Quantitative Evaluation of the In vivo Biocompatibility and Performance of Freeze-cast Tissue Scaffolds

Abstract

Little used for the in vivo assessment of tissue scaffolds are quantitative methods for the evaluation of biocompatibility. To complement current histological techniques, we introduce as a measure of biocompatibility a straightforward, geometric analysis for the quantitative assessment of encapsulation thickness, cross-sectional area, and shape of biomaterials. Advantages of this new technique are that it enables, on the one hand, a more complete and objective comparison of scaffolds with differing compositions, architectures, and mechanical properties, and, on the other, a more objective approach to their selection for a given application. In this contribution, we focus on freeze-cast polymeric scaffolds for tissue regeneration and their subcutaneous implantation in mice for biocompatibility testing. Initially, seven different scaffold types are screened. Of these, three are selected for systematic biocompatibility studies based on histopathological criteria: EDC-NHS-crosslinked bovine collagen, EDC-NHS-crosslinked bovine collagen-nanocellulose, and chitin. Geometric models developed to quantify scaffold size, ovalization, and encapsulation thickness are tested, evaluated, and found to be a powerful and objective metric for the in vivo assessment of biocompatibility and performance of tissue scaffolds.

1. Introduction

Surprisingly few in vivo studies of freeze-cast biopolymer scaffolds have been reported, to date, despite their promise for applications in tissue regeneration thanks to their porosity and complex hierarchical architectures, which parallel those found in native tissues. Freeze-cast scaffolds have the desired high specific surface area and additionally been shown to promote cell and tissue integration while maintaining structural integrity [1-11]. Scaffold structure, mechanical properties, and degradation rate can be custom-designed through material composition, processing parameters (e.g. the freezing rate), and the degree of cross-linking [5,6,10,12-18] In addition to a robust architecture, freeze-cast biopolymer scaffolds offer attractive mechanical and chemical cues that invite cell and tissue integration in vivo with minimal inflammatory and fibrotic responses; additionally, they have desirable biodegradation and bioresorbability profiles [19,20]. Various forms of proteins and polysaccharides such as collagen, chitin, and their derivatives, such as gelatin and chitosan, respectively, stand out as particularly promising biomaterials for a broad range of medical applications [10,12,13,21-25]. Recently and despite its resistance to proteolytic breakdown in its native form, plant-based nanocellulose has been explored for biomaterial applications, because of its attractive mechanical properties, [2,26,27].

Although considerable progress has been made in recent years, the need for new and improved biomaterials and scaffolds persists, because tissue regeneration and repair remain suboptimal in the case of many applications, such as osteogenesis [28-31], cartilage regeneration [32,33], and nerve repair [3,6,7,34,35], and the frequently desired release of drugs and growth factors [36]. Only few in vivo studies have been reported, to date, for highly porous, freeze-cast biomaterials. As a result, little is known about the fundamental structural tissue and immune responses that follow the biomaterial’s implantation [37,38] and the type and magnitude of the foreign body response [39-42] that it invokes, which traditionally are evaluated and scored through qualitative histological examination by a blinded pathologist.

Of great promise is a quantitative analysis and a histomorphometric approach for the assessment of cross-sectional area and shape of biomaterials after implantation [43] with the goal of a more comprehensive, biomaterial evaluation and a more objective scaffold-to-scaffold comparison. Overall, current methods for quantitative biocompatibility include encapsulation thickness measurements, leukocyte cell counts and density, lymphocyte assays, cell infiltration distance, capillary counting, and histological scoring (e.g. scales from 0 to 3 or 0 to 5) systems [20,35,44-47].

Described in this contribution is the development and application of new quantitative metrics in a subcutaneous murine model to characterize both the in vivo foreign body response and structural performance of freeze-cast biopolymer scaffolds, which are suitable to serve as regenerative devices for a wide range of biomedical applications.

2. Materials and Methods

2.1. Fabrication

2.1.1. Scaffold Composition and Slurry Preparation

Seven different scaffold types were manufactured: (A) 1% w/v uncrosslinked bovine collagen (Worthington Biochemical Corporation, Lakewood, NJ), (B) 1% w/v EDC-NHS crosslinked bovine collagen, (C) 1% EDC-NHS crosslinked bovine collagen and nanocellulose blend, (D) 2% nanocellulose blend of Bioplus™ nanocellulose fibrils and crystals (American Process, Atlanta, GA), (E) crosslinked jellyfish collagen (Jellagen, Cardiff, Wales), (F) 0.6% Chitin (Berglund Lab, Wallenberg Wood Science Center, KTH, Stockholm, Sweden), and (G) 1% xanthan gum – konjac glucomannan (Modernist Pantry, Portsmouth, NH).

Water-based slurries were prepared through the stepwise additions of powders and/or stock solutions before homogenizing these with a Fisher Scientific™ Homogenizer 152 (Fisher Scientific International, Inc., Hampton, NH) and/or shear mixing them for 2–3 minutes at 2100-2500 rpm with a Speed Mixer™ (DAC 150FVZ-K, FlackTek, Landrum, SC) to achieve homogeneity and uniform viscosity. The nanocellulose slurry was prepared by diluting the stock suspension with distilled water followed by shear mixing. The 1% w/v collagen suspensions were prepared by raising 1 g collagen powder to 100 mL of 0.05 M acetic acid, refrigerating the heterogeneous mixture overnight, and homogenizing the mixture thoroughly for 1.5 hours at (¾ maximum rpm). The 50:50 (v:v) collagen-nanocellulose blend was made by shear mixing equal volumes of 1% suspensions of each component. Xanthan gum and konjac glucomannan powders in a xanthan:konjac mass ratio of 3:2 were dissolved in double distilled water with 0.02% sodium azide (Sigma-Aldrich, St. Louis, MO ) and buffered to a pH of 5 with citric acid and sodium hydroxide to obtain a concentration of 1% w/v [48]. Chitin was used as received (Berglund Lab, Wallenberg Wood Science Center, KTH, Stockholm, Sweden).

2.1.2. Scaffold Freeze casting and Lyophilization

Slurries were injected with a needle and syringe (16^1/2 gauge) into cylindrical bores of 4 mm diameter and 40 mm length in a cylindrical aluminum mold of 25.4 mm diameter and 45 mm overall length. The bottom of the mold was sealed with a removable Teflon™ cap. After filling the mold with a given slurry composition, it was placed on a copper coldfinger and equilibrated to 4°C before a 10°C/min cooling rate was applied to the copper mold bottom until a temperature of −150°C was reached. Careful control of the applied cooling rate was achieved with a standard freeze-casting system [17] that uses liquid nitrogen to cool the copper cold fingers and a band heater-PID-controller system to ensure desired processing conditions.

Once the samples were frozen, the molds were removed from the cold finger and equilibrated to −20°C in a freezer prior to sample punch out from the mold by hand with a multi-pronged plunger. Samples were placed in a Freezone 6 Plus Lyopholizer (Labconco, Kansas City, MO) at 0.008 mBar and a cooling coil temperature of −85°C for at least 36 hours to remove the ice phase by sublimation.

Xanthan gum – konjac glucomannan (XK) scaffolds were fabricated with an intermediate gelation step prior to freeze casting. The XK solution was sealed in a container and placed in a water bath at 85°C for two hours for mechanical crosslinking. Once the solution had cooled to room temperature it was injected into the aluminum mold to form a semi-solid gel before starting the freezing process.

2.1.3. Scaffold Crosslinking

For crosslinking, the freeze-dried, freeze-cast collagen and collagen-nanocellulose scaffolds were fully submersed and gently stirred for 6 hours at room temperature in a solution of 33 mM 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) and 6 mM N-hydroxysuccinimide (NHS) (both Sigma-Aldrich, St. Louis, MO) in 200 proof ethanol as the solvent. To remove residual crosslinking agents, the crosslinked scaffolds were submersed and washed three times stirring them in fresh distilled water for 2 hours, 12 hours, and 1 hour, respectively. In a final processing step, the fully hydrated, crosslinked scaffolds were flash frozen in liquid nitrogen and lyophilized a second time under the same freeze-drying conditions.

2.2. Structural Characterization

The scaffold microstructure was quantified using scanning electron microscopy (SEM). All SEM micrographs were acquired with a Tescan VEGA 3 scanning electron microscope (Tescan, Brno, Czech Republic). Scale bars were added using the ImageJ software.

2.3. In vivo Biocompatibility

2.3.1. Surgery and Implantation

Scaffolds were sectioned into 6 mm long cylinders, massed, and sterilized with ethylene oxide gas under vacuum for 24 hours (12 hours sterilization followed by 12 hours outgassing) at 22°C before subcutaneous implantation in mice.

The mice (C3H, Charles River Laboratories, Wilmington, MA) were three-month-old (21-22 grams). Before surgery, all mice received 0.9 mL of ketoprofen/saline cocktail with 0.1 mg/mL ketoprofen, and were anesthetized under vaporized isoflurane.

The lower body of the mice was shaved, and a one-centimeter transverse incision was made in the side body wall. For scaffold placement, the scaffold was loaded into a tapered rubber catheter (inner diameter > 4 mm) and inserted into the surgical pocket. The implant was deposited by inserting a rubber plunger and slowly retracting the catheter. Proline suture (6-0) was used to close the incision. As control, the operation was repeated on both flanks of mice exactly as described above without a scaffold.

Post-anesthesia, three sequential antiseptic scrubs with chlorohexidine, ethanol, and betadine were performed on every surgical site. Less than 24 hours post-surgery, 0.9 mL of ketoprofen/saline cocktail was administered. Ketaprofen was used to manage pain in the short-term and is not expected to affect foreign body response outcomes.

Sutures were removed 7 days post-surgery. Animals were sacrificed two weeks post-surgery; blood was drawn directly from the heart and stored in 0.5 mL EDTA (purple top) tubes. All cadavers were fixed in 10% formalin post-sacrifice for at least 24 hours before the left and right body wall containing the implants were resected from each mouse with surgical scissors. Multiple measures were taken to ensure histological sections of the cylindrical implants were transverse to the cylindrical axis when analyzed under a microscope: i) at surgery, scaffolds were all implanted perpendicular to the transverse/axial body plane of the mouse; this ensured similar tissue contact/orientation to the scaffold for all implanted scaffolds; ii) upon sacrifice, a square was cut from the body wall/flank that contained the scaffold; the implant could easily be distinguished from the surrounding tissue by visual inspection of the underside of the skin during dissection; thus, the orientation of the cylindrical scaffold was clearly visible; iii) to ensure that cutting was uniform and precise, given the pliable and slippery nature of the scaffold and tissue even after formalin fixation, samples were processed in paraffin and eosin. This enabled clean and uniform sectioning of the scaffold with a known orientation. Each processed body wall sample was thus sectioned transversely, blocked in paraffin, thinly (4 μm) sliced with a microtome, and stained with hematoxylin and eosin (H&E).

All animals in this study were handled as approved by the Dartmouth IACUC and all surgeries were performed in compliance with the NIH Guide for Care and Use of Laboratory Animals.

2.3.2. Histology

Histological slides were viewed with an Olympus BX50 transmission light microscope (Olympus Corporation, Tokyo, Japan) at 2x, 4x, and 40x magnification and using its SPOT software. Scale bars were added and images quantified using the ImageJ software. Measurements were taken on 1600 pixel by 1200 pixel images; 272 pixels corresponded to 1 mm. High magnification micrographs of the implants were evaluated qualitatively to assess the type of foreign body response to the scaffold implant.

2.3.3. Quantification of Geometry and Size of Encapsulation

The novelty of this approach for assessing in vivo biocompatibility of freeze-cast scaffolds is the quantification of both the implant geometry and the thickness of encapsulation as a straightforward indicator of the severity of the foreign body response. Histological slides reveal the shape of the deformed, but originally circular face of the cylindrical scaffolds sectioned transverse to the cylinder axis.

For quantification of the local foreign body encapsulation, two parameters were determined on n = 9-10 samples of each scaffold type: i) the area of the scaffold, A_S, and ii) the area of the scaffold plus the encapsulation area, A_SC. Margins that demarcated A_SC were determined by a pathologist based on the morphology of the capsule. While staining can vary, these measurements were taken based on morphology of the encapsulation, and tinctorial quality and density (i.e. staining properties) were used for validation. The area of the capsule A_C was determined as the difference of A_SC and A_S (Eq. 1).

$$A_C=A_{SC}-A_S$$ (Eq. 1)

The area ratio A_ratio is defined by Eq. 2:

$$A_{\mathit{ratio}}=\frac{A_{SC}}{A_S}$$ (Eq. 2)

Additionally, the long axis, L_axis, and, orthogonally to it, the short axis, S_axis, of the samples that ovalized in vivo, were measured on each implanted scaffold. Long and short axis were measured manually using ImageJ. Axes were defined as both passing through the center of the scaffold section and being perpendicular (at 90°) to one another, without first fitting an oval. Their ratio is calculated:

$$R_{\mathit{axis}}=\frac{L_{\mathit{axis}}}{S_{\mathit{axis}}}$$ (Eq. 3)

The closer the ratio to unity, the more circular the cross section remained in vivo; the larger the ratio, the more ovalized the cross section is. Lastly, the thickness of the encapsulation layer, t_exp, around each implant was measured to quantify the local response. This is achieved by averaging eight measurements taken every 45° around the histological scaffold section, as shown in Figure 1. An angle of 45° (approximated with respect to the long axis of the scaffold to ensure that the line of measurement was perpendicular to the tangent of the scaffold) is then used to determine capsule thickness t₃ t₄ t₅ and t₆.

Figure 1.

Histological schematic and definition of parameters for which measurements were taken: the area of the scaffold, A_S; the area of the capsule A_C:; the scaffold long axis, L_axis, and short axis, S_axis; the average encapsulation thickness, t_exp.

2.4. Mathematical Models of Scaffold Encapsulation

Three geometric models (Figure 2) were developed (as detailed in the Appendix), for comparison to find the most suitable to quantify the degree of scaffold encapsulation correctly by i) describing the encapsulation thickness and ii) determining a normalized thickness parameter (Figure 2).

Figure 2.

Three different geometric models to calculate encapsulation thickness from measured axes and area parameters.

In Model A, the capsule thickness, t_shell, is calculated from the measurement of the scaffold plus capsule area, A_SC, and scaffold only area, A_S, assuming a circular cross-section and using a thin shell approximation (Eqs 4, 5).

In Model B, the capsule thickness, t_circle, is calculated from the measurement of the scaffold radius, r, and the scaffold plus capsule area, A_SC, assuming a circular cross-section (Eqs 6, 7).

In Model C, the capsule thickness, t_ellipse, is calculated from the measurements of the scaffold’s long and short axes, L_axis, and S_axis,, and the scaffold plus capsule area, A_SC, assuming an elliptical cross-section (Eqs 8, 9).

The area of the scaffold, A_S, and the area of the scaffold plus capsule, A_SC as well as the long, L_axis, and short, S_axis, axes of the ovalized implant could easily and precisely be measured on histological sections. Implant radius, r, was calculated from A_S (Eq. 10), assuming a circular scaffold cross-section:

$$r=\sqrt{\frac{A_S}{\pi }}$$ (Eq. 10)

To approximate an ellipse model for scaffold cross-section, semi-major/minor axes, a and b, were calculated from A_S and R_axis following Eqs 11 & 12:

$$\frac{a}{b}=R_{axis}$$ (Eq. 11)

$$A_S=\pi ab$$ (Eq. 12)

Finally, normalized biocompatibility indices F_A, F_B, and F_C were calculated to describe the scaffold area with respect to the gross amount of encapsulation: the higher the index, the smaller the relative encapsulation, the better the biocompatibility.

2.5. Systemic Immune Response

The systemic immune response was evaluated based on a complete white blood cell (WBC) count performed by ANTECH Diagnostics after arrival within 24-36 hours of sampling. Sample size of n=4-5 were used.

2.6. Statistical Methods

Statistical analyses were performed with MATLAB (R2016B) and Microsoft Excel (2016). Assuming a Gaussian distribution of results, small sample two-tail (equal variance) hypothesis t-tests were performed comparing mean white blood cell count (WBC) of the treatment groups to the control. One-way Analysis of Variance (ANOVA) was performed to compare all three biocompatibility response parameters (A_S, A_SC, and A_ratio) for the three treatment groups. Small sample one-sided hypothesis t-tests were performed to compare the axis ratio (R_axis) for the three treatment groups. Paired t-tests were performed to compare each estimated encapsulation thickness parameter (t_shell, t_circle, and t_ellipse) to the experimentally measured t_exp. Calculations for error progression were performed with Maple™ 18 (Maplesoft, division of Waterloo Maple Inc., Waterloo, Ontario) and Microsoft Excel (2016).

3. Results

3.1. Qualitative Assessments of Histological Sections

The seven scaffold types were studied histologically to characterize the composition and structure of the foreign body response. Different capsule thicknesses (Figure 3 & 4) formed, reflecting the varying severity of the reaction. The biomaterial-tissue interface for each scaffold type at a higher magnification (Figure 4) revealed that the encapsulation was made of fibrillar layers of organized and/or unorganized fibroblasts, natively deposited collagen, and immune cells. Results are summarized in Table 1.

Figure 3.

Histological sections (2 weeks) of the implanted scaffolds. (A) uncrosslinked bovine collagen; (B) crosslinked bovine collagen; (C) crosslinked bovine collagen-nanocellulose blend (D) pure nanocellulose blend; (E) crosslinked jellyfish collagen; (F) chitin; (G) xanthan-konjac glucomannan; (H) control.

Figure 4.

High magnification histological sections (2 weeks) of implanted scaffolds; labelled are organized fibroblasts (OF), unorganized fibroblasts (UF), enlarged fibroblasts (EF), neutrophilic region (N), adipose cells (AC), and the scaffold (S) material. A) uncrosslinked bovine collagen; (B) crosslinked bovine collagen; (C) crosslinked bovine collagen-nanocellulose blend (D) pure nanocellulose blend; (E) crosslinked jellyfish collagen; (F) chitin; (G) xanthan-konjac glucomannan; (H) control.

Table 1.

Foreign Body Response Histological Assessment Summary from 2 week Scaffold Screening Study

	Scaffold Type	Scaffold Appearance	Conserved Porosity	Scaffold Shape	Capsule Composition	Immune Cell Infiltration
(A)	Uncrosslinked Collagen	Highly compacted pink fibrillar material	No	Ovalized	Outer layer of organized fibroblasts; inner layer of unorganized fibroblasts and immune cells	Throughout scaffold material
(B)	Crosslinked Collagen	Pink porous macrostructure	Yes	Ovalized	Organized fibroblasts	Minimal
(C)	Crosslinked Collagen- Nanocellulose	Purple porous macrostructure	Yes	Circular	Organized fibroblast exterior and cellular interior	Extending from capsule into scaffold
(D)	Nanocellulose	Compacted light purple fibrillar material	No	Ovalized	External layer of organized fibroblasts and dense interior of neutrophils and polymorphonuclear cells	Minimal
(E)	Crosslinked Jellyfish Collagen	Highly compacted dark purple fibrillar material	No	Ovalized	Outer layer of organized fibroblasts; inner layer of unorganized fibroblasts and immune cells	Throughout scaffold material
(F)	Chitin	Pink porous fibrillar structure; some vascularization	Yes	Ovalized	Organized fibroblasts	Minimal
(G)	Xanthan-konjac glucomannan	Compacted purple hyaline material with dehydrated gellike appearance	No	Ovalized	Pyogranulomatous response; thick and dense capsule with immune cells, giant cells, and enlarged fibroblasts.	Extensive

3.2. Systematic Study of Three Scaffold Compositions

Based on the screening of seven different scaffold types, crosslinked bovine collagen (Col), crosslinked bovine collagen-nanocellulose (Col-NB), and chitin (Chi) were downselected as promising materials for a systematic in vivo study to quantitatively compare local foreign body encapsulations and morphologies (Table 1).

3.3. Systemic Response

Statistical analysis of the WBC counts for Col, Col-NB, and Chi compared to the control WBC count indicated that the difference in the mean WBC count for the control group and the tested biomaterial was not statistically significant (Table 2).

Table 2.

White blood cell count (mean ± SD) for Col (n = 5), Col-NB (n= 4), and Chi (n = 4) as compared to the control (n = 5) after 2 weeks

Scaffold Type	WBC (103/μL)	t statistic	p value
Control	1.8 ± 0.49	-	-
Collagen	2.9 ± 0.94	−2.197	0.0592
Collagen- Nanocellulose	2.7 ± 1.2	−1.412	0.2008
Chitin	1.7 ± 0.83	0.4311	0.6794

3.4. Experimental Results for Scaffold Encapsulation

Col-NB best preserved its size and shape and had both the largest scaffold (A_S) and scaffold plus capsule areas (A_SC), followed by Chi and Col. The capsule thickness decreased from Col-NB to Col to Chi; however, A_ratio decreased from Col to Col-NB to Chi (Figure 5). Measurements of the long, L_axis, and short, S_axis, cross-sectional axes of the implanted scaffolds and their ratios (Figure 6) revealed that the crosslinked Col-NB had the axis ratio, R_axis, closest to 1, indicating a more circular cross-section than all others, and one that differed significantly (p < 0.001) from chitin. In contrast, the R_axis values did not differ significantly from one another (p > 0.50) for chitin and crosslinked collagen.

Figure 5.

Boxplots for local and structural responses (2 weeks) for crosslinked bovine collagen (Col, n = 10), crosslinked collagen-nanocellulose (Col-NB, n = 10), and chitin (Chi, n = 9) scaffolds after two weeks in vivo.

Figure 6.

SEM micrographs of transverse cross-sections (left) and scaffold surface morphologies at low (middle) and high magnification (right) of (A) crosslinked bovine collagen, (B) crosslinked bovine collagen – nanocellulose, and (C) Chitin.

3.5. Results Comparison of Models A, B and C

To determine, whether the capsule thickness, t_exp, can appropriately be approximated from area measurements using Models A (shell) and B (circle), and area and axis measurements for Model C (ellipse), experimentally determined thickness values were compared with calculated ones (t_shell, t_circle, and t_ellipse) obtained using the geometric models (Table 3).

Table 3.

Scaffold encapsulation thickness (mean ± SD) values for experimental and geometric models.

	Scaffold Encapsulation Thickness (μm)
	texp	tellipse	tcircle	tshell
Col (n = 10)	107 ± 29.4	102 ± 31.8	110 ± 32.0	117 ± 35.5
Col-NB (n = 10)	115 ± 37.5	119 ± 33.4	121 ± 33.1	128 ± 36.3
Chi (n = 9)	87.9 ± 28.2	85.5 ± 37.9	94.9 ± 42.3	99.9 ± 47.5

Error progression analysis for the calculation of the three thickness parameters calculated with the models for the three materials, indicated that the t_ellipse exhibits errors (±12.1-20.2) that are more than twice that of either t_circle (±5.92-7.88) or t_shell (±6.62-9.12). t_ellipse was the most accurate (Table 3) as a predictor of t_exp. The shell Model A thickness, t_shell was least accurate compared to t_exp (Table 3); model A differed significantly for Col-NB (p = 0.0273). Both the circle Model B and ellipse Model C were more accurate in predicting encapsulation thickness values that agreed well with the experimental values, t_exp, for all three materials.

Lastly, the encapsulation thickness indices, F_B and F_C, for circular or ellipsoidal geometry, respectively, which correlate scaffold size with capsule thickness shape were applied to Col, Col-NB, and Chi for a comparison in their accuracy and indicated trends similar to those of the area ratios, as expected (Table 4). F_A was not considered given the significant difference between t_shell and t_exp. Chi had the highest index of 13.8 (F_B) and 6.86 (F_C), suggesting it is the most biocompatible and invokes the lowest foreign body response. Col had the lowest indices of 9.44 (F_B) and 4.69 (F_C); lastly, Col-NB had indices of 10.9 (F_B) and 5.45 (F_C).

Table 4.

Foreign body response indices (mean ± SD) for circle and ellipse models.

Col (n = 10)	Col-NB (n = 10)	Chitin (n = 9)
Circle, $F_B:\frac{r}{t_{\mathit{circle}}}$
9.44 ± 3.46	10.9 ± 4.03	13.8 ± 6.09
Ellipse, $F_C:\frac{ab}{(a+b)t_{\mathit{ellipse}}}$
4.69 ± 1.75	5.45 ± 2.01	6.86 ± 3.05

Structural analysis of the pore morphology of the three candidate materials suggested that Col possesses a more radially oriented porosity with more film-like and closed cell walls than the other two compositions (Figure 6). Additionally, the surface of the scaffold was formed by a skin-like outer layer that has a small amount of porosity and very small pore diameters. Col-NB and chitin did not exhibit radial porosity; instead a porosity gradient was apparent with pore diameters increasing from the perimeter to the center of the scaffold (Figure 6). Overall, Col possessed the largest pores with long and short axes of 275 μm and 47.8 μm, respectively, followed by Col-NB (80 μm; 63.3 μm) and Chi (70.2 μm; 57.3 μm). Both Col-NB and Chi were lacking the skin observed on Col (Figure 6).

A direct comparison of SEM micrographs of the respective scaffolds to histological images of the scaffolds in vivo revealed that the original shape, morphology and cell wall structure of the pores were well preserved, while there was shrinkage due to scaffold ovalization under subcutaneous compression and sample processing (Figure 7).

Figure 7.

A comparison of transverse histological sections (2 weeks) of (A) crosslinked bovine collagen scaffolds, (B) crosslinked bovine collagen–nanocellulose, (C) chitin with corresponding transverse SEM micrographs of dry scaffolds pre-implantation (right). Note, in all cases, the in vivo conserved porosity and microstructures in comparison to the scaffold structure pre-implantation.

4. Discussion

Considerable differences in the local foreign body responses were observed when evaluating in vivo seven freeze-cast scaffold types. While collagen [1,9] jellyfish collagen [20,44], plant-derived nanocellulose [27] and chitin [49] had been tested in vivo prior to this study, collagen-nanocellulose composites [2], and xanthan gum–konjac glucomannan have not been reported in vivo.

It is well-known that adjusting processing conditions for a given material can greatly change structure (pore size, pore aspect ratio, cell wall thickness) and mechanical properties (modulus, yield strength, and toughness) [5] and thus the resulting in vivo performance. Hence, this study focuses on the use of quantitative techniques to characterize histomorphometric outcomes, after first qualitatively evaluating histopathology, rather than making absolute claims into the etiology of one response compared to another.

4.1. Screening Study of Seven Scaffold Compositions

The in vivo performance of uncrosslinked bovine collagen, crosslinked bovine collagen, a crosslinked bovine Col-NB composite, nanocellulose, crosslinked jellyfish collagen, Chi, and, a xanthan-konjac glucomannan composite scaffolds differed considerably, because of differences in composition, structure, and mechanical properties.

Low mechanical properties of the uncrosslinked bovine collagen scaffold resulted in its severe compaction and considerable loss of porosity, when compressed in vivo. Scaffold biodegradation likely elicited a further foreign body response with a thick encapsulation composed of a combination of fibroblasts, native collagen deposition, and immune cells. The considerable encapsulation and lack of structural integrity made this scaffold unattractive for further analysis.

Collagen is known to be proteolytically degraded and remodeled by the body. Subsequent crosslinking of the collagen is known to inhibit this degradation and reduce cell infiltration; in addition to peptide bond formation, the crosslinking eliminates free lysine, asparagine, and glutamine residues, which reduce protein and cell binding [10,12,50,51]. In the freeze-cast scaffold, the paper-like cell-wall structure was likely reinforced with covalent bonding between the collagen fibrils. Thus, the chemical crosslinking, which results in higher stiffness, strength, and resilience, explain Col’s well preserved macro- and microstructural features as well its lower foreign body response when compared to its uncrosslinked counterpart.

The Col-NB composite best retained its shape and porosity in vivo. The presence of stiff and strong fibrillar nanocellulose, enhanced the mechanical properties of the cell-wall material in comparison to the pure collagen [26]. In contrast, pure nanocellulose scaffolds lacked the benefit of a cross-linked composite cell wall material, and thus lost their pore structure and ovalized, despite a higher overall density; the duration of implantation is expected to result in the abundance of polymorphonuclear cells [27]. A neutrophilic response may be further upregulated by the inability for proteolytic degradation of cellulose molecules. Thus, the hybrid response in the Col-NB of both fibroblasts, immune cells, and increased neutrophils compared to pure collagen scaffolds is thought to be a result of its semi-degradable composition.

While it is known that EDC-NHS crosslinker chemistry does function effectively for marine collagen derived from jellyfish [20,52,53], the evaluation of Type II jellyfish collagen in vivo revealed that the crosslinking process did not structurally and mechanically enhance the material to the same extent as was observed in the case of Type I bovine collagen and in fact, looked structurally similar to the uncrosslinked bovine collagen. Potential reasons for ineffective chemical crosslinking include, differences in the degree of crosslinking in the stock solid, the collagen fibrillation, and/or the crosslinking chemistry. Further studies are required to explore these hypotheses in detail.

The least encapsulation was observed for the Chi scaffold, which also exhibited well-conserved micro-porosity. In the absence of crosslinking or structural reinforcement through composite formation some macro-structural compression was observed albeit less than expected, given its lower solid content, which was almost half that of the collagen-based scaffolds. Also, the interfacial response was minimal, suggesting that this scaffold has promise.

Xanthan-konjac glucomannan scaffolds were the least attractive of all. The lack of retained porosity and the presence of a very dense capsule of inflammatory cells rendered them unsuitable in their current form. The concentric white space in the histological section and the highly compacted and hyaline appearance of the scaffold material mimics the histological behavior of serous fluid or exudate, suggesting that the scaffolds likely absorbed large amounts of abscess-like fluid and dehydrated during histological processing. The encapsulation of the implant appeared inhomogeneous and varied in its cellular makeup, both of which are undesirable characteristics. The innate composition and lack of purity in the components could account for the poor outcome.

4.2. Systematic Study of Col, Col-NB, and Chi

Col, Col-NB, and Chi were selected for our systematic in vivo biocompatibility study, because of their promising in vivo performance. Comparing the scaffolds’ in vitro with their in vivo structures (Figure 7), all compositions were found to preserve the freeze-cast porosity and pore morphology well. However, there was evidence of pore compression that accompanied the overall scaffold ovalization. Given the noticeable differences in scaffold shape for the different compositions, this response reveals the difficulty in knowing a scaffold’s true in vivo porosity, P_IV, given its compressed cross-sectional scaffold area A_IV, in comparison to the porosity P₀ and cross-sectional are A₀ before implantation.

We propose to address this gap in the following way. Based on the conventional equation for scaffold porosity as a function of the densities of the scaffold ρ₀ and that of the solid material ρ_s from which it was made:

$$P_0=1-\frac{{\rho }_0}{{\rho }_S}$$ (Eq. 13)

and assuming an original cross-sectional area of the scaffold, A₀, as well as that the scaffold length and mass remain unchanged in this short-term subcutaneous model, and along with these the cell wall thickness, and pore diameter.

With these assumptions and an equivalence between the ratios of the scaffold porosities and cross-sectional areas, we generate a simplified equation for the true scaffold porosity in vivo as a function of experimentally measurable parameters. These parameters are the cross-sectional areas A₀ and A_IV before and after implantation, respectively:

$$\frac{{\rho }_{IV}}{{\rho }_0}=\frac{A_0}{A_{IV}}$$ (Eq. 14)

to calculate with these the porosity retained in vivo:

$$P_{IV}=1-\frac{A_0}{A_{IV}}(1-P_0)$$ (Eq. 15)

where A_IV = A_S (1 + S) with the tissue shrinkage, S, due to processing and sectioning ranges between 5% to 40%, i.e. S takes values from 0.05 to 0.40 [54].

Using this model, we can approximate reductions in scaffolds porosity after implantation (Table 5) and find the overall porosity well preserved, while the shape of individual pores changes.

Table 5.

Scaffold density, porosity, cross-sectional area before/after implantation with/without tissue shrinkage (5-40%), and in vivo porosity

	Density (mg/cm3)	P0 (%)	A0 (mm2)	AS (mm2)	AIV (mm2)	PIV (%)
Col (n=10)	14.0 ± 0.3	98.92 ± 0.02	8.6 ± 0.3	3.1 ± 1.3	3.3 – 4.3	~97-99
Col-NB (n=10)	16.4 ± 0.8	98.80 ± 0.06	7.1 ± 0.8	4.9 ± 1.8	5.1 – 6.9	~98-99
Chi (n=9)	14.1 ± 0.3	99.06 ± 0.01	5.9 ± 0.3	3.9 ± 0.75	4.1 – 5.5	~98-99

Col retained its distinct radial porosity well. Its more film-like cell-wall material structure results from the degree of fibrillation and cross-linking of the collagen used as well as slurry composition, viscosity and pH, combined with the applied processing conditions, such as the application of a radial thermal gradient and freezing rate.

The structure of Col-NB and Chi scaffolds is distinctly different. Both compositions have a more equiaxed pore and considerably more porous cell-wall material structure, which results from the entrapment of the nanofibrils in the ice-phase during freezing. This entrapment not only causes fiber bundles to cross the ice-templated pore space and bridge the cell walls at regular intervals, but also results in a more fibrillar and open cell-wall material structure. Because cell-wall material and bridges become little distinguishable as a result, the overall appearance of the structure is one of an equiaxed foam rather than a more honeycomb-like architecture and pore alignment. The presence of collagen in the collagen-nanocellulose composite makes these effects a little less pronounced than in the case of chitin. The effect of nanofibrillar entrapment is likely also the reason for the absence of a skin layer on the radial surfaces at which freezing starts, resulting in a more open scaffold surface, which may ease cell infiltration and be preferable in some, but not all, biomedical applications.

The presence or absence of this surface skin likely contributes to the differences in cell infiltration observed in this study. Of the three scaffold compositions, the least cellular infiltration was visible in Col, which possesses a scaffold skin that likely impedes cell infiltration. In contrast, Col-NB, a protein-polysaccharide composite, displayed the largest surface porosity and the most cellular infiltration, while Chi, a polysaccharide, exhibited a smaller surface porosity and some immune cell infiltration.

It is important to note that, due to biodegradation, the scaffold morphology is expected to significantly change with time and site of implantation. The rate of scaffold degradation and resulting scaffold structure will depend factors such as composition, pore structure and orientation, cell-wall thickness, and degree of cross-linking, type and distance from the living tissue, and severity of the foreign body response, capsule formation, and the type and degree of cell infiltration.

Our new quantitative method for the histomorphometric evaluation of tissue scaffold biocompatibility uses scaffold and encapsulation dimensions, areas, and their ratios. This approach offers several advantages since it enables gross and relative comparisons between different materials and because the method remains applicable, as we could show, even when an originally round scaffold cross-section deforms and ovalizes in vivo.

When comparing area and axes measurements, it becomes clear that both the overall scaffold area and the axis ratio are important predictors for scaffold success; both are measures of retained overall porosity and individual pore size, and both are affected by in vivo compressive forces, resulting from a combination of subcutaneous compression and compression due to encapsulation. The higher the axis ratio, the lower the overall porosity and individual pore size, because of lower mechanical properties such as stiffness and compressive strength.

The significance of scaffold area and degree of encapsulation depend on a scaffold’s application. For example: while chitin performs best in terms of its relative and absolute biocompatibility, it ovalizes significantly more than collagen-nanocellulose. This ovalization reflects a reduction in pore size and results, as observed, in differences in cell and tissue infiltration into the scaffold.

In contrast, scaffold or total area and area of capsule capture differences in both gross shape and foreign body response. This data addresses biocompatibility as defined by encapsulation thickness. Therefore, axis ratio (a measure of a scaffold’s mechanical properties) and scaffold versus total area (a measure of encapsulation) are both important metrics to quantify biocompatibility and foreign body response.

A careful comparison between experimental encapsulation thickness values and those calculated using the method of simplified geometries (circle, ellipse, thin shell) were found to coincide well for the circle and ellipse models.

Despite differences in cell type, degree of infiltration, and morphological shape, it is important to note that the ratio of scaffold plus encapsulation area to the scaffold area, A_ratio, for the three materials were in a similar value range; this can be interpreted as an indication that all three materials are within the same tolerance for biocompatibility as oppose to the implication that they are indistinguishable; in fact, the Chi clearly had the lowest relative encapsulation while Col had the highest. This trend was also consistent with the normalized thickness indices determined by F_B and F_C using the geometric Models B and C, respectively. Similarly, this trend was consistent with histopathological observation, which also illustrates the same order of severity of the response from Col to Col-NB to Chi.

The gross capsule thickness was actually highest in the Col-NB and lowest in the Chi both qualitatively and based on experimental thickness measurements. This is an important outcome illustrating that normalizing scaffold encapsulation is an important consideration when making material comparisons; traditional gross encapsulation measurements by themselves are not a complete reflection of the biomaterial-tissue interface.

Comparing the different approaches, Model B performs slightly better in cases of a more circular scaffold crosssection, while Model C is more suitable for more ovalized samples; as reflected in the model equations, the primary limitation of Model B is the dependence on a radius to fit non-circular ovalized sections while Model C is limited by the error in the two axis measurements that define ellipse eccentricity. The thin shell circle model (Model A) displayed a high percent error that renders it unattractive for evaluation purposes but useful for a comparison in methodology.

The successful use of geometric models to characterize the biocompatibility of scaffolds in vivo, suggests that a similar approach may be taken to systematically analyze histological data for other in vivo biomaterial-tissue interfaces. Overall, the results of this study illustrate the advantages of this newly developed, straightforward and precise method. With it, the encapsulation thickness can objectively be quantified for implanted tissue scaffolds using area measurements of the scaffold and its surrounding foreign body response.

Conclusions

In vivo biocompatibility testing of biomaterials is a critical step that informs material selection. This paper first surveys a range of both novel and established freeze-cast scaffolds to understand their histopathological responses. Subsequently, a new quantitative histomorphometric approach and geometric models were developed to characterize the biomaterial-tissue interface.

Screening seven compositions, uncrosslinked bovine collagen was found to suffer from compaction, excessive encapsulation, and a loss in microstructural porosity. Jellyfish collagen exhibited a similar response. Nanocellulose lacked structural integrity and displayed a neutrophil-heavy response. Xantham-konjac glucomannan performed poorly with a pyogranulomatous response. Crosslinked bovine collagen, crosslinked bovine collagen-nanocellulose, and chitin scaffolds were selected for a systematic investigation. While all three were highly biocompatible in this in vivo model, the structural and foreign body responses of these scaffold compositions observed on H&E-stained histological cross-sections differed.

A successful quantitative description of the observed encapsulation was found through straightforward models for both circular and elliptical scaffold shapes in vivo, indicating the promise of such a geometric approach. The new quantitative metrics complement well the traditional qualitative histopathological assessments and enable more objective scaffold comparisons. Future work should focus on a systematic investigation of the effects of composition and processing parameters on the response, using the newly developed quantitative framework.

Appendix

A.1. Geometric Models for Foreign Body Encapsulation

A.1.1. Model A: Capsule Thickness Using a Thin Shell Approximation

The thin-shell model assumes that the histological scaffold section can be treated as a circle with radius, r, capsule thickness, t_shell, and a capsule area of:

$$A_C=2\pi r{t}_{shell}$$ (Eq. A1)

Assuming that an error of 10% in the capsule area, A_C, approximation is acceptable—a value that corresponds to the experimental, error, as will be detailed below— this model may be used for implants whose ratio of scaffold radius to capsule thickness, r/t_shell is larger than 4.5, with r being calculated from the experimentally determined scaffold area, A_S following Eq. C2:

$$r=\sqrt{\frac{A_S}{\pi }}$$ (Eq. A2)

The shell thickness can then be calculated from Eq. 3:

$$t_{shell}=\frac{A_C}{2\pi r}$$ (Eq. A3)

A.1.2. Model B: Capsule Thickness Assuming a Circular Scaffold Cross-Section

Assuming that the histological scaffold cross-section can be treated as a circle, the capsule thickness t_circle can be determined directly from the difference between the scaffold plus the encapsulation area, A_SC, area and the scaffold area, A_S, following Eq. 4:

$$t_{\mathit{circle}}=\frac{\sqrt{A_{SC}}-\sqrt{A_S}}{\sqrt{\pi }}$$ (Eq. A4)

A.1.3. Model C: Capsule Thickness Assuming an Ellipsoidal Scaffold Cross-Section

Assuming that the histological scaffold cross-section can be treated as an ellipse with semi-major and semi-minor axes a and b, and an axis ratio

$$\frac{a}{b}=R_{axis}$$ (Eq. A5)

a scaffold area

$$A_S=\pi ab$$ (Eq. A6)

a scaffold plus capsule area with axes that are one capsule thickness t_ellipse greater than that of the scaffold

$$A_{SC}=\pi (a+t_{\mathit{ellipse}})(b+t_{\mathit{ellipse}})$$ (Eq. A7)

a scaffold capsule area that for a, b >> t may be approximated as

$$A_C=\pi (a+b)t_{\mathit{ellipse}}$$ (Eq. A8)

and the capsule thickness can be calculated accurately as

$$t_{\mathit{ellipse}}=\frac{\sqrt{(a-b{)}^2+4A_{SC}∕\pi }-(a+b)}{2}$$ (Eq. A9)

Assuming, as we did for Model A, that an error of 10% in the capsule area, A_C, approximation is acceptable, this Model C may be used for implants whose ratio of the shell thickness to the sum of the long and short ellipse axis (a +b)/t is larger than 9.

For all three models we can define a non-dimensional parameter that allows a comparison to be made between different scaffolds. In the case of Models A and B with a circular geometry, we use as the foreign body index, F_A,B, the ratio of scaffold radius, r, to capsule thickness, t_shell or t_circle.

$$F_{A,B}=\frac{r}{t_{\mathrm{shell},\mathrm{circle}}}$$ (Eq. A10)

In the case of the ellipse, the foreign body index, F_C, is the ratio of scaffold the scaffold to shell areas:

$$F_C=\frac{ab}{(a+b)t_{\mathit{ellipse}}}$$ (Eq. 11)

We will below apply and compare these three different approaches in their accuracy for the description of the different scaffold materials and the foreign body responses that they caused.

A.2. Experimental and Model Results, Statistics, and Errors

Table A1.

Mean ± SD and ANOVA of scaffold + capsule area (A_SC), scaffold area (A_S), and area ratio for collagen (Col), collagen-nanocellulose (Col-NB), and chitin (Chi); the scaffold + capsule and scaffold areas are not similar among the three materials while the area ratios are.

	Scaffold + Capsule Area (mm2)
	Col	Col-NB	Chi
	3.822 ±1.54	5.904 ± 0.693	4.584 ± 1.97
Fstatistic		4.797
pvalue		0.0169
	Scaffold Area (mm2)
	Col	Col-NB	Chi
	3.108 ± 1.32	4.918 ± 1.77	3.903 ± 0.756
Fstatistic		4.438
ftapvalue		0.0220
	ASC to AS Ratio
	Col	Col-NB	Chi
	1.261 ± 0.116	1.218 ± 0.0796	1.186 ± 0.103
Fstatistic		1.370
pvalue		0.2772

Table A2.

Long axis (L_axis), short axis (S_axis), and axis ratio (R_axis) mean ± SD for collagen (Col), collagen nanocellulose blend (Col- NB), and chitin (Chi).

Collagen (mm)
Laxis	Saxis	Raxis
2.896 ± 0.368	1.400 ± 0.533	2.340 ± 1.326
Collagen-Nanoblend (mm)
Laxis	Saxis	Raxis
2.941 ± 0.384	2.096 ± 0.608	1.512 ± 0.443
Chitin (mm)
Laxis	Saxis	Raxis
3.607 ± 0.474	1.392 ± 0.215	2.631 ± 0.400

Table A3.

Unpaired t-test of Axis Ratio (R_axis) for collagen (Col), collagen nanocellulose blend (Col-NB), and Chitin (Chi); Chi and Col-NB are significantly different in their R_axis; Col is similar in shape to both Chi than Col-NB.

	Axis Ratio, Raxis
	Col vs Chi	Col vs Col-NB	Chi vs Col-NB
p value	0.522	0.130	0.000129

Table A4.

Standard error after error progression analysis

(μm)	tellipse	tcircle	tshell
Col	20.2	7.88	9.12
Col-NB	12.1	5.92	7.20
Chi	15.6	6.58	6.62

Table A5.

Paired t-test p values of t_ellipse, t_circle, and t_shell vs. experimental thickness t_exp for collagen (Col), collagen-nanocellulose (Col-NB), and chitin (Chi); t_ellipse and t_circle are statistically similar to t_exp for all three materials, while t_shell is significantly different from t_exp for collagen-nanoblend.

Collagen
tellipse	tcircle	tshell
0.247	0.619	0.149
Collagen-Nanoblend
tellipse	tcircle	tshell
0.393	0.213	0.0273
Chitin
tellipse	tcircle	tshell
0.595	0.273	0.151

Table A6.

Pore size (mean ± SD) for collagen (Col), collagen-nanocellulose (Col-NB), and chitin (Chi)

	Length μm)	Width μm)	Pore Aspect Ratio
Col	275 ± 127	47.8 ± 3.43	5.69 ± 2.48
Col-NB	80.0 ± 30.1	63.3 ± 6.32	1.30 ± 0.325
Chi	70.2 ± 10.2	57.3 ± 14.0	1.31 ± 0.361