Comparative Structural and Functional Analysis of Micronized Collagen-Based Scaffolds

Abstract

Micronized collagen-based bioscaffolds are increasingly used in clinical applications for wound repair and soft tissue regeneration. This study compared the structural properties of four different commercially available micronized products derived from either reconstituted collagen (pRC), urinary bladder matrix (pUBM), or ovine forestomach matrix (mOFM, mOFMµ). The test articles were characterized by laser diffraction analysis, scanning electron microscopy (SEM), micro-computed tomography (micro-CT), packing density, differential scanning calorimetry, rheometry, proteolytic stability, agarose gel electrophoresis, and blood clotting index. Particle size and surface morphology, assessed by laser diffraction, SEM, and micro-CT, revealed marked differences in particle size, shape, and aggregation. Packing density ranged from 80.3 ± 2.7 mg/cm³ (mOFM) to 484.7 ± 17.8 mg/cm³ (pRC). Thermal analysis demonstrated the native structure of the OFM-based test articles (T_m, 59.80 ± 0.11°C and 58.15 ± 0.15°C) relative to pUBM and pRC (T_m, 41.06 ± 0.06°C and 40.59 ± 0.23°C). Rheological testing revealed that mOFM and mOFMµ had increased cohesive energy, indicating better mechanical resilience when the micronized materials were rehydrated to form a paste. The OFM-based test articles exhibited the greatest resistance to proteolytic digestion (T_1/2, 12.730 ± 1.232 and 5.759 ± 0.1296). All the test articles, except for the reconstituted collagen product, demonstrated hemostasis in whole blood. Micronized reconstituted collagen showed immediate dissolution and no fluid absorption, hemostasis, or resistance to proteolytic digestion, whereas micronized OFM showed the greatest proteolytic stability and packing density. Substantial differences among the micronized bioscaffolds were revealed from the analysis, most likely due to their different source materials and manufacturing processes. Careful consideration of these parameters is warranted when selecting a micronized product for soft tissue applications.

Impact Statement

This study provides a comprehensive comparative analysis of four commercially available micronized collagen-based bioscaffolds, highlighting substantial differences in their structural and mechanical properties. Our findings suggest that bioscaffolds derived from the native extracellular matrix, particularly ovine forestomach matrix, exhibit superior proteolytic stability, mechanical cohesion, and hemostasis compared with the products made from reconstituted collagen. These differences are likely driven by the source material and processing method and underscore the critical importance of material selection in clinical applications. This work advances the understanding of bioscaffold performance and informs evidence-based decision-making.

Keywords

micronized extracellular matrix ovine forestomach matrix urinary bladder matrix collagen

Introduction

While collagen-based bioscaffolds presented as sheet-based grafts have been commercially available for many decades, there is an emergence of micronized or powdered collagen-based bioscaffold materials for applications in wound healing and soft tissue repair procedures.¹ These micronized presentations of bioscaffolds offer many advantages over sheet-based grafts, including ease of use, improved apposition to irregular wound surfaces, and rapid delivery of biological components. Similar to sheet-based bioscaffolds, the primary purpose of these micronized products is to serve as a scaffold for rapid cell infiltration and proliferation, which may be improved given the large surface area of these micronized products in comparison with sheet-based grafts.

Micronized collagen-based bioscaffolds can be generally categorized into either reconstituted collagen scaffolds or decellularized extracellular matrix (dECM) bioscaffolds.² Reconstituted collagen bioscaffolds are prepared from collagen that has been extracted under relatively harsh conditions to solubilize the collagen, typically as mixtures of collagen I and III. Soluble or “hydrolyzed” collagen solutions are then reconstituted into solid particles via freeze drying, spray drying, or other similar processing. Whereas, dECM bioscaffolds are derived from a suitable xenogeneic or allogenic tissue that is decellularized leaving only the ECM.^3,4 A key feature of these dECM bioscaffolds is the retention of naturally occurring ECM-associated proteins that are believed to contribute to regenerative processes, as suggested by previous research.^5–10 For example, prior studies have shown that dECM-based bioscaffolds can stimulate cell migration, angiogenesis, and vascularogenesis, leading to the restoration of functional tissue with reduced scaring.^11–13 Whereas, reconstituted collagen and synthetic bioscaffolds have been reported to be relatively biologically inert.^5,14–19

Sheet-based bioscaffolds that incorporate reconstituted collagen have a long history of clinical use dating back to early research in tissue engineering by Yannis and Burke.^20,21 Since then, many reconstituted collagen-based bioscaffolds have entered clinical practice for the treatment of various dermal defects, including burns,²² complex traumatic wounds,^23,24 tumor excisions,²⁵ scar revisions,²⁶ donor sites,²⁷ and chronic wounds.^28,29 Similarly, there has been the widespread adoption of dECM bioscaffolds, including porcine urinary bladder matrix (UBM) and ovine forestomach matrix (OFM), for both implant procedures^30–33 as well as dermal regeneration and wound healing.^34–40

While these types of products have been extensively characterized as sheet-based grafts, little has been presented in the scientific literature to describe the structural features of these products when micronized. The following study was undertaken using standardized testing to benchmark four micronized collagen-based bioscaffolds through structural and functional material analysis, with the aim of informing their clinical use and guiding future biological evaluation.

Materials and Methods

General

The test articles included micronized coarse OFM (“mOFM,” Myriad Morcells™, Aroa Biosurgery Limited, Auckland, New Zealand), micronized fine OFM (“mOFMµ,” Myriad Morcells–Fine™, Aroa Biosurgery Limited), micronized reconstituted bovine collagen (“pRC,” CellerateRX®, Sanara MedTech Inc., Texas, USA/Wound Care Innovations LLC, Texas, USA/Applied Nutritionals LLC, Pennsylvania, USA), and micronized UBM (“pUBM,” MicroMatrix®, Integra Lifesciences, New Jersey, USA), which were obtained from commercial sources. Human whole blood was provided by the New Zealand Blood Service (ethics approval number 2023/13; expiry, 12 July, 2025; facility code, N00764). Descriptive statistics (median, interquartile range, mean, standard error of the mean) were calculated using GraphPad Prism (version 9.0.0, GraphPad Software LLC, USA). Significance between the groups was determined using unpaired one-way analysis of variance (ANOVA) (Tukey’s multiple comparison test). A p value of <0.05 was considered statistically significant.

Particle size analysis—laser diffraction

The test articles were immersed in hexane prior to testing to avoid sample swelling and reduce aggregation. Samples were briefly sonicated prior to testing. Laser diffraction analysis was conducted using a laser diffractometer (Malvern Instruments, Malvern, UK), size range: 10 mm–10 nm, using a silica reference (refractive index, 1.544; absorption index, 0.1). The results were reported as volume mean diameter (D[4,3]); D10, D50, and D90, defined as the particle size where 10%, 50%, and 90% of the particles in the sample are smaller than the measured size, respectively. Span, a measure of particle size distribution, was determined from the following formula:

Span = \frac{D 90 - D 10}{D 50}

Scanning electron microscopy

The test articles were placed onto a carbon disc and distributed using a spatula. The sample height was calibrated, and the samples and metal stud were placed into a tabletop scanning electron microscopy (SEM) instrument (TM3030Plus, Hitachi, Japan). Images were captured in the secondary electron mode, with 15 kV accelerated voltage and 50× and 250× magnification.

Micro-computed tomography

The test articles mOFM, mOFMµ, and pUBM were mounted using 4.5-mm plastic straws, while pRC was mounted in a 3.0-mm plastic straw. The difference in the straw diameter between samples was necessary due to the relatively small particle size and high packing density of pRC. These differences do not impact the comparative analysis via micro-computed tomography (micro-CT), as differences in the straw diameter are accounted for during image acquisition. Data acquisition was performed using a Bruker Skyscan 1272 micro-CT instrument (Bruker, USA) with the following parameters: voltage = 54 kV, current = 200 mA, exposure time = 4000 ms. Images were captured at 4904 × 3280 pixels with at a 1-µm camera pixel resolution, and data were acquired every 0.175° for 360° of rotation. The micro-CT data were reconstructed to give grayscale transaxial planes using InstaRecon CPR Premium 15 K (InstaRecon Inc., USA) software. The micro-CT data were reconstructed to give grayscale transaxial planes at 1-µm intervals using InstaRecon and Nrecon (Bruker, Belgium) software. Representative two-dimensional (2D) images were prepared in ImageJ (version 1.54f, NIH, USA). First, the grayscale images were converted into binary black/white images using the thresholding operation. Black/white images were processed to render a particle-filled image, whereby “pores” within each of the particles were filled to create a space-filled rendering of each particle in each 2D image frame. The test articles were additionally visualized in three dimensions (3D) using CTVox (version 3.3, Bruker, Belgium).

Packing density

The test articles were added to a preweighed microcentrifuge tube ( $m_{initial}$ ) and centrifuged (5 s) using a mini-centrifuge to settle the material within the tube. The test articles were added until 1 mL of volume was reached, and the mass was then recorded ( $m_{final}$ ). Testing was conducted in triplicate. Packing density was expressed in mg/cm³, according to the following equation:

Packing density (m g per {c m}^{3}) = \frac{(m_{final}) - (m_{initial})}{1 {c m}^{3}}

Differential scanning calorimetry

Testing was conducted as previously described, whereby linear sections of the thermogram on either side of the thermal event peak were selected to calculate the melt onset temperature (T_m, °C).⁴¹ Controls included OFM and ovine forestomach tissue (OFT), prepared as previously described.⁴¹

Rheometry

Testing was conducted using a Modular Compact Rheometer (Model # MCR 702e, Anton Paar, Germany) in the parallel plate configuration. The plate diameter was 25 mm. Samples of mOFM, mOFMµ, and pUBM were rehydrated using reverse-osmosis (RO) water at a ratio of 1:5 (g/mL), while pRC was rehydrated 1:1 (g/mL) prior to testing. Test gaps (plate-to-plate separation distance) of 1, 2, and 2.5 mm were used for pRC, mOFMµ and pUBM, and mOFM, respectively, aiming to maintain a consistent normal force of 2 N prior to measurement. The test environment was uncontrolled, at ambient temperature (approximately 21°C). An oscillatory strain sweep test was conducted from 0.01% to 100% strain at a constant angular frequency of 1 Hz.

The storage modulus (G’) and loss modulus (G’’) (GPa) were plotted versus the % strain curves. From the plots of G’ versus % strain, the linear viscoelastic region (LVR) was determined for each test article, and G’ at the LVR (G’_LVR) was determined. Critical strain (ϒ_crit) was determined as the intersection of linear tangents fitted to the LVR (Newtonian plateau) and linear non-Newtonian region. Cohesive energy E_coh⁴² was determined for each test article based on the G’_LVR and ϒ_crit, and E_coh was determined for each test article from triplicate experiments using the following formula:

E_{coh} = \frac{1}{2} {G'}_{L V R} {¡_{crit}}^{2}

Agarose gel electrophoresis

The test articles (50 mg) were papain digested prior to agarose gel electrophoresis, as previously described.⁴³

Fluid absorbency

The test articles (100 mg) were added to a microcentrifuge tube, and the mass was recorded ( $m_{initial}$ ). Phosphate buffered saline (PBS) (2 mL) was added to the sample, and the sample was incubated (37°C, 10 min), with gentle agitation. Samples were centrifuged (∼20,000 g, 10 min), the supernatant was aspirated, and the tube was wicked dry. The mass of the tube was recorded ( $m_{final}$ ). Samples were tested in triplicate, and fluid absorbency was expressed as a percentage of the starting mass ( $m_{initial}$ ) according to the following equation⁴⁴:

Fluid absorbency (%) = \frac{(m_{final}) - (m_{initial})}{(m_{initial})}

Proteolytic stability

Proteolytic stability was quantified for each test article (∼20 mg) in the presence of collagenase (Sigma-Aldrich, USA) solution (1 mL, 50 µg/mL), as previously described.⁴⁵ Samples were tested in triplicate experiments, using quadruplicate samples of each test article. The proteolytic half-life (T_1/2) was determined from interpolation of the linear quadratic regressions at 50% mass remaining.

Blood clotting index

Testing was conducted according to the method previously described.⁴⁶ The test articles (50 mg) were weighed into a microcentrifuge tube and incubated with PBS (1 mL) for 10 min at room temperature (RT) on the orbital shaker. A fresh, empty microcentrifuge tube was also incubated with PBS (1 mL) for 10 min at RT on the orbital shaker as the control. Samples were then centrifugated (7000 rpm, 5 s), and PBS was removed from the samples and control tubes. Human blood (50 µL) was added to all tubes and incubated for 15 min at RT with shaking. After incubation, RO water (950 µL) was added to all the tubes, and samples were incubated for a further 5 min at RT with shaking to lyse the unbound red blood cells and release hemoglobin. Samples were then centrifugated (7000 rpm, 5 s). The hemoglobin solution was transferred to a fresh microcentrifuge tube and diluted 1:10 with RO water; then, 200 µL was transferred to a nontreated 96-well microtiter plate. Absorbance by optical density (OD) was measured at 540 nm (Fluostar Omega, BMG Labtech, Germany) using a microtiter plate reader. Testing was conducted using triplicate samples. Blood clotting index (BCI%) was expressed as the mean from three independent experiments, where each experiment used different donor blood. BCI% was calculated using the formula, where hemoglobin binding (“clotting”) is inversely proportional to BCI%:

BCI (%) = (\frac{O D Sample}{O D Control}) \times 100 %

Results

Particle size analysis—laser diffraction

Particle size distributions of the test articles as a function of size interval are presented in Figure 1. Volume mean diameter (D[4,3]), an estimate of the average particle size, was 825.88, 271.07, 210.91, and 119.23 µm for mOFM, pUBM, mOFMµ, and pRC, respectively (Table 1). D90, the particle size of which 90% of the test particles are smaller than the measured, followed a similar trend, with 90% of mOFM, pUBM, mOFMµ, and pRC being smaller than 1364.55, 430.55, 350.37, and 225.42 µm, respectively. The particle size distribution for each of the test articles was determined by calculating the sample span, from D90, D50, and D10 (Table 1). A larger span value indicates a wider particle size distribution, meaning that there is a greater difference between the largest and smallest particles in the sample. The particle size distribution was the greatest in mOFMµ (span = 2.02) and the lowest in mUBM (span = 1.13).

FIG. 1.

Representative particle size distribution plots from laser diffraction, smoothed plots of particle size (µm) versus % volume by interval.

Table 1.

Results Summary

	mOFM	mOFMµ	pRC	pUBM
Laser diffraction particle size
D[4,3] (µm)	825.88	210.91	119.23	271.07
D10 (µm)	388.81	49.37	30.22	142.66
D50 (µm)	774.57	148.98	106.93	255.76
D90 (µm)	1364.55	350.37	225.42	430.55
Span	1.26	2.02	1.83	1.13
Packing density (mg/cm³) (n = 3)	80.3 ± 2.7	75.4 ± 3.5	484.7 ± 17.8	127.6 ± 3.1
Onset melt temperature (T_m), mean ± SD (°C) (n)	59.80 ± 0.11 (18)	58.15 ± 0.15 (19)	40.59 ± 0.23 (3)	41.06 ± 0.06 (3)
% Fluid absorbency, mean ± SD (n = 3)	551.0 ± 39.3	442.0 ± 40.4	0.0	384.0 ± 12.7
Proteolytic stability (T_1/2), mean ± SD (days) (n = 12)	12.730 ± 1.232	5.759 ± 0.1296	0.003 ± 0.001	2.861 ± 0.127
Blood clotting index (BCI%), mean ± SD (n = 3)	68.01 ± 24.38	52.59 ± 20.69	97.17 ± 4.51	61.54 ± 23.48
Rheometry
Modulus, G’_LVR (Pa), mean ± SD (n = 3)	8951 ± 824	16251 ± 5397	ND	11477 ± 975
Critical strain, ϒ_crit (%), mean ± SD (n = 3)	10.10 ± 0.01	11.83 ± 1.50	ND	3.06 ± 1.00
Cohesive energy, E_coh, (J/m³) mean ± SD (n = 3)	45.7 ± 4.2	109.4 ± 17.7	ND	5.9 ± 3.9

n, sample size; SD, standard deviation; ND, not determined.

Scanning electron microscopy

Representative SEM images are provided in Figure 2. mOFM (Fig. 2A and E) was observed as distinct amorphous-shaped particles, with sizes ranging from approximately 0.5 to 2.0 mm in diameter. mOFMµ (Fig. 2B and F) showed smaller amorphous particles (less than approximately 1.0 mm in diameter), with filamentous surface projections (Fig. 2F). pRC (Fig. 2C and G) was observed as spherical particles that were relatively uniform in appearance when compared with mOFM, mOFMu, and pUBM. Particles of pUBM (Fig. 2D and H) had an amorphous shape but appeared flatter than mOFM particles, with a maximum diameter of 300 μm.

FIG. 2.

Representative SEM images at 50× (scale bar = 2 mm) (A–D) and 250× (scale bar = 300 µm) (E–H) of the test articles mOFM (A, E), mOFMµ (B, F), pRC (C, G), and pUBM (D, H). SEM, scanning electron microscopy; mOFM, ovine forestomach matrix; pRC, reconstituted collagen; pUBM, urinary bladder matrix.

Micro-computed tomography

Representative 2D images from micro-CT for each of the test articles are provided in Figure 3A–D. Both mOFM and mOFMµ (Fig. 3A and B, respectively) had irregular particle geometries with surface projections that had been visualized via SEM. pRC (Fig. 3C) particles were relatively uniform spheres, and where image processing had not entirely filled a particle, a hollow center could be seen. pUBM had rod-like particles (Fig. 3D). Similar morphology was also observed in the 3D images of the test articles (Fig. 3E–H). Most notable was the particle arrangement and 3D packing of pRC (Fig. 3G). The 3D arrangement of pRC contrasted with that of mOFM and mOFMµ, which were more open, with voids between adjacent particles.

FIG. 3.

Representative two-dimensional (scale bar = 1 mm) (A–D) and three-dimensional (E–H) micro-CT images of the test articles mOFM (A, E), mOFMµ (B, F), pRC (C, G), and pUBM (D, H). Note that the test article pRC (C) was imaged using a 3.0-mm straw; thus, the cross-section appears reduced versus test articles presented in panels (A), (B), and (D). micro-CT, micro-computed tomography.

Packing density

Packing density was determined based on the mass (mg) required for each of the test articles to occupy a fixed volume (i.e., 1 mL or 1 cm³). There was no significant difference in the packing density of mOFM and mOFMµ (80.3 ± 2.7 and 75.4 ± 3.5 mg/cm³, respectively), and both OFM-based products had a packing density that was significantly smaller relative to pUBM (127.6 ± 3.1 mg/cm³) (Table 1 and Fig. 4A). The packing density of pRC (484.7 ± 17.8 mg/cm³) was approximately 5-fold higher than the other test articles.

FIG. 4.

(A) Packing density. Error bars represent the standard error of the mean from triplicate samples. (B) Representative DSC thermograms, and (C) onset melt temperature. Error bars represent the standard error of the mean from replicate samples as follows: OFT, n = 30; OFM, n = 60; mOFM, n = 18; mOFMµ, n = 19; pRC, n = 3; pUBM, n = 3. Significance between the groups was determined using unpaired one-way ANOVA (Tukey’s multiple comparison test), where ns = p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. DSC, differential scanning calorimetry.

Differential scanning calorimetry

Representative thermograms for each of the test articles are presented in Figure 4B. Thermograms for the reference samples OFT and OFM are provided as controls to evaluate the impact of processing methods. OFT represents the unprocessed ovine source tissue used to manufacture OFM, mOFM, and mOFMµ. OFM is a sheet form of decellularized ECM prior to morcellation to prepare mOFM and mOFMµ. The test articles mOFM and mOFMµ gave well-defined melt onset temperatures (T_m), similar to OFT and OFM. In contrast, pRC and pUBM thermograms were shallow during the melt transition event, with little heat flux or step-change in the pre/post melt heat capacity. Both mOFM (59.80 ± 0.11°C) and mOFMµ (58.15 ± 0.15°C) had a lower T_m relative to OFT (65.45 ± 0.14°C)⁴¹ and OFM (61.60 ± 0.07°C),⁴¹ reflecting the processing of the source tissue and subsequent morcellation (Fig. 4C and Table 1). Both ovine test articles had a significantly increased T_m relative to pRC (40.59 ± 0.23°C) and pUBM (41.06 ± 0.06°C). There was no significant difference between the T_m of pRC and pUBM (Fig. 4C).

Rheometry

Representative modulus G’ and G’’ versus % strain curves are presented in Figure 5. Rehydrated samples of mOFM, mOFMµ, and pUBM behaved as viscoelastic solids with defined solid/liquid transitions, G’>G’’, and a defined LVR (Fig. 5A,B and 5D). In contrast, pRC did not show a LVR, but instead the soluble material showed viscous flow (Fig. 5C). As pRC did not behave as a viscoelastic solid, LVR could not be determined, and therefore, pRC was not analyzed further. The mean G’_LVR values for mOFM, mOFMµ, and pUBM were 8951 ± 824, 16251 ± 5397, and 11477 ± 975 Pa, respectively (Table 1), but there were no significant differences in G’_LVR (Fig. 5E). The mean critical strain (ϒ_crit) was significantly reduced in pUBM (3.06 ± 1.00%) relative to mOFM (10.10 ± 0.01%) and mOFMµ (11.83 ± 1.50%) (Table 1 and Fig. 5F). Similarly, the mean cohesive energy (E_coh) of pUBM (5.9 ± 3.9 J/m³) was significantly reduced relative to mOFM (45.7 ± 4.2 J/m³) and mOFMµ (109.4 ± 17.7 J/m³) (Table 1), and differences were significant between mOFM and mOFMµ (Fig. 5G).

FIG. 5.

Rheometry analysis. Representative modulus (Pa) versus % strain curves for mOFM (A), mOFMµ (B), pRC (C), and pUBM (D), where G’ and G’’ are plotted with a solid and dashed lines, respectively. For each representative plot, linear tangents were fitted to the LVR (Newtonian plateau) and the linear non-Newtonian region, where the intercept represents ϒ_crit (dotted vertical line). (E) Modulus, G’_LVR. Errors represent the standard error of the mean from triplicate samples. (F) Critical strain, ϒ_crit. Errors represent the standard error of the mean from triplicate samples. (G) Cohesive energy, E_coh. Errors represent the standard error of the mean from triplicate samples. Significance between groups was determined using unpaired one-way ANOVA (Tukey’s multiple comparison test), where ns = p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

Agarose gel electrophoresis

The nucleic acid content of the test articles was qualitatively assessed via agarose gel electrophoresis (Fig. 6). As expected, nuclear material was not detected in the reconstituted collagen product, pRC. Faint nuclear banding (∼0.1 to 0.5 kbp) was present in both mOFM and mOFMµ, while the sample of pUBM gave intense bands from ∼0.1 to 3 kbp.

FIG. 6.

Agarose gel electrophoresis. M = marker, where 10, 3, 1, 0.5, 0.1 kbp.

Fluid absorbency

The reconstituted collagen sample, pRC, was found to be water soluble. As such, percent fluid absorbency was determined to be 0.0% based on the method employed (Table 1 and Fig. 7A). The percent fluid absorbency of mOFM and mOFMµ was 551.0 ± 39.3% and 442.0 ± 40.4%, respectively, and was significantly greater than pUBM (384.0 ± 12.7%).

FIG. 7.

(A) Percentage fluid absorbency. Error bars represent the standard error of the mean from triplicate samples. (B) Proteolytic stability. Error bars represent the standard deviation of the mean from triplicate experiments. Line of best fit determined using a linear quadratic regression. Dotted line at 50% weight remaining used to determine the half-life (T_1/2) for each test article. Dashed line (pRC) represents an estimate of the digestion curve. (C) Proteolytic half-life (T_1/2) determined from interpolation of the linear quadratic regressions (B). Error bars represent the standard error of the mean from triplicate experiments. Significance between groups was determined using unpaired one-way ANOVA (Tukey’s multiple comparison test), where ns = p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

Proteolytic stability

The stability of the test articles in the presence of digestive proteases was assessed via collagenase incubation. The mass of the remaining sample was assessed at multiple time points to derive a stability curve of mass versus time (Fig. 7B), and from this, the proteolytic half-life (T_1/2) of the test article was determined from interpolation of the linear regressions (Table 1, Fig. 7C). As noted previously, pRC was found to be a water-soluble formulation, and as such, T_1/2 (T_1/2 = 0.003 ± 0.001 days) could only be estimated from the digestion curve (Fig. 7B) but is likely to be significantly less than this estimate. The proteolytic stability of mOFM (T_1/2 = 12.730 ± 1.232 days) was approximately twice that of mOFMµ (T_1/2 = 5.759 ± 0.1296 days) and three times that of pUBM (T_1/2 = 2.861 ± 0.127 days) (Table 1, Fig. 7C).

Whole blood clotting

BCI% reflects the relative amount of unbound hemoglobin, where the lower the BCI%, the higher the relative blood clotting for a given test article. There were no significant differences in the BCI% value for mOFM, mOFMµ, and pUBM (68.01 ± 24.38%, 52.59 ± 20.69%, and 61.54 ± 23.48%, respectively) (Table 1, Fig. 8A). pRC did not appreciably bind hemoglobin, and the BCI% value (97.17 ± 4.51%) was significantly higher than the other test articles. Representative images of the test articles after incubation with whole blood are provided in Figure 8B.

FIG. 8.

(A) Quantification of mean BCI%. Error bars represent the standard deviation from three independent experiments. Significance between groups was determined using unpaired one-way ANOVA (Tukey’s multiple comparison test), where ns = p > 0.05, * = p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. (B) Representative images of the test articles after 15-min incubation with whole blood. Co = control.

Discussion

Collagen-based bioscaffolds for soft tissue repair and regeneration are now an established part of the reconstructive ladder.⁴⁷ While these products were originally introduced into clinical practice as sheet-based grafts,²¹ there are now a growing number of micronized collagen-based scaffolds available for clinical use (Supplementary Table S1). However, there is little presented in the literature describing the comparative characterization of these micronized collagen-based bioscaffolds. For the current analysis, we included both a micronized reconstituted collagen (pRC) and representative micronized dECMs (mOFM, mOFMu, and pUBM).

There was diversity in particle size, morphology, and packing density between the test articles, as assessed via laser diffraction (Table 1 and Fig. 1), SEM (Fig. 2), micro-CT (Fig. 3), and packing density (Fig. 4A). The particle size of pRC (D[4,3] = 119.23 µm) was significantly smaller than the dECM test articles (D[4,3] = 210.91 to 825.88 µm), and as a consequence, packing density was significantly higher (Fig. 4A). In clinical terms, this means that approximately five times the mass of pRC would be required to achieve the same coverage or volumetric fill achieved with one of the dECM products. Visual inspection of the pRC particles by SEM and micro-CT showed a relatively uniform spherical appearance, with hollow voids (Fig. 3C), suggesting that pRC is prepared via spray drying aqueous solutions of solubilized collagen.^48,49 The particle sizes (D[4,3]) of the dECM test articles mOFMµ, pUBM, and mOFM were 210.91, 271.07, and 825.88 µm (Table 1), respectively. Our results for the particle size of commercial pUBM are slightly reduced relative to prior reports for pUBM prepared in the laboratory.⁵⁰ There were noticeable differences in the morphology of micronized pUBM and OFM (mOFMµ and mOFM) by SEM (Fig. 2) and micro-CT (Fig. 3). pUBM is manufactured to a micronized format from intact sheets of UBM using the process of cryo-milling,⁵⁰ yielding irregular and relatively solid flakes, consistent with prior reports.^5,50 In contrast, micronized OFM was highly amorphous with irregular fibrous surface projections. These surface projections tended to restrict the close packing of the particles, as evidenced by the reduced packing density of mOFM and mOFMµ (Fig. 4A). Differences in the morphology of mOFM/mOFMµ versus pUBM particles may be due to the differences in the dECM, from which the micronized materials are produced (OFM and UBM), and/or the micronization processes. For example, it is known that OFM has a dense fibrillar collagen architecture⁵¹ and is both thicker and stronger than UBM.⁵² We speculate that milling OFM to create particles may lead to “shredding” of the material that gives rise to the observed filamentous surface projections. This is in contrast to the cryo-milling technique used to produce pUBM,⁵⁰ which potentially “shatters” UBM to yield particles with relatively defined edges. We additionally observed marked differences in the extent of decellularization between the dECM test articles (Fig. 6), consistent with the prior comparative analyses of the decellularization of OFM and UBM.⁴³

The most notable difference between the reconstituted collagen product (pRC) and the dECM test articles were differences in the melt onset temperature (Fig. 4C), fluid absorbency (Fig. 7A), and resistance to proteolytic digestion (Fig. 7C). Reconstituted collagen bioscaffolds are prepared from animal tissues (e.g., dermis or tendon) using relatively harsh chemical processes to extract or solubilize the collagen fibers via hydrolysis. The relatively harsh chemical reagents used to solubilize collagen cause protein denaturation,⁵³ resulting in a reduction in the melt onset temperature (T_m). The reduced T_m seen here for pRC (Table 1) is consistent with prior studies comparing T_m between reconstituted collagen and dECM bioscaffolds.⁴¹ The melt onset temperature (Tm), as derived from differential scanning calorimetry (DSC), is an established analytical method that is highly sensitive to changes in the protein structure resulting from processing.^54,55 The processing steps (e.g., terminal sterilization, temperature cycles, exposure to chemicals including detergents and disinfectants) may denature protein components, which reduce their stability and result in a lower T_m.⁵⁶ In the current study, unprocessed animal tissue (OFT, Fig. 4B and C) that had not been exposed to potentially damaging processes was included as a reference for evaluating the degree of protein denaturation in the test samples. These findings (Fig. 4) are consistent with previous studies showing that reduced T_m is a direct indicator of protein denaturation induced by processing.^41,56–58

Unlike the dECM test articles, pRC was soluble in aqueous solutions and showed no fluid absorbency. The immediate solubilization and lack of structural cohesiveness of pRC particles may account for the relatively high rates of postoperative complications seen with this material.^59,60 Another consequence of any denaturation of the matrix proteins is the relative resistance of the test articles to proteolytic digestion (Fig. 7B). Tissue proteases, including matrix metalloproteinases and neutrophil elastase, play a critical role in tissue remodeling and soft tissue regeneration.^61–63 In the context of collagen-based bioscaffolds, endogenous tissue proteases remodel a bioscaffold, in parallel to host cell infiltration and repopulation. Manufacturing processes that damage or denature proteins within the bioscaffold increase their susceptibility to proteolytic digestion and reduce the bioscaffold’s persistence and efficacy during soft tissue regeneration. Clinically, this may translate to requiring repeat applications of the bioscaffold to achieve the desired reconstructive effect. The mean T_m of pUBM was significantly reduced relative to mOFMµ (41.06 ± 0.06°C and 58.15 ± 0.15°C, Table 1). This was consistent with previous reports⁵ and suggests the denaturation of the proteinaceous matrix of pUBM. Correspondingly, pUBM had significantly reduced proteolytic stability (T_1/2) (2.861 ± 0.127 days) relative to mOFMµ (5.759 ± 0.1296 days), even though the mean particle size of mOFMµ (D[4,3] = 210.91 µm) was smaller than pUBM (D[4,3] = 271.07 µm). Our findings herein are supported by clinical studies describing the need for repeat applications of pUBM,^64–66 whereas micronized OFM typically requires only a single application.^38–40

The surfaces of soft tissue defects are characterized by surface irregularities, tunnelling within the soft tissue, and undermining between tissue planes. Micronized collagen-based bioscaffolds have found a clinical niche in addressing surface irregularities. Micronized collagen-based bioscaffolds are often rehydrated in sterile saline (or wound exudate) to create a paste that can be packed into the irregularities of the soft tissue defect. Rheology was used to assess the viscoelastic properties of the test articles and gauge the useability of the test articles in clinical applications.⁵ A reduced rehydration ratio of 1:1 was used to produce the most concentrated preparation possible for pRC. However, even this more concentrated solution did not result in a viscoelastic hydrogel, and the sample lacked measurable viscoelastic properties under rheological testing (Fig. 5C). This is in contrast to the dECMs that maintained mechanical integrity after rehydration. pUBM demonstrated a significantly lower mean cohesive energy (5.9 ± 3.9 J/m³) relative to the OFM test articles. In practical terms, this means that a hydrated paste of pUBM will disaggregate with less energy relative to micronized OFM. These differences will have an impact on the handling of these materials during placement as well as their resistance to shear forces due to patient movement.

The improved viscoelastic properties of the micronized OFM test articles may be attributed to the irregular fibrous projections observed in micronized OFM. We speculate that these morphological features may enable entanglement between adjacent particles that would lead to the increased cohesive energy of the resulting hydrated pastes. Blood clotting is known to be influenced by surface morphology, and several studies have shown that the clotting efficiency of fibrous surfaces is increased relative to smooth surfaces.^67,68 Given the observed differences in the surface features of OFM particles compared with pRC and pUBM, we characterized the BCI% value for each of the test articles. All materials were rehydrated in sterile saline prior to testing, consistent with the clinical practice for product preparation immediately before application. This approach was intended to simulate real-world conditions under which the materials are deployed clinically, thereby enhancing the translational relevance. In this study, pRC was essentially unable to clot human blood, while hemostasis with the dECM test articles was greatly increased, and approaching significance for mOFMµ (Fig. 8A).

This study was intentionally focused on in vitro characterization to establish foundational material comparisons. While in vitro analyses are valuable for characterizing the material properties and baseline performance of bioscaffolds, they cannot fully replicate the complex, dynamic environment of living tissue. Specifically, processes such as immune cell recruitment, vascularization, and cellular recruitment and remodeling are not captured here. As such, there are limitations in extrapolating these results directly to in vivo tissue remodeling outcomes. However, the performance differences observed here, including thermal stability, clotting behavior, and surface morphology, provide meaningful indicators of scaffold persistence, integration potential, and usability, such as the number of reapplications required in a clinical context. Future studies will be needed to assess in vivo degradation kinetics, host immune responses, and regenerative efficacy in relevant preclinical models.

In summary, while all four test articles are routinely used clinically for soft tissue reconstruction and wound healing, significant differences exist between the micronized collagen-based bioscaffolds. As a result of processing methodology, pRC lacks structural cohesiveness and proteolytic stability, which could limit its functional durability in vivo. Of the micronized dECM bioscaffolds, the OFM-based test articles showed less protein denaturation, reduced DNA content, and increased resistance to proteolytic digestion compared with pUBM. Additionally, the unique surface morphology of micronized OFM gives rise to the differences in the packing density, cohesive energy, and blood clotting. Performance differences observed herein may help predict bioscaffold persistence and usability in a clinical setting.

Authors’ Contributions

S.G.D. and B.C.H.M.: Conceptualization. M.J.S., A.D., A.W., D.G., S.K., Y.N., X.Y., and R.W.F.V.: Methodology. M.J.S., A.D., A.W., D.G., S.K., Y.N., X.Y., H.Y., N.T., D.G., and R.W.F.V.: Investigation. S.G.D., B.C.H.M., M.J.S., S.K., Y.N., and D.G.: Visualization. S.G.D.: Writing—original draft. S.G.D. and B.C.H.M.: Writing—review and editing. S.G.D., I.T.T.M., and B.C.H.M.: Supervision. S.G.D. and B.C.H.M.: Project administration. B.C.H.M.: Funding acquisition.

Footnotes

Acknowledgments

The authors wish to acknowledge the staff and donors of the New Zealand Blood Service.

Funding Information

Research funding was provided by Aroa Biosurgery Limited (Auckland, New Zealand) and Callaghan Innovation (Wellington, New Zealand).

Disclosure Statement

The following authors are employees of Aroa Biosurgery Limited (Auckland, New Zealand): S.G.D., M.J.S., A.D., A.W., S.K., Y.N., X.Y., H.Y., N.T., I.T.T.M., and B.C.H.M.

Supplemental Material

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