v/v of neutralization solution to the sample and mixing gently, but thoroughly, to avoid creating bubbles. The pH of the pregel solutions was monitored using midrange test strips at each time point (pH range: 6.0–8.0; Hydrion Cat # 345), to confirm that the sample was in the desired range ∼7.4 ± 0.2. The pregel solutions, now at ∼8 mg/mL following neutralization, can be used immediately or stored at 4°C for up to 1 week. To prepare an ECM hydrogel, the desired volume of pregel solution is transferred to a vessel or tissue culture plate and then incubated at 37°C for 30 min to induce gelation.

Tracking matrix solubilization as a function of digestion time

Soluble and total ECM sample preparation and total protein assays

To track ECM digestion and protein concentration of total and soluble ECM, a bicinchoninic acid Protein quantification assay (Pierce BCA Protein Assay Kit; ThermoFisher) was conducted according to the manufacturer's instructions. To prepare the ECM samples for this assay, total and soluble pregel fractions were diluted 1:4 in protein lysis buffer (2% sodium dodecyl sulfate [SDS], 62.5 mM tris, pH 7.4). The samples were transferred to a 96-well flat bottom plate (Greiner Bio-One) in triplicate and exposed to assay reagents for 30 min at 37°C. Absorbance was quantified at 562 nm using a plate reader, and a bovine serum albumin standard was used to determine protein concentration.

Tracking solubility and gelation potential using a turbidity assay

A turbidity assay was also conducted using the total and soluble ECM samples to examine the digestion progress and gelation potential as a function of digestion time. For this experiment, 50 μL of each pregel sample was transferred, in triplicate, to a 96-well round bottom plate (Greiner Bio-One), while both the sample and plate were maintained at 4°C. Using a plate reader, absorbance of each well was recorded at 405 nm as a measure of each pregel solution's turbidity. Following this initial measurement, the same samples were incubated at 37°C for at least 30 min to induce gelation and absorbance at 405 nm was measured again. Gelation potential was calculated at each digestion time point, as the percent change in absorbance resulting from gelation. The change in turbidity in these samples is a direct result of soluble ECM polymerization into insoluble hydrogels network architecture, and therefore an indirect assessment of the potential of the soluble protein to form a gel.

To further assess the effect of digestion time on ECM hydrogels, a turbidimetric gelation kinetics characterization experiment was conducted using a previously published strategy.^35,58 In brief, a second set of timed digest samples were prepared and pregel samples were again transferred to a 96-well round bottom plate, mirroring the first turbidity experiment. Before reading the initial turbidity values at 405 nm, the environmental temperature of the plate reader was preset to 37°C. Then the plate was loaded into the reader, an initial t = 0 turbidity measurement was taken, and then additional measurements were made every 3 min, for 45 min. The resulting turbidimetric gelation data were used to calculate normalized absorbance (Eq. (1)] to generate gelation kinetics curves. $N A = (A - A_{0}) ∕ (A_{m a x} - A_{0}) (1)$ (1)

Using these curves several parameters were calculated allowing us to compare the effect of digestion time on the gelation behavior of the ECM hydrogels. This includes the time parameters: t_1/2 (time to reach 50% maximum turbidity), t_95% (time to reach 95% maximum turbidity), and t_lag (time to start gelation), as well as S (speed of gelation) defined as the maximum slope at t_1/2, which is assumed to be during the linear growth phase of gelation.

Tracking protein size distribution using SDS-polyacrylamide gel electrophoresis

Soluble and total ECM fractions were diluted 1:4 in protein lysis buffer (2% SDS, 62.5 mM tris, pH 7.4). Protein concentration values were used to balance each sample to load 7.25 μg of protein per lane. Samples were then fluorescently labeled with Cy5 fluorescent probe using the Amersham Quickstain kit according to the manufacturer's instructions (GE Healthcare). The labeling reaction was stopped with a 1:1 dilution of each sample with 2 × lithium dodecyl sulfate buffer containing 1 × reducing agent (ThermoFisherScience). Samples were then heated at 70°C for 10 min, cooled to RT, and centrifuged (10,000 g for 10 min). Twenty microliter (7.25 μg protein) of each sample was loaded per lane on a NuPAGE tris-acetate gradient (ThermoFisherScience; 3–7%, 12-well, 1.0 mm), and every sample set of five-time points was run next to a HiMark protein standard. Electrophoresis was conducted at 150 V for ∼55 min using a bolt mini gel box (ThermoFisherScience).

Following electrophoresis, the tris acetate gels were imaged using an Amersham 600 imager (fluorescent Cy5 channel) and the images analyzed using Bio-Rad Image Lab software. Semiquantitative analysis of the images was conducted to examine the size distribution of larger and smaller proteins as a function of digestion time. To do this a digital molecular weight cutoff was generated at 250 kDa using the HiMark standard and the signal from all individual bands above and below the cutoff summed and normalized by total signal from all the bands in a given lane.

Tracking total protein and collagen 1 digestion using fluorescent western blotting

After the SDS-polyacrylamide gel electrophoresis (PAGE) gels were imaged, they were equilibrated in NuPAGE transfer buffer (1 × NuPAGE Transfer Buffer, 10% MeOH, 0.05% SDS) and transferred (Bio-Rad Mini Trans-Blot Cell) to a low-fluorescence polyvinylidene difluoride transfer membrane (Thermofisher; 0.2 μM). Following transfer, (4°C, 340 mA, 90 min), the membranes were blocked with 5% ECL-Prime blocking agent (GE Lifesciences), dissolved in 0.01% TBS-t₂₀, for 1 h at RT, rinsed three times in 0.01% TBS-t₂₀, and then imaged to confirm efficient protein transfer (Cy5 channel). The membranes were then incubated for 1 h at RT using a collagen 1 rabbit polyclonal primary antibody (Abcam; ab34710) diluted 1:2000 in the original blocking buffer and then rinsed three times in wash buffer.

The membrane was then probed using a Cy2 conjugated goat-anti-rabbit fluorescent secondary (ThermoFisherScience; A32731) for 1 h at RT and then rinsed three times in wash buffer. The membrane was imaged using the Amersham 600 imager (fluorescent Cy2 and Cy5 channel). Semiquantitative collagen 1 analysis was broken up to assess the relative change in trimeric (α1 α1 α2, ∼400 kDa), dimeric (α1 α1 or α1 α2, ∼270/260 kDa), and monomeric (α1 or α2, ∼138/128 kDa) collagen using the HiMark standard to establish digital molecular weight cutoffs. Western blots were evaluated using Bio-Rad Image Lab software. For each individual sample lane, the trimeric, dimeric, and monomeric collagen 1 signal was normalized by total collagen I signal.

Physical characterization of extracellular matrix hydrogels

SEM imaging, interconnectivity, and fiber diameter analysis

Hydrogels prepared from dECM digested for 4, 12, 24, 48, or 72 h were fixed in 5% glutaraldehyde and 4% osmium tetroxide solution for 1 h at RT and then dehydrated in a series of graded ethanol solutions. The samples were then dried using a critical point dryer (Tousimis) and plasma sputter coated for SEM imaging. Three representative images were taken from each sample at 10,000 × magnification on either a JEOL JSM-5610LV SEM or a Hitachi SU-70 FE SEM. Image analyses were conducted using ImageJ to quantify the interconnectivity of the self-assembled ECM hydrogel network and to measure the average diameter of the self-assembled fibers are the resulting hydrogels at each digestion time point.

This approach was adapted from established protocols to measure the fiber diameter of porcine lung hydrogels.^63,80 In brief, using ImageJ a 5 μm² grid was applied to each SEM image and the perpendicular cross section of 25 fibers was measured in three of the grid segments per image. Average fiber diameter was calculated using measurements taken from three separately prepared hydrogels at each time point. To calculate interconnectivity, the total number of branched or entangled ECM fibers in each grid segment section was recorded. The total number of interconnections within the 12 segments was divided by the combined area to calculate network interconnectivity as [interconnections/μm²].

Evaluating the mechanical properties of ECM hydrogels

Both shear rheological testing and linear compression testing of ECM hydrogels were completed using a Discovery HR-3 rheometer (TA Instruments). Rheological properties were evaluated using a 20 mm diameter parallel plate geometry and linear compression was conducted using an 8 mm diameter parallel plate geometry. To prepare each hydrogel for rheologic assessment, 100 μL of pregel solution was transferred to the center of the Peltier plate, which had been preset to 4°C. The gap distance was set to 300 μm, lowering the geometry until the pregel solution filled the space between the two parallel surfaces. The temperature of the Peltier plate was then raised to 37°C to induce gelation and held constant for 30 min. Following gelation, storage moduli of the hydrogel under 0.5% oscillatory strain at a frequency of 0.1 Hz was measured.

Hydrogels were prepared for linear compression testing as 4 mm thick disks by transferring 1.4 mL of pregel solution into a 12 well flat-bottom tissue culture plate having a surface area of ∼0.35 cm². The wells were sealed with plastic film to prevent sample dehydration, and the plate was incubated at 37°C for 30 min to induce gelation. The plastic film was removed from each well before each individual test. Linear compression strain was applied using a constant speed of 10 μm/s and stress was measured and used to calculate normalized young's modulus for each experiment. Additional samples, prepared using rat tail collagen 1 (3 mg/mL final concentration; Corning) were assessed using the same method for rheometry (n = 3), and linear compression testing (n = 2). Collagen 1 control data generated in these follow-up experiments serve as a visual reference and was not used for statistical comparison to the experimental data. Rheometry and linear compression data generated from both the experimental samples and collagen 1 controls were normalized to the 4 h timepoint for presentation.

Evaluating the effects of ECM hydrogel preparation on representative lung cell behaviors

Cell types and general culture description

Cell lines: (i) human lung fibroblasts (hLFs; hLF-CCD19Lu, ATCC CCL-210, passages 4–6) cultured in 1 × Minimum Essential Media (Cat # 11095114, Gibco) supplemented with 10% fetal bovine serum (FBS) and 1% Pen/Strep; (ii) pulmonary vascular endothelial cells (PVECs; courtesy of Mervin Yoder MD, passages 4–6) cultured in complete endothelial growth medium (Vasculife); and (iii) human mesenchymal stromal cells (hMSCs from NHLBI PACT program David McKenna MD, University of Minnesota, passages 3–5) cultured in 1 × MEM supplemented with 20% FBS and 1% Pen/Step. All cell culture media was changed every two days, both for regular maintenance and for experimental use.

Evaluating cell viability on 3D ECM hydrogels

To prepare hydrogels for cell culture studies, 100 μL of pregel solution was added per well of a 48 well plate (USA Sci; Cyto-One) and then incubated at 37°C for 1 h to allow for gelation. Rat tail collagen 1 gels (3 mg/mL final concentration; Corning) were prepared as a 3D culture control. ECM and collagen control gels were plated with n = 4 plate replicates. After gelation, cells were seeded onto the hydrogels (hLFs, PVECs, and hMSCs seeded at 5000 cells/well) and cultured for 5 days. Metabolic activity was assessed every 2 days utilizing the PrestoBlue assay according to manufacturer's instruction (Thermofisher). On the last day of culture, following the final PrestoBlue assay, Calcein-AM live cell stain was added to the cells for 1 h and images were taken at 4 × and 10 × magnification using an EVOS inverted microscope (ThermoFisher) to assess live-cell morphology.

Results

An overview of the sample preparation and experimental approach is depicted in Figure 1.

FIG. 1.

Schematic of experimental approach. To prepare tissue-derived hydrogels matrix for 3D cell culture, decellularized tissue was lyophilized, milled into a fine protein powder, and enzymatically digested. Soluble matrix proteins in the pregel solution undergo fibrillogenesis when incubated to form a self-assembled hydrogel for 3D cell culture. Alterations to structural and biochemical properties of the material due to pepsin ECM digestion duration will be investigated. 3D, three-dimensional; ECM, extracellular matrix. Color images are available online.

Tracking matrix solubilization

Tracking time dependence of protein solubilization gelation potential, and hydrogel stability

The progress of the ECM digestion progress was tracked by monitoring the pH and by quantifying protein concentration and turbidity of the pregel solutions and resulting hydrogels. The pH of all neutralized pregel samples were found to be in the correct range (∼7.4 ± 0.2); however, there was a trend in which the digest buffer pH increased with digestion time was identified. At time zero, after ECM addition, the pH was ∼2.0 as expected; however measurements at each subsequent time point showed a trend, with buffer pH increasing to: ∼2.5 at 4 h, 2.75 at 12 h, 3.25 at 24 h, and plateauing at ∼3.5 after 48 h (all pH measurements ±0.2). Two sample types were used to assess the protein concentration and turbidity of the pregel solution at each time point: (i) the uncentrifuged total protein sample and (ii) the soluble protein sample (Fig. 2A, B).

FIG. 2.

ECM solubility, gelation potential, and gelation kinetics of dECM change significantly with pepsin digestion time. The digestion of insoluble dECM powder in the total and soluble ECM fractions as a function of digestion time was tracked using (A) BCA protein quantification and (B) turbidimetric assessment and demonstrates that soluble protein in the pregel solution increases with digestion time. Turbidimetric experiments were used to quantify the (C) gelation potential and assess the (D) gelation kinetics, and further demonstrated that digestion time had a significant effect on the gelation behavior of the resulting pregel solutions. Data are presented as mean ± standard deviation with N = 9 replicates from three separate experiments (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 represents significance within each digestion time between total and soluble ECM). BCA, bicinchoninic acid; dECM, decellularized ECM. Color images are available online.

When using centrifugation to prepare the soluble ECM samples, pellet formation of undigested ECM was observed at all time points; however, the pellet formed at from the 4 h digest was visually much larger than any other time point. Further, the protein concentration and turbidity measurements of the soluble ECM sample were found to be lower than those of the total ECM sample, consistent with the insoluble protein pelleting observed at all digestion time points. This difference was found to be significant at 4 h of digest, but became nonsignificant as the soluble protein sample converged with the total ECM sample at 12 h. The convergence of the protein concentration values was maintained for the remainder of the digest; however, the turbidity values split again after 24 h, as the absorbance of the soluble ECM decreased while total ECM remained relatively stable.

Following the initial turbidity assessment the same pregel solutions were incubated to induce gelation and turbidity of the fully formed hydrogel was measured allowing for the calculation of gelation potential (Fig. 2C). The gelation potential of both samples was found to increase with digestion time. This trend plateaued around the 24 h time point with both the total protein and soluble protein samples following a similar trajectory. A second experiment to characterize gelation kinetics was conducted to directly investigate the effect of digestion time of the resulting samples, by monitoring turbidity change during gelation. The resulting normalized absorbance curves demonstrate that each digestion time results in a unique gelation curve (Fig. 2D). Qualitatively, samples prepared from shorter digestion time points started gelling earlier, and reached maximum gelation faster, then samples prepared from longer digestion time points.

Quantitative comparison of this behavior was to be made by calculating several parameters from these curves (S, t_1/2, t_95%, t_lag), which are shown in Table 1. These results were helpful, allowing us to confirm that samples prepared from longer digestion time generally took longer to start and finish their gelation process when incubated at 37°C (t_95%, t_lag), especially in comparisons between the 4 h sample and any sample prepared from a longer digest. Parameters describing the linear growth portion of the gelation curves (S, t_1/2) showed a much more consistant trend down (S) or up (t_1/2) with each subsequent digestion time point. A more detailed description of data normalization, calculation, and statistical comparisons within the parameters is shown in Supplementary Figure S1.

Table 1.

Summary of Gelation Kinetics Parameters for Total and Soluble Extracellular Matrix Pregel Samples

Parameter	S		t_1/2		t_95%		t_lag
Digest time (h)	Average (±)	Change ↑↓, p-value	Average (±)	Change ↑↓, p-value	Average (±)	Change ↑↓, p-value	Average (±)	Change ↑↓, p-value
Total protein
4	0.180 (0.005)		8.02 (0.12)		10.52 (0.09)		5.24 (0.18)
		↓,^***		↑,^***		↑,^***		↑,^***
12	0.160 (0.011)		10.69 (0.38)		13.51 (0.21)		7.55 (0.60)
		↓,^***		—,n.s.		—, n.s.		↓,^**
24	0.142 (0.004)		10.45 (0.30)		13.63 (0.23)		6.92 (0.39)
		↓,^***		↑,^***		↑,^***		↑,^***
48	0.120 (0.006)		11.94 (0.26)		15.70 (0.43)		7.77 (0.11)
		—, n.s.		—, n.s.		—,n.s.		—, n.s.
72	0.121 (0.007)		11.78 (0.44)		15.53 (0.65)		7.62 (0.22)
Soluble protein
4	0.188 (0.13)		8.49 (0.09)		10.90 (0.21)		5.81 (0.20)
		↓,^***		↑, ^***		↑,^***		—, n.s.
12	0.136 (0.007)		9.81 (0.39)		13.12 (0.29)		6.12 (0.54)
		↓,^*		↑, ^***		↑,^***		↑,^***
24	0.147 (0.006)		11.24 (0.33)		14.30 (0.44)		7.85 (0.22)
		↓,^***		↑, ^***		↑,^***		—, n.s.
48	0.117 (0.006)		12.45 (0.56)		16.30 (0.67)		8.17 (0.49)
		—, n.s.		—, n.s.		—, n.s.		—, n.s.
72	0.119 (0.006)		12.82 (0.80)		16.62 (0.88)		8.60 (0.75)

Data are presented as mean ± standard deviation with N = 9 replicates from three separate experiments (^*p < 0.05, ^**p < 0.01, ^***p < 0.001 represents significance between each subsequent time point)

n.s., nonsignificant.

Protein size distribution SDS-PAGE

Following the initial tracking assessments, the total protein pregel samples were further evaluated using SDS-PAGE to examine the effect of digestion time on the protein size distribution. A representative image of the resulting SDS-PAGE gels (Fig. 3A) demonstrates a qualitative shift in size distribution with a large proportion of high-molecular-weight proteins at earlier time points, which decreases with longer digestion times, becoming faint or disappearing entirely by 72 h. Conversely, the proportion of low-molecular-weight proteins increases at longer digestion times, with additional bands below 100 kDa appearing after 24 h of pepsin digestion. Semiquantitative image analysis of the gels (Fig. 3B) supports the qualitative observations. As digestion time increases, the results show a significant decrease in proteins larger than 250 kDa and an increase in proteins smaller than 250 kDa when compared to the 4 h time point.

FIG. 3.

Longer pepsin digestion causes ECM fragmentation. (A) Representative images of SDS-PAGE shows change in protein size distributions during digestion progression of total ECM. (B) Analysis of SDS-PAGE band intensities in the molecular weight range above 250 kDa and below 250 kDa demonstrates that solubilized matrix proteins are continuously cleaved as digestion time increases. Data are presented as mean ± standard deviations from three separate experiments. (*, **, and *** represents significance p < 0.05, 0.01, and 0.005, respectively) to 4 h of digestion and # represents significance [p < 0.05] to 12 h digestion). SDS, sodium dodecyl sulfate; PAGE, polyacrylamide gel electrophoresis.

Tracking collagen 1 digestion using fluorescent western blotting, quantification, and analysis

Fluorescent images from the collagen I western blots qualitatively demonstrate the relative amount of trimeric, dimeric, and monomeric collagen in each sample relative to the time of digestion (Fig. 4A). At lower digestion time, there is higher molecular weight collagen 1 in relationship to lower molecular weight bands. With longer digestion times, the relative amount of trimeric collagen decreases and monomeric collagen 1 increases. Semiquantitative analysis of the fluorescent signal demonstrates increased monomeric and decreased dimeric and trimeric collagen I with increasing times of pepsin digestion (Fig. 4B). The relative amount of monomeric collagen continues to increase through 72 h of digestion time. These results indicate that not all type 1 collagens are cleaved even with up to 72 h of pepsin digestion.

FIG. 4.

Collagen helical structure disassembles with longer digestion times. (A) Collagen I soluble protein profile, depicted in one of three similar replicate western blots, was used to (B) quantify collagen I monomer (140 kDa), dimer (270 kDa), and trimer (400 kDa) structure band intensity with respect to digestion time. As the digestion continues, an increasing soluble collagen is further digested from its trimeric forms to its dimer and monomer forms. Data are presented as mean ± standard deviations from three separate experiments. (*p < 0.05 and **p < 0.01 represent significance from all other groups at each time point).

Physical characterization of porcine lung extracellular matrix hydrogels

SEM imaging and calculating hydrogel interconnectivity

Shorter digestion times result in hydrogels that appear as a highly branched and woven mesh, with increasing digestion time leading to a more porous, less interwoven structure (Fig. 5A). Increasing digestion times progressively result in more linear hydrogel networks, with long spaghetti-like fibers stacked on each other. Using ImageJ analysis, the number of fiber intersections over the area of the image was used to quantify hydrogel “Interconnectivity” (Fig. 5B). Hydrogel interconnectivity was highest at lower digestion times, and significantly decreased following 4 and 12 h of digestion but then not significantly changed with up to 72 h of pepsin digestion. Collagen fiber diameter (Fig. 5C) was quantified from the same set of SEM images using ImageJ. A trend of increasing fiber diameter through 24 h was observed, after which the fiber size dropped again with longer digestion times.

FIG. 5.

Duration of pepsin digestion effects soluble ECM assembly and alters the physical and mechanical properties of the resulting hydrogels. (A) Representative SEM images of ECM hydrogels prepared from matrix solubilized for 4, 12, 24, 48, and 72 h show enzymatic digestion effects on material architecture (scale bar = 1 μm). Analysis of hydrogel SEM images conducted using ImageJ to evaluate (B) fiber diameter, which increases with digestion time and peaks at 24 h, decreasing with longer digest times, and (C) interconnectivity, which is highest at shorter digestion times, decreasing with longer digestion times. Mechanical testing of the resulting ECM hydrogels using (D) shear rheology and (E) linear compression testing demonstrate a significant reduction in normalized storage modulus at 48 h of digestion, while Young's modulus increases with digestion time to a peak at 24 h, before falling off at longer digestion times. Collagen 1 controls conducted separately are presented as a visual reference to the experimental mechanics data. Data are shown as mean ± standard deviations from three separate experiments, calculated using three representative areas per experiment (^n.sp > 0.05, *p < 0.05 and **p < 0.01 denotes significance between groups designated by brackets; while ^###p < 0.001, denotes significance to all other groups). SEM, scanning electron microscopy; n.s., nonsignificant.

Mechanical evaluation of ECM hydrogels

Shear rheological testing (Fig. 5D) demonstrated that the storage modulus was higher and relatively stable for hydrogels prepared from shorter 4, 12, and 24 h digestion times and then dropped significantly for hydrogels prepared from the longer 48 and 72 h digestion times. Alternatively, linear compression testing (Fig. 5E) demonstrated a trend in increasing Young's modulus with digestion time from 4 to 24 h, decreasing at 48 h following the 24 h sample peak. The 72 h digests formed hydrogels; however, they were too weak to be tested using this approach.

Cell response to ECM hydrogels

All three cell lines evaluated were successfully cultured on ECM hydrogels prepared from all digestion time points (Fig. 6A and Supplementary Fig. S2). However, Calcien AM cell staining demonstrated that there were qualitatively more cells observed on gels prepared from the 4, 12, and 24 h time points in comparison to cells grown on gels prepared from 48 and 72 h digests. Qualitatively, both hLFs and hMSCs appear to be more elongated and demonstrated some clustering organization on gels from shorter 4 and 12 h digests compared to gels derived from longer digests. PVECs, when grown on hydrogels prepared from 4 and 12 h digests, appeared to organize into semibranched clusters, when compared to cells grown on gels prepared from longer digestion times, which did not. Quantitative assessment of all three cell lines after 5 days of culture with PrestoBlue demonstrated that the metabolic activity of viable cells was higher on gels prepared from 4, 12, and 24 h digests, and decreased significantly with longer digestion time for all cell types (Fig. 6B).

FIG. 6.

Physical changes in hydrogel with digestion time effect the appearance and metabolic activity of viable cells. PVECs, hLFs, or hMSCs were cultured on top of ECM hydrogels digested for 4, 12, 24, 48, or 72 h or on collagen hydrogels for 5 days to assess (A) morphology with Calcein AM staining and (B) metabolic activity of viable cells with a PrestoBlue viability assay. Data are presented as mean ± standard deviation. N = 12 replicates from three experiments (*p < 0.05, **p < 0.01, ***p < 0.001 denote significance between groups). Scale Bars = 400 μm. hLF, human lung fibroblast; hMSC, human mesenchymal stromal cell. Color images are available online.

Discussion

Over the past 10 years, the adaptability and unique advantages offered by ECM hydrogels have driven their development for an increasing number of impactful applications in both the laboratory and clinic. As an in vitro research model, hydrogels derived from a wide range of organ sources have demonstrated the capacity to induce tissue-specific cell behaviors.^{31,45,53,60,62,70,81} As a clinical material, ECM hydrogels are readily remodeled by immune cells and can induce a proregenerative response in vivo.^82–84 In addition, the resulting degradation products attract proregenerative endogenous stem cell populations, reduce additional proinflammatory immune cell recruitment,^39,85–87 and have been demonstrated to have antibacterial effects.^88,89 The rapid adoption of ECM hydrogels has been driven by both the availability of standardized ECM processing protocols and the ability of the resulting hydrogels to mimic the ECM microenvironment.^36,61,90,91 However, the connection between ECM digestion parameters and the physical, mechanical, and biologic properties of the resulting hydrogels has been understudied.

The isolation of pepsin-solubilized collagen 1 and preparation of collagen hydrogels have been studied extensively over the past 30 years and are a useful comparison as collagens make up a significant proportion of tissue-derived dECM. Several studies have demonstrated that pepsin isolation of soluble collagen and the resulting hydrogel properties are tied to digestion parameters, including time, temperature, and enzyme to substrate ratio.^78,92 In this work, we hypothesized that like collagen 1, digestion parameters play an important role in determining the properties of ECM hydrogels as well, and digestion time was selected for further evaluation. Currently, the majority of ECM hydrogels are prepared by adherence to standard 48 to 72 h long digestion, a strategy that asserts that maximizing the proportion of soluble ECM should be the main focus for preparing hydrogels.³⁶ To challenge this principle, we followed dECM solubilization over time and characterized the resulting ECM hydrogels using samples isolated from ongoing digestions at 4, 12, 24, 48, and 72 h.

The initial digestion tracking studies suggest that a large portion of the dECM powder is solubilized by 12 h, while the remaining insoluble powder was solubilized at a significantly reduced rate. Qualitative observations of the insoluble protein pellet were consistent with the quantitative protein and turbidity assay results, in which a significant difference between the total and soluble ECM samples at 4 h converged to nonsignificant by 12 h of digestion. One possible explanation is that the enzymatic activity of pepsin was reduced as the pH of the digestion buffer increased with longer digestion times. Pepsin exhibits peak activity at a pH of ∼2.0, however, as the pH increased steadily to ∼3.5 at 48 h, the activity would decrease to around 70%.⁷²

Interestingly, the gelation potential results may suggest that the undigested ECM does not play a role in hydrogel assembly as both the total and soluble ECM samples increased with digestion time until plateauing between 12 and 24 h. Further, the results support the hypothesis that longer digestion times are not required for successful hydrogel formation as the majority of gellable ECM proteins were already soluble by 24 h. A deeper assessment of the gelation kinetics and the calculated parameters further demonstrated that sample behavior was directly related to digestion time. Parameters describing the linear “growth” region of the gelation kinetic curve had a similar trend as other tracking experiments, with increases in digestion time reducing the speed of gelation (S), while increasing the duration of the gelation curve (t_1/2, t_95%). However, the parameter describing the time until gelation starts (t_lag) was slightly different, with a gap between the shortest time point, 4 h, and the longer time points, which may suggest that this behavior is related to more complex aspects of the pregel solution composition.

While these tracking assessments were useful, they were not perfect. The relative sensitivity of protein assays is directly related to protein size and some like the Bradford assay have difficulty detecting proteins below a certain size threshold for quantification.^93,94 This adds another layer of complexity when comparing enzymatically digested ECM samples which have heterogeneous molecular weight distributions.

To more thoroughly evaluate the possible effect of nonspecific cleavage, SDS-PAGE demonstrated that the relative proportion of larger soluble proteins (>250 kDa) decreased, while the relative proportion of smaller soluble proteins (<250 kDa) increased with longer digestion times. Specific western blotting analysis of collagen 1 distribution demonstrated similar effects, with larger trimeric collagen 1 decreasing with digestion time, while monomeric and submonomeric collagen 1 increased. Both these results further support the suggestion that already soluble proteins are subject to continued cleavage with longer digestion times. Increasing fragmentation of soluble proteins may explain the relative decrease in protein and turbidity measurements observed at longer digestion times as increasingly heterogeneous samples complicate colorimetric protein quantification, while smaller protein and peptide fragments absorb and defract less light during turbidity assessment.

The next stage of our investigation assessed correlations between time-dependent digestion effects on protein solubilization and the physical architecture and mechanical properties of the resulting hydrogels. Qualitative SEM analysis demonstrated that hydrogels produced at shorter digestion times were more branched like a woven mesh, while the architecture became increasingly linear with longer digestion times. Hydrogels produced from 4 h digests were determined to have the highest interconnectivity, significantly decreasing at 12 h, decreasing further from 12 to 24 h, and then remaining relatively stable at longer digest times. Comparatively, the fiber diameter was the lowest at 4 h, increased with digestion time to a peak at 24 h, and then fell off with longer 48 and 72 h digestions. Despite the observed relationships, understanding the functional implications between digestion time and resulting hydrogel architecture remains challenging.

One possible explanation of the hydrogel architecture results is that the participation of individual proteins in hydrogel polymerization is also time-dependant. Previous investigations of collagen 1 assembly, a less complex system, have demonstrated the importance of nucleating proteins during collagen 1 fibrillogenesis or polymerization. Increasing concentrations of dermatan sulfate, heparin, fibronectin, and collagen VI have been shown to decrease the fiber density, delay fibril formation, and increase fiber thickness of resulting collagen hydrogels, while increases in chondroitin sulfate or decorin delay fibril formation promoting more branched thinner fibrils.^95–101

In another key study, investigators used mass spectrometry to evaluate digestion time-dependent differences in the composition of pepsin solubilized cardiac dECM. Interestingly, this study found that the isolation of specific ECM components from the insoluble powder was time-dependent, as basement membrane proteins such as fibrillin-1, fibrinogen, and laminins were soluble at shorter digestion times, while large fibrillar ECM components such as collagen 1 took longer.⁷⁹

In this context, the complex architecture of ECM hydrogels may be explained by the relative availability of nucleating proteins in the soluble fraction. At shorter digestion times, a larger proportion of nucleators results in a complex branched hydrogel, while a steady increase in soluble collagens is responsible for the transition to a more linear architecture by 24 h. For example, t_l_ag is one parameter that might be effected if nucleating proteins were playing a role in the gelation behavior, fibril assembly, and hydrogel architecture. The presence of nucleating proteins would induce faster gelation. If this is the case, our results may suggest the presence of more nucleating proteins (total or proportionally), in pregel samples produced from 4 h digests when compared to other samples. While this is an exciting possiblility, additional investigation of soluble ECM composition is required to better understand the relationship.

Interestingly, while interconnectivity and fiber diameter followed separate trends with digestion time, each measure correlated with one of the mechanical properties assessed. The oscillatory shear rheometry results followed a trend similar to that measured with hydrogel interconnectivity, where the storage modulus was highest at digestion times between 4 and 24 h, but decreased significantly with longer 48 and 72 h digestion times. This result suggests that hydrogels prepared with shorter pepsin digest times provide more resistance to shear deformation due to their highly branched and interconnected architecture, while hydrogels with linear architecture had less resistance to shear forces. Conversely, the linear compression results were closely correlated with average fiber diameter. At 4 h, the Young's modulus and fiber diameter of the resulting hydrogels was relatively low but increased significantly with digestion time to a peak at 24 h, before trending down with longer digestion times.

In summary, the cumulative architectural and mechanical analyses indicate a clear transition at 24 h, whereafter longer digestion times result in less complex hydrogels with significantly weaker mechanical properties.

Assessing how the time-dependant physical and mechanical properties of the resulting ECM hydrogels affected the behavior of relevant lung cells demonstrated that, overall, qualitatively more cells were observed on hydrogels prepared from shorter 4 to 24 h digestion times and more characteristic morphology of fibroblasts and vascular endothelial cells observed. Further, there was increased metabolic activity of cells grown on hydrogels prepared from dECM digested between 4 and 24 h, when compared to those grown on hydrogels prepared form 48 or 72 h digests. Although these are relatively simple studies, the results allow us to draw initial connections between the physical properties of the resulting hydrogels and some cell behaviors. While all the cell types were able to survive and grow on hydrogels generated from every time point, the results suggest that the cells performed better on mechanically robust and architecturally complex hydrogels in comparison to weaker, more linear hydrogels.

Overall, longer digestion times of pig lung-derived dECM resulted in less mechanically robust hydrogels, with less structural heterogeneity and reduced ability to support cultured cells. These findings suggest that the protocols used to solubilize dECM can be optimized to improve the performance of ECM hydrogels as they are adapted for a widening variety of applications. For example, investigators developing an ECM model might identify the parameters to prepare the most compositionally and architecturally complex hydrogel, while those preparing materials for biomanufacturing or a bioink might prioritize parameters to create the most mechanically robust hydrogel.

The generalized trends discovered in this study are limited, and only describe the relationship between hydrogel properties and digestion time for porcine lung ECM hydrogels prepared using the specific materials processing and enzymatic digestion methods described here. This study did not include dECM from other tissue sources; however, it is reasonable to suggest that similar relationships between digestion time and hydrogel properties exist for ECM hydrogels derived from other tissues or using prepared using other methods. We have identified that manipulation of the enzymatic digestion process provides another pathway for controlling ECM hydrogel properties and function, but it is clear that these releationships will have to be defined for other systems individually.

Conclusion

The duration of enzymatic digestion of pig lung dECM significantly affects protein solubilization, soluble protein size distribution, gelation potential, and downstream compositional, mechanical, and biologic properties of derivative hydrogels. Notably, potentially deleterious effects on hydrogel architecture, mechanical properties, and capacity for supporting live cell culture of several relevant lung cell lines were observed at longer digestion times. These are significant observations as most ECM hydrogels are enzymatically digested for relatively long times at 48 or 72 h to maximize soluble ECM. Overall, adherence to standardized ECM digestion practices may be limiting the potential utilities of ECM hydrogels as they are increasingly adapted for impactful application as tissue engineering materials, as tools to study matrix biology, and as products for clinical translation. While this study clarifies the importance of a single parameter, digestion time, it also highlights the possibility that other parameters might have equal or greater ability to optimize or tailor ECM hydrogel properties.

Footnotes

Disclosure Statement

No competing financial interests exist.

Funding Information

This work was supported by the National Institutes of Health (R01 HL127144 and R21 EB024329 to D.J.W.; R01 AG04182 to R.L.H.) and the National Science Foundation (CMMI 135162 to R.L.H.).

Supplementary Material

References

Petersen

T.H.

, Calle

E.A.

, Zhao

, et al. Tissue-engineered lungs for in vivo implantation. Science, 329, 538, 2010.

Lin

, Zhang

, Rippon

H.J.

, Bismarck

, and Bishop

A.E.

Tissue engineering of lung: the effect of extracellular matrix on the differentiation of embryonic stem cells to pneumocytes. Tissue Eng Part A, 16, 1515, 2009.

Cortiella

, Niles

, Cantu

, et al. Influence of acellular natural lung matrix on murine embryonic stem cell differentiation and tissue formation. Tissue Eng Part A, 16, 2565, 2010.

Price

A.P.

, England

K.A.

, Matson

A.M.

, Blazar

B.R.

, and Panoskaltsis-Mortari

Development of a decellularized lung bioreactor system for bioengineering the lung: the matrix reloaded. Tissue Eng Part A, 16, 2581, 2010.

Song

J.J.

, Kim

S.S.

, Liu

, et al. Enhanced in vivo function of bioartificial lungs in rats. Ann Thorac Surg, 92, 998, 2011.

Daly

A.B.

, Wallis

J.M.

, Borg

Z.D.

, et al. Initial binding and recellularization of decellularized mouse lung scaffolds with bone marrow-derived mesenchymal stromal cells. Tissue Eng Part A, 18, 1, 2012.

Booth

A.J.

, Hadley

, Cornett

A.M.

, et al. Acellular normal and fibrotic human lung matrices as a culture system for in vitro investigation. Am J Respir Crit Care Med, 186, 866, 2012.

Sokocevic

, Bonenfant

N.R.

, Wagner

D.E.

, et al. The effect of age and emphysematous and fibrotic injury on the re-cellularization of de-cellularized lungs. Biomaterials, 34, 3256, 2013.

Wagner

D.E.

, Bonenfant

N.R.

, Parsons

C.S.

, et al. Comparative decellularization and recellularization of normal versus emphysematous human lungs. Biomaterials, 35, 3281, 2014.

10.

Wagner

D.E.

, Bonenfant

N.R.

, Sokocevic

, et al. Three-dimensional scaffolds of acellular human and porcine lungs for high throughput studies of lung disease and regeneration. Biomaterials, 35, 2664, 2014.

11.

Jensen

, Roszell

, Zang

, et al. A rapid lung de-cellularization protocol supports embryonic stem cell differentiation in vitro and following implantation. Tissue Eng Part C Methods, 18, 632, 2012.

12.

Uriel

, Labay

, Francis-Sedlak

, et al. Extraction and assembly of tissue-derived gels for cell culture and tissue engineering. Tissue Eng Part C Methods, 15, 309, 2009.

13.

Choudhury

, Tun

H.W.

, Wang

, and Naing

M.W.

Organ-derived decellularized extracellular matrix: a game changer for bioink manufacturing? Trends Biotechnol, 36, 787, 2018.

14.

Kim

B.S.

, Kim

, Gao

, Jang

, and Cho

D.-W.

Decellularized extracellular matrix: a step towards the next generation source for bioink manufacturing. Biofabrication, 9, 034104, 2017.

15.

Xing

, Qian

, Jia

, Ghosh

, Tahtinen

, and Zhao

Natural extracellular matrix for cellular and tissue biomanufacturing. ACS Biomater Sci Eng, 3, 1462, 2017.

16.

Krishtul

, Baruch

, and Machluf

Processed tissue-derived extracellular matrices: tailored platforms empowering diverse therapeutic applications. Adv Funct Mater, 1900386, 2019.

17.

Sundarakrishnan

, Chen

, Black

L.D.

, Aldridge

B.B.

, and Kaplan

D.L.

Engineered cell and tissue models of pulmonary fibrosis. Adv Drug Deliv Rev, 129, 78, 2018.

18.

Gilbert

T.W.

, Stolz

D.B.

, Biancaniello

, Simmons-Byrd

, and Badylak

S.F.

Production and characterization of ECM powder: implications for tissue engineering applications. Biomaterials, 26, 1431, 2005.

19.

Choi

J.S.

, Yang

H.-J.

, Kim

B.S.

, et al. Human extracellular matrix (ECM) powders for injectable cell delivery and adipose tissue engineering. J Control Release, 139, 2, 2009.

20.

Beachley

, Ma

, Papadimitriou

, Gibson

, Corvelli

, and Elisseeff

Extracellular matrix particle-glycosaminoglycan composite hydrogels for regenerative medicine applications. J Biomed Mater Res A, 106, 147, 2018.

21.

Manni

M.L.

, Czajka

C.A.

, Oury

T.D.

, and Gilbert

T.W.

Extracellular matrix powder protects against bleomycin-induced pulmonary fibrosis. Tissue Eng Part A, 17, 2795, 2011.

22.

Hoganson

D.M.

, Owens

G.E.

, Meppelink

A.M.

, et al. Decellularized extracellular matrix microparticles as a vehicle for cellular delivery in a model of anastomosis healing: decellularized ECM microparticles. J Biomed Mater Res A, 104, 1728, 2016.

23.

Link

P.A.

, Ritchie

A.M.

, Cotman

G.M.

, Valentine

M.S.

, Dereski

B.S.

, and Heise

R.L.

Electrosprayed extracellular matrix nanoparticles induce a pro-regenerative cell response. J Tissue Eng Regen Med, 12, 2331, 2018.

24.

Young

B.M.

, Shankar

, Allen

B.P.

, et al. Electrospun decellularized lung matrix scaffold for airway smooth muscle culture. ACS Biomater Sci Eng, 3, 3480, 2017.

25.

Jakus

A.E.

, Laronda

M.M.

, Rashedi

A.S.

, et al. “Tissue papers” from organ-specific decellularized extracellular matrices. Adv Funct Mater, 27, 1700992, 2017.

26.

Pouliot

R.A.

, Link

P.A.

, Mikhaiel

N.S.

, et al. Development and characterization of a naturally derived lung extracellular matrix hydrogel. J Biomed Mater Res A, 104, 1922, 2016.

27.

Link

P.A.

, Pouliot

R.A.

, Mikhaiel

N.S.

, Young

B.M.

, and Heise

R.L.

Tunable hydrogels from pulmonary extracellular matrix for 3D cell culture. J Vis Exp, 119, 55094, 2017.

28.

, Nguyen

K.T.

, and Hong

Engineering a three-dimensional cancer model from lung extracellular matrix hydrogel. Abstract presented at the 10th World Biomaterials Congress, Montreal, Canada, 2016.

29.

Jiang

, Chaimov

, Patel

S.N.

, et al. 3-D physiomimetic extracellular matrix hydrogels provide a supportive microenvironment for rodent and human islet culture. Biomaterials, 198, 37, 2019.

30.

Zhou

, Wu

, Sun

, Zhou

, Zhang

, and Xiao

Lung tissue extracellular matrix-derived hydrogels protect against radiation-induced lung injury by suppressing epithelial–mesenchymal transition. J Cell Physiol, 235, 2377, 2020.

31.

Young

B.M.

, Shankar

, Tho

C.K.

, Pellegrino

A.R.

, and Heise

R.L.

Laminin-driven Epac/Rap1 regulation of epithelial barriers on decellularized matrix. Acta Biomater, 100, 223, 2019

32.

de Hilster

R.H.J.

, Sharma

P.K.

, Jonker

M.R.

, et al. Human lung extracellular matrix hydrogels resemble the stiffness and viscoelasticity of native lung tissue. Am J Physiol Lung Cell Mol Physiol, 318, L698, 2020.

33.

Farre

, Otero

, Falcones

, Marhuenda

, Almendros

, and Navajas

Bioprinted 3D model to study the crosstalk between lung mesenchymal stem cells and lung extracellular matrix. Presented at the American Thoracic Society International Conference. Dallas, TX, 2019.

34.

Santis

M.D.

, Ota

, Costa

, et al. Development of a hybrid alginate-ECM hydrogel as a potential bioink for 3D bioprinting. Eur Respir J, 52, LSC, 2018.

35.

Freytes

D.O.

, Martin

, Velankar

S.S.

, Lee

A.S.

, and Badylak

S.F.

Preparation and rheological characterization of a gel form of the porcine urinary bladder matrix. Biomaterials, 29, 1630, 2008.

36.

Saldin

L.T.

, Cramer

M.C.

, Velankar

S.S.

, White

L.J.

, and Badylak

S.F.

Extracellular matrix hydrogels from decellularized tissues: structure and function. Acta Biomater, 49, 1, 2017.

37.

Voytik-Harbin

S.L.

, Brightman

A.O.

, Waisner

B.Z.

, Robinson

J.P.

, and Lamar

C.H.

Small intestinal submucosa: a tissue-derived extracellular matrix that promotes tissue-specific growth and differentiation of cells in vitro. Tissue Eng, 4, 157, 1998.

38.

Massensini

A.R.

, Ghuman

, Saldin

L.T.

, et al. Concentration-dependent rheological properties of ECM hydrogel for intracerebral delivery to a stroke cavity. Acta Biomater, 27, 116, 2015.

39.

Reing

J.E.

, Zhang

, Myers-Irvin

, et al. Degradation products of extracellular matrix affect cell migration and proliferation. Tissue Eng Part A, 15, 605, 2008.

40.

Slivka

P.F.

, Dearth

C.L.

, Keane

T.J.

, et al. Fractionation of an ECM hydrogel into structural and soluble components reveals distinctive roles in regulating macrophage behavior. Biomater Sci, 2, 1521, 2014.

41.

Young

D.A.

, Ibrahim

D.O.

, Hu

, and Christman

K.L.

Injectable hydrogel scaffold from decellularized human lipoaspirate. Acta Biomater, 7, 1040, 2011.

42.

Jeffords

M.E.

, Wu

, Shah

, Hong

, and Zhang

Tailoring material properties of cardiac matrix hydrogels to induce endothelial differentiation of human mesenchymal stem cells. ACS Appl Mater Interfaces, 7, 11053, 2015.

43.

Wassenaar

J.W.

, Braden

R.L.

, Osborn

K.G.

, and Christman

K.L.

Modulating in vivo degradation rate of injectable extracellular matrix hydrogels. J Mater Chem B, 4, 2794, 2016.

44.

Johnson

T.D.

, DeQuach

J.A.

, Gaetani

, et al. Human versus porcine tissue sourcing for an injectable myocardial matrix hydrogel. Biomater Sci, 2, 735, 2014.

45.

Duan

, Liu

, O'Neill

, Wan

L.Q.

, Freytes

D.O.

, and Vunjak-Novakovic

Hybrid gel composed of native heart matrix and collagen induces cardiac differentiation of human embryonic stem cells without supplemental growth factors. J Cardiovasc Transl Res, 4, 605, 2011.

46.

Singelyn

J.M.

, Sundaramurthy

, Johnson

T.D.

, et al. Catheter-deliverable hydrogel derived from decellularized ventricular extracellular matrix increases endogenous cardiomyocytes and preserves cardiac function post-myocardial infarction. J Am Coll Cardiol, 59, 751, 2012.

47.

Hong

, Kim

, Han

S.J.

, et al. Compressed collagen intermixed with cornea-derived decellularized extracellular matrix providing mechanical and biochemical niches for corneal stroma analogue. Mater Sci Eng C, 103, 109837, 2019.

48.

Wolf

M.T.

, Daly

K.A.

, Brennan-Pierce

E.P.

, et al. A hydrogel derived from decellularized dermal extracellular matrix. Biomaterials, 33, 7028, 2012.

49.

Lee

, Han

, Kim

, et al. Development of liver decellularized extracellular matrix bioink for three-dimensional cell printing-based liver tissue engineering. Biomacromolecules, 18, 1229, 2017.

50.

Lee

J.S.

, Shin

, Park

H.-M.

, et al. Liver extracellular matrix providing dual functions of two-dimensional substrate coating and three-dimensional injectable hydrogel platform for liver tissue engineering. Biomacromolecules, 15, 206, 2014.

51.

Coronado

R.E.

, Somaraki-Cormier

, Natesan

, Christy

R.J.

, Ong

J.L.

, and Halff

G.A.

Decellularization and solubilization of porcine liver for use as a substrate for porcine hepatocyte culture: method optimization and comparison. Cell Transplant, 26, 1840, 2017.

52.

Loneker

A.E.

, Faulk

D.M.

, Hussey

G.S.

, D'Amore

, and Badylak

S.F.

Solubilized liver extracellular matrix maintains primary rat hepatocyte phenotype in-vitro. J Biomed Mater Res A, 104, 957, 2016.

53.

Keane

T.J.

, DeWard

, Londono

, et al. Tissue-specific effects of esophageal extracellular matrix. Tissue Eng Part A, 21, 2293, 2015.

54.

Sawkins

M.J.

, Bowen

, Dhadda

, et al. Hydrogels derived from demineralized and decellularized bone extracellular matrix. Acta Biomater, 9, 7865, 2013.

55.

Burnsed

O.A.

, Schwartz

, Marchand

K.O.

, Hyzy

S.L.

, Olivares-Navarrete

, and Boyan

B.D.

Hydrogels derived from cartilage matrices promote induction of human mesenchymal stem cell chondrogenic differentiation. Acta Biomater, 43, 139, 2016.

56.

Kwon

J.S.

, Yoon

S.M.

, Shim

S.W.

, et al. Injectable extracellular matrix hydrogel developed using porcine articular cartilage. Int J Pharm, 454, 183, 2013.

57.

Mercuri

J.J.

, Patnaik

, Dion

, Gill

S.S.

, Liao

, and Simionescu

D.T.

Regenerative potential of decellularized porcine nucleus pulposus hydrogel scaffolds: stem cell differentiation, matrix remodeling, and biocompatibility studies. Tissue Eng Part A, 19, 952, 2013.

58.

Medberry

C.J.

, Crapo

P.M.

, Siu

B.F.

, et al. Hydrogels derived from central nervous system extracellular matrix. Biomaterials, 34, 1033, 2013.

59.

Kočí

, Výborný

, Dubišová

, et al. Extracellular matrix hydrogel derived from human umbilical cord as a scaffold for neural tissue repair and its comparison with extracellular matrix from porcine tissues. Tissue Eng Part C Methods, 23, 333, 2017.

60.

Palade

, Pal

, Rawls

, Stabenfeldt

, and Wilson-Rawls

Molecular analysis of muscle progenitor cells on extracellular matrix coatings and hydrogels. Acta Biomater, 97, 296, 2019.

61.

Sasikumar

, Chameettachal

, Cromer

, Pati

, and Kingshott

Decellularized extracellular matrix hydrogels—cell behavior as a function of matrix stiffness. Curr Opin Biomed Eng, 10, 123, 2019.

62.

Agmon

, and Christman

K.L.

Controlling stem cell behavior with decellularized extracellular matrix scaffolds. Curr Opin Solid State Mater Sci, 20, 193, 2016.

63.

Johnson

T.D.

, Lin

S.Y.

, and Christman

K.L.

Tailoring material properties of a nanofibrous extracellular matrix derived hydrogel. Nanotechnology, 22, 494015, 2011.

64.

Singelyn

J.M.

, and Christman

K.L.

Modulation of material properties of a decellularized myocardial matrix scaffold. Macromol Biosci, 11, 731, 2011.

65.

Ahearne

, and Coyle

Application of UVA-riboflavin crosslinking to enhance the mechanical properties of extracellular matrix derived hydrogels. J Mech Behav Biomed Mater, 54, 259, 2016.

66.

Jang

, Kim

T.G.

, Kim

B.S.

, Kim

S.-W.

, Kwon

S.-M.

, and Cho

D.-W.

Tailoring mechanical properties of decellularized extracellular matrix bioink by vitamin B2-induced photo-crosslinking. Acta Biomater, 33, 88, 2016.

67.

Fujita

, Feng

, Sato

, et al. Modulation of the mechanical properties of ventricular extracellular matrix hydrogels with a carbodiimide crosslinker and investigation of their cellular compatibility. AIMS Mater Sci, 5, 54, 2018.

68.

Visser

, Levett

P.A.

, te Moller

N.C.R.

, et al. Crosslinkable hydrogels derived from cartilage, meniscus, and tendon tissue. Tissue Eng Part A, 21, 1195, 2015.

69.

Skardal

, Devarasetty

, Kang

H.-W.

, et al. A hydrogel bioink toolkit for mimicking native tissue biochemical and mechanical properties in bioprinted tissue constructs. Acta Biomater, 25, 24, 2015.

70.

Ali

, Pr

A.K.

, Yoo

J.J.

, Zahran

, Atala

, and Lee

S.J.

A photo-crosslinkable kidney ECM-derived bioink accelerates renal tissue formation. Adv Healthc Mater, 8, 1800992, 2019.

71.

Grover

G.N.

, Rao

, and Christman

K.L.

Myocardial matrix–polyethylene glycol hybrid hydrogels for tissue engineering. Nanotechnology, 25, 014011, 2014.

72.

Kondjoyan

, Daudin

J.-D.

, and Santé-Lhoutellier

Modelling of pepsin digestibility of myofibrillar proteins and of variations due to heating. Food Chem, 172, 265, 2015.

73.

Miller

E.J.

Structural studies on cartilage collagen employing limited cleavage and solubilization with pepsin. Biochemistry, 11, 4903, 1972.

74.

Northrop

J.H.

The effect of various acids on the digestion of proteins by pepsin. J Gen Physiol, 1, 607, 1919.

75.

Ahn

, Cao

M.-J.

, Yu

Y.Q.

, and Engen

J.R.

Accessing the reproducibility and specificity of pepsin and other aspartic proteases. Biochim Biophys Acta, 1834, 1222, 2013.

76.

Holmes

, Kirk

, Tronci

, Yang

, and Wood

Influence of telopeptides on the structural and physical properties of polymeric and monomeric acid-soluble type I collagen. Mater Sci Eng C, 77, 823, 2017.

77.

Hamuro

, Coales

S.J.

, Molnar

K.S.

, Tuske

S.J.

, and Morrow

J.A.

Specificity of immobilized porcine pepsin in H/D exchange compatible conditions. Rapid Commun Mass Spectrom, 22, 1041, 2008.

78.

Lin

Y.K.

, and Liu

D.C.

Effects of pepsin digestion at different temperatures and times on properties of telopeptide-poor collagen from bird feet. Food Chem, 94, 621, 2006.

79.

Williams

, Sullivan

, and Black

L.D.

Partially digested adult cardiac extracellular matrix promotes cardiomyocyte proliferation in vitro. Adv Healthc Mater, 4, 1545, 2015.

80.

Baker

S.C.

, Atkin

, Gunning

P.A.

, et al. Characterisation of electrospun polystyrene scaffolds for three-dimensional in vitro biological studies. Biomaterials, 27, 3136, 2006.

81.

Sellaro

T.L.

, Ranade

, Faulk

D.M.

, et al. Maintenance of human hepatocyte function in vitro by liver-derived extracellular matrix gels. Tissue Eng Part A, 16, 1075, 2009.

82.

Badylak

S.F.

, Valentin

J.E.

, Ravindra

A.K.

, McCabe

G.P.

, and Stewart-Akers

A.M.

Macrophage phenotype as a determinant of biologic scaffold remodeling. Tissue Eng Part A, 14, 1835, 2008.

83.

Brown

B.N.

, Londono

, Tottey

, et al. Macrophage phenotype as a predictor of constructive remodeling following the implantation of biologically derived surgical mesh materials. Acta Biomater, 8, 978, 2012.

84.

Brown

B.N.

, Valentin

J.E.

, Stewart-Akers

A.M.

, McCabe

G.P.

, and Badylak

S.F.

Macrophage phenotype and remodeling outcomes in response to biologic scaffolds with and without a cellular component. Biomaterials, 30, 1482, 2009.

85.

Agrawal

, Brown

B.N.

, Beattie

A.J.

, Gilbert

T.W.

, and Badylak

S.F.

Evidence of innervation following extracellular matrix scaffold-mediated remodelling of muscular tissues. J Tissue Eng Regen Med, 3, 590, 2009.

86.

Agrawal

, Johnson

S.A.

, Reing

, et al. Epimorphic regeneration approach to tissue replacement in adult mammals. Proc Natl Acad Sci U S A, 107, 3351, 2010.

87.

Agrawal

, Tottey

, Johnson

S.A.

, Freund

J.M.

, Siu

B.F.

, and Badylak

S.F.

Recruitment of progenitor cells by an extracellular matrix cryptic peptide in a mouse model of digit amputation. Tissue Eng Part A, 17, 2435, 2011.

88.

Brennan

E.P.

, Reing

, Chew

, Myers-Irvin

J.M.

, Young

E.J.

, and Badylak

S.F.

Antibacterial activity within degradation products of biological scaffolds composed of extracellular matrix. Tissue Eng, 12, 2949, 2006.

89.

Gupta

, Dinda

, and Mishra

N.C.

Antibacterial activity and composition of decellularized goat lung extracellular matrix for its tissue engineering applications. Biol Eng Med, 2, 1000112, 2017.

90.

Hussey

G.S.

, Cramer

M.C.

, and Badylak

S.F.

Extracellular matrix bioscaffolds for building gastrointestinal tissue. Cell Mol Gastroenterol Hepatol, 5, 1, 2018.

91.

Smithmyer

M.E.

, Sawicki

L.A.

, and Kloxin

A.M.

Hydrogel scaffolds as in vitro models to study fibroblast activation in wound healing and disease. Biomater Sci, 2, 634, 2014.

92.

, Zong

, Jin

, et al. Optimization of extraction conditions and characterization of pepsin-solubilised collagen from skin of giant croaker (Nibea japonica). Mar Drugs, 16, 29, 2018.

93.

Bass

J.J.

, Wilkinson

D.J.

, Rankin

, et al. An overview of technical considerations for Western blotting applications to physiological research. Scand J Med Sci Sports, 27, 4, 2017.

94.

Krohn

R.I.

The colorimetric detection and quantitation of total protein. Curr Protoc Cell Biol, 52, A.3H.1, 2011.

95.

Izu

, Ansorge

H.L.

, Zhang

, et al. Dysfunctional tendon collagen fibrillogenesis in collagen VI null mice. Matrix Biol, 30, 53, 2011.

96.

Stamov

D.R.

, Khoa Nguyen

T.A.

, Evans

H.M.

, Pfohl

, Werner

, and Pompe

The impact of heparin intercalation at specific binding sites in telopeptide-free collagen type I fibrils. Biomaterials, 32, 7444, 2011.

97.

, Patra

, and Faezipour

Structure of collagen-glycosaminoglycan matrix and the influence to its integrity and stability. Presented at 36th Annual International Conference IEEE Engineering in Medicine and Biology Society. Chicago, IL, USA, 2014, pp. 3949–3952.

98.

Mattson

J.M.

, Turcotte

, and Zhang

Glycosaminoglycans contribute to extracellular matrix fiber recruitment and arterial wall mechanics. Biomech Model Mechanobiol, 16, 213, 2017.

99.

Milan

A.M.

, Sugars

R.V.

, Embery

, and Waddington

R.J.

Modulation of collagen fibrillogenesis by dentinal proteoglycans. Calcif Tissue Int, 76, 127, 2005.

100.

McDonald

J.A.

, Kelley

D.G.

, and Broekelmann

T.J.

Role of fibronectin in collagen deposition: fab′ to the gelatin-binding domain of fibronectin inhibits both fibronectin and collagen organization in fibroblast extracellular matrix. J Cell Biol, 92, 485, 1982.

101.

Kubow

K.E.

, Vukmirovic

, Zhe

, et al. Mechanical forces regulate the interactions of fibronectin and collagen I in extracellular matrix. Nat Commun, 6, 8026, 2015.

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