Impact of Release Kinetics on Efficacy of Locally Delivered Parathyroid Hormone for Bone Regeneration Applications

Abstract

Characterizing the release profile for materials-directed local delivery of bioactive molecules and its effect on bone regeneration is an important step to improve our understanding of, and ability to optimize, the bone healing response. This study examined the local delivery of parathyroid hormone (PTH) using a thiol-ene hydrogel embedded in a porous poly(propylene fumarate) (PPF) scaffold for bone regeneration applications. The aim of this study was to characterize the degradation-controlled in vitro release kinetics of PTH from the thiol-ene hydrogels, in vivo hydrogel degradation in a subcutaneous implant model, and bone healing in a rat critical size bone defect. Tethering PTH to the hydrogel matrix eliminated the early timepoint burst release that was observed in previous in vitro work where PTH was free to diffuse out of the matrix. Only 8% of the tethered PTH was released from the hydrogel during the first 2 weeks, but by day 21, 80% of the PTH was released, and complete release was achieved by day 28. In vivo implantation revealed that complete degradation of the hydrogel alone occurred by day 21; however, when incorporated in a three-dimensional printed osteoconductive PPF scaffold, the hydrogel persisted for >56 days. Treatment of bone defects with the composite thiol-ene hydrogel–PPF scaffold, delivering either 3 or 10 μg of tethered PTH 1-84, was found to increase bridging of critical size bone defects, whereas treatment with 30 μg of tethered PTH resulted in less bone ingrowth into the defect area. Continued development of this biomaterial delivery system for PTH could lead to improved therapies for treatment of nonunion fractures and critical size bone defects.

Impact statement

Characterizing the release profile for materials-directed local delivery of bioactive molecules and its effect on bone regeneration is an important step to improve our understanding of, and ability to optimize, the bone healing response. The presented work details the effect of modifying the release of parathyroid hormone from a biomaterial scaffold on bone regeneration and preliminary work on the use of noninvasive imaging to monitor in vivo biomaterial scaffold degradation. These data provide a foundation for future studies aimed to understand and optimize the use of bioactive molecule delivery for bone regeneration and tissue engineering applications.

Introduction

Understanding release kinetics and degradation of biomaterial/biomolecule delivery systems is essential for their optimization and use in regenerative medicine. For example, in a bone regeneration model, bone morphogenetic protein 2 (BMP-2) delivered from a material through a burst release, or a burst release followed by sustained release, resulted in more new bone formation than a slow sustained release.¹ However, for novel alternative osteoanabolic molecules for bone regeneration applications, the optimal release profiles are unknown. Because of its anabolic effect on bone, parathyroid hormone (PTH) is a molecule of interest for bone regeneration applications.² When administered as daily injections, PTH treatment results in an increase in bone volume and bone mineral density in women with osteoporosis.^3–6 A number of studies have shown PTH to have beneficial effects on bone healing when given as daily injections.^7–9 Pulsatile release of PTH from biomaterial scaffolds has been investigated for treatment of osteoporosis. However, developing scaffolds for local delivery of PTH for bone regeneration remains relatively unexplored.^10,11

It is generally accepted that continuously elevated PTH has a catabolic effect on bone.^12,13 Cortical bone loss is characteristic in primary hyperparathyroidism; however, trabecular bone is preserved or enhanced.^14–16 Because chronically elevated PTH does not universally have catabolic effects on bone,^14,16 and mild PHPT may be beneficial for trabecular bone, local delivery of PTH for bone regeneration applications may not be limited to an intermittent release profile. Advances in biomaterial delivery systems may provide an opportunity to optimize the local delivery of PTH to promote a bone healing response, while decreasing or eliminating systemic PTH exposure.

Various biomaterial delivery systems, including the system used in this study,¹⁷ indicate local delivery of PTH can promote healing of critical size bone defects.^18–20 PTH delivered by a poly(ethylene glycol) (PEG) matrix resulted in increased bone formation in bone chambers in rabbits, and in cylindrical defects around titanium implants in dog mandibles.^18,21 Similarly, Arrighi et al. observed increased healing in 8 mm diameter cylindrical bone defects treated with PTH delivered by fibrin hydrogel.¹⁹ In vitro release indicated 11.6% of the PTH was released from the fibrin hydrogels during the first 72 h.¹⁹ Previously, we entrapped PTH in a thiol-ene hydrogel allowing it to freely diffuse from the hydrogel–poly(propylene fumarate) (PPF) scaffold system.¹⁷ In vitro release kinetics indicated a burst release, resulting in 80% of the PTH being released in the first 72 h. Treatment of critical size femur defects with the PPF scaffold and thiol-ene hydrogel with 10 μg PTH resulted in improved defect bridging by a combination of bony and cartilaginous tissue.¹⁷ However, complete bony bridging was not achieved in all samples.

Whether or not different release kinetics of PTH from the thiol-ene hydrogel and PPF scaffold would enhance bone regeneration efficacy is unclear and motivated this study. For example, would a PTH burst release of 11.6%, as in the Arrighi study, result in better bone healing than the 80% burst release in our previous study? To address this question we tethered PTH to the hydrogel matrix using an MMP degradable crosslinker, to reduce the burst release and tailor release to local enzyme levels, instead of relying on simple diffusive processes.^22,23 We hypothesized that reducing the burst and prolonging the release of locally delivered PTH using thiol-ene hydrogel and PPF scaffold would improve healing in a rat critical size defect bone model.

Methods

Preparation of PTH 1-84

Black bear PTH 1-84 was used for this study because PTH has been implicated in the mechanism hibernating bears use to prevent disuse osteoporosis.²⁴ It is a potent osteoanabolic agent that reverses osteoporosis in rodent models,²⁵ and can be more osteoanabolic than human PTH.²⁶ Furthermore, we recently used it in a bone defect healing study.¹⁷ Recombinant black bear PTH 1-84 was produced by Proteos (Kalamazoo, MI) and stored lyophilized at −80°C. Before use, a 1 μg/μL stock solution of PTH was prepared in Dulbecco's phosphate-buffered saline (dPBS; Gibco by Life Technologies, Waltham, MA).

To accommodate crosslinking of PTH to the hydrogel matrix, PTH was modified by adding a sulfhydryl group. The sulfhydryl group was added to PTH using Traut's reagent (2-iminothiolane-HCl; ThermoFisher Scientific, Waltham, MA). The 1 μg/μL stock solution of PTH was exposed to a 20 × molar excess of Traut's reagent and 3 mM ethylenediaminetetraacetic acid (EDTA; Sigma-Aldrich, St Louis, MO) and incubated at room temperature for 2 h. Excess Traut's reagent was then removed using a desalting column, and modification was verified by high-performance liquid chromatography (HPLC).

PPF scaffold preparation

PPF was synthesized as previously described.²⁷ Cylindrical scaffolds (external length of 6 mm and diameter of 3 mm) with square pores (1 × 1 mm in size with a strut thickness of 500 μm) were fabricated using a Viper Si2 stereolithography system (3D Systems, Valencia, CA). After fabrication, scaffolds were coated with synthetic bone mineral in a chemical bath using a precipitation type method, as previously described.²⁸ Manufacturing and dimension details are consistent with those used in the previous study where PTH was free to diffuse out of the thiol-ene hydrogels.¹⁷

Thiol-ene hydrogel preparation

Monomer solutions used to make the hydrogels were prepared as previously described.²⁹ In brief, monomer solutions were as follows: 6 wt %/vol 10 K four-arm poly(ethylene glycol)–norbornene (PEG-NB), 1 mM adhesion peptide (CRGDS), 5.5 mM di-cysteine MMP-degradable crosslinker peptide (KCGPQGIAGQCK), 0.01% of the photoinitiator lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP), and PTH in dPBS. The monomer solutions were created using PEG (JenKem, Plano, TX) functionalized with a norbornene group,³⁰ adhesion peptide (CRGDS) and MMP-degradable crosslinker purchased from Bachem (Torrance, CA), and LAP was synthesized as previously described.²⁹ A 1 mL syringe was used as a mold to create 90 μL cylindrical hydrogels (diameter: ∼4 mm, height: ∼6 mm). The PPF scaffold was placed in the mold and the hydrogel solution was pipetted into the mold, filling the pores of the scaffold. The solution was then polymerized around the scaffold under a 2.5 mW/cm² ultraviolet lamp (365 nm) for 2.5 min.

In vitro release kinetics

In vitro release kinetics of PTH 1-84 crosslinked to the hydrogel matrix was determined for hydrogels alone and hydrogels embedded in PPF scaffolds (hydrogels+PPF). Hydrogels (n = 3 per group, per timepoint) containing no PTH (vehicle) or 30 μg PTH 1-84 crosslinked to the gel were placed in 1 mL of dPBS and kept in an incubator at 37°C. PBS (1 mL) was collected at days 3, 7, 14, 21, and 28, and stored at −80°C until the last sample was collected. After all samples were collected, the amount of PTH released from the hydrogels at each timepoint was determined by μBCA protein assay kit (Thermo Scientific, Waltham, MA). The collected supernatant was combined with working reagent and incubated according to kit instructions. Absorbance was measured using plate reader after incubation and compared with a standard curve.

Verification of bioactivity

To verify bioactivity of PTH 1-84 released from hydrogels and hydrogels+PPF, MC3T3-E1 cells were seeded in 6-well plates in basal medium (α-minimum essential medium [Hyclone, Logan, UT], supplemented with 10% fetal bovine serum [Peak Serum, Wellington, CO], and 1% penicillin/streptomycin solution [Hyclone]). After 24 h, the basal medium was changed to osteogenic medium (basal +50 μg/mL l-ascorbic acid +10 mM β-glycerophosphate; day 0). On day 3 the medium was replaced. On day 5 cells were stimulated for 10 min with 1 mL dPBS containing PTH released from hydrogels (collected on days 3 and 7 as described for in vitro release kinetics), or a control treatment as follows: vehicle (dPBS—control), forskolin (Sigma-Aldrich; positive control), 100 nM stock PTH 1-84 solution, and dPBS collected from hydrogels containing no PTH. Cyclic AMP was measured in duplicate for each sample by enzyme-linked immunosorbent assay (Cayman Chemical, Ann Arbor, MI) to determine if the PTH released from the scaffolds was bioactive.

In vivo hydrogel degradation

Thiol-ene hydrogels used for in vivo degradation studies were prepared as described previously, with the substitution of Alexa Fluor 680 (ThermoFisher Scientific) crosslinked to the hydrogel matrix instead of PTH. As a control, ex vivo fluorescent imaging was performed on the PPF scaffolds, thiol-ene hydrogels, thiol-ene hydrogels surrounding PPF scaffolds, and sutures to verify that these materials did not fluoresce at the excitation/emission wavelengths for Alexa Fluor 680 (excitation wavelength: 679 nm and emission wavelength: 702 nm).

All procedures were approved by the Colorado State University Institutional Animal Care and Use Committee (IACUC protocol number 15–555A). Twelve male Sprague-Dawley rats (∼350 g) were anesthetized by isoflurane inhalation (2–3%, to effect). Animals received buprenorphine SR (0.6–0.8 mg/kg) and cefazolin (20 mg/kg) subcutaneously 20–30 min before surgery, for pain management and prophylactic antibiotic, respectively. The hydrogels were implanted subcutaneously over the lateral aspect of the femur. Four rats received Alexa Fluor 680-labeled thiol-ene hydrogels. Eight rats received Alexa Fluor 680-labeled thiol-ene hydrogels embedded in PPF scaffolds.

In vivo fluorescent imaging was conducted out to 56 days. To reduce autofluorescence background the subcutaneous pocket was isolated using black paper as given in Figure 1. The fluorescent label from the hydrogel was detected, integrated, digitized and displayed via an IVIS Spectrum in vivo imaging system (Perkin Elmer, Waltham MA). Fluorescence was analyzed for a 4 mm circular region of interest, centered in the hydrogel area, using Living Image software.

FIG. 1.

(A) In vivo fluorescent imaging of rat with fluorophore-labeled thiol-ene hydrogel. The hydrogels were implanted subcutaneously above the femur as indicated by the white arrow. (B) IVIS imaging of implanted thiol-ene hydrogels isolated using black paper. Color images are available online.

Animals with hydrogels alone were imaged daily until fluorescence from the hydrogel could no longer be detected. When fluorescence from the hydrogel was no longer detectable 2 days in a row, the implant site was opened and examined to qualitatively assess hydrogel degradation. Animals with hydrogels combined with PPF scaffolds were imaged daily for the first 10 days, then every other day until their respective timepoint: days 10 (n = 2), 20 (n = 2), 40 (n = 2), and 56 (n = 2). At each timepoint the implant site was opened and examined to qualitatively assess hydrogel degradation.

Femur defect: surgery

Male Sprague-Dawley rats (340–380 g) underwent surgery to create a 6 mm segmental defect in the midshaft of the femur. Surgeon and animal aseptic preparation before the surgery was according to the Colorado State University IACUC Guidelines for Rodent Survival Surgery (Protocol number 15-6309A). Animals received buprenorphine SR (0.6–0.8 mg/kg) and cefazolin (20 mg/kg) subcutaneously 20–30 min before surgery for pain management and prophylactic antibiotic, respectively. Rats were anesthetized by isoflurane inhalation (2–3%, to effect). The diaphysis of the femur was approached, clearing away soft tissue with a combination of sharp and blunt dissection. A four-pin internal fixation plate was attached to the femur with 0.045 inch (∼1.14 mm) threaded titanium k-wires and a 6 mm defect was created using gigli wire.

After the defect was created, one of five treatments (n = 7/group) was placed in the defect: PPF+thiol-ene hydrogel +0 μg PTH 1–84 (vehicle) containing no PTH, PPF+thiol-ene hydrogel +1 μg PTH 1-84, PPF+thiol-ene hydrogel +3 μg PTH 1-84, PPF+thiol-ene hydrogel +10 μg PTH 1-84, or PPF+thiol-ene hydrogel +30 μg PTH 1-84. Bone healing in the defect volume was monitored using in vivo micro-computed tomography (μCT) scans at 4 and 8 weeks. At the end of the study (12 weeks), animals were killed, femurs were fixed in neutral-buffered formalin for 72 h, and then stored in 70% ethanol, and healing was assessed using ex vivo μCT.

Femur defect: in vivo μCT

Healing was monitored by in vivo μCT (VivaCT 80; Scanco Medical) at weeks 4 and 8. Rats were anesthetized by isoflurane inhalation (2–3%, to effect). The femur was scanned 70 kVp, 114 μA, 8 W, field of view/diameter (mm) of 49.8. Bone volume and mineral content in the defect area were evaluated at each timepoint. Radiographs were used to calculate the percent of the defect area that had bridged with new bone. Percent defect bridged was calculated as original defect size (6 mm) minus the average length between bone ends (measured in mm), divided by original defect size (Equation 1) (Fig. 2).

FIG. 2.

(A) Radiograph of a femur defect at week 8. Yellow measurement lines represent length measurements used to obtain an average “length between bone ends” for Equation (1). (B) μCT scans region (11 mm), the entire defect (6 mm), and the analysis region (3.5 mm). μCT, micro-computed tomography. Color images are available online.

$$\begin{array}{c}\%\mathrm{\hspace{0.17em}d}efect\mathrm{\hspace{0.17em}b}ridged=\left(\frac{6\mathrm{\hspace{0.17em}m}m\hspace{0.17em}-\mathrm{\hspace{0.17em}l}ength\mathrm{\hspace{0.17em}b}etween\mathrm{\hspace{0.17em}b}one\mathrm{\hspace{0.17em}e}nds}{6\mathrm{\hspace{0.17em}m}m}\right)\end{array}\times 100$$ (1)

Femur defect: ex vivo μCT

At 12 weeks bone volume and mineral density were assessed (ex vivo) by μCT (VivaCT 80; Scanco Medical). During scans, samples were held in a 50 mL conical tube surrounded by 70% ethanol. The area between the two inner fixation pins was scanned at 70 kVp, 114 μA, 8 W, FOV/diameter (mm) of 31.5. As there was sufficient healing that native bone ends could not be easily discerned, the volume of the defect that was evaluated was limited to the middle 3.5 mm of the defect area (Fig. 2) for the 4- and 8-week in vivo and the 12-week ex vivo evaluations. The region of interest was evaluated with a lower threshold of 220, gauss support = 1 and sigma = 0.8. The samples were observed for the presence of full bridging. If complete bridging of the defect was achieved for any portion of the defect volume, samples were given a “yes” indication, otherwise they were given a “no” indication.

Femur defect: histomorphometry

Five samples from each group were randomly selected for histological analysis. Samples were embedded in plastic, sectioned, and stained (three thin sections per sample, per stain) with VonKossa MacNeil, Goldners Trichrome, and hematoxylin and eosin. Slides were used to assess whether differences between groups in the type of soft tissue in the defect area were observable. Slides were digitized using a ZEISS Discovery V20 stereoscope and Axiocam 512 color ccd (ZEISS, Oberkochen, Germany). Mineralized tissue area, areas of cartilaginous matrix, and areas of fibrous tissue in the defect were measured on the VonKossa MacNeil slides using the hematoxylin and eosin and Goldner's Trichrome to confirm tissue types. Qualitative observations were also made regarding the type of tissue bridging the defect and healing response. If there was complete bridging of the defect by a combination of bone and cartilage for any portion of the defect area, samples were given a “yes” indication with regard to full bony bridging. All other samples (i.e., samples bridged by a combination of bone and fibrous tissue) were given a “no” indication.

Statistics

One-way analysis of variance (ANOVA) was used to determine differences between treatment groups. If significant differences were detected by ANOVA, Tukey's post hoc test was used compare individual groups to one another. Categorical data from histologically prepared samples were compared using Fisher's exact test. A value of p < 0.05 was considered significant for all tests.

Results

In vitro release

No difference was observed in PTH released from hydrogels with or without PPF scaffolds (p > 0.200), thus data presented are combined. Figure 3 provides the release profile of PTH from thiol-ene hydrogels containing PTH tethered to the matrix. The bulk of PTH was retained at early timepoints, with no observable burst effect and <10% of the PTH being released from the scaffold during the first 2 weeks. However, with substantial degradation at later stages, >80% of the PTH was released by day 21, and complete release by day 28. Complete release of PTH did not correspond to complete degradation of the hydrogel in vitro.

FIG. 3.

(A) In vitro PTH release profile data from thiol-ene hydrogels containing 30 μg PTH tethered to the matrix. (B) PTH released from hydrogels retained bioactivity. Treatment groups were as follows: vehicle (dPBS), forskolin (positive control), PTH control (100 nM PTH 1-84 stock solution), hydrogel + PPF (hydrogel containing no PTH, embedded in PPF scaffold), released PTH (PTH 1-84 collected from release experiments at days 3 [D3] and 7 [D7]). Forskolin, PTH control, and released PTH at days 3 and 7 were all higher than vehicle controls (p < 0.009). Different letters indicate significant difference in cAMP activity between groups. dPBS, Dulbecco's phosphate-buffered saline; PPF, poly(propylene fumarate); PTH, parathyroid hormone.

Verification of PTH bioactivity

As there were no differences in bioactivity of PTH released from hydrogels compared with hydrogels with PPF scaffolds, data were combined. Cyclic AMP activity was higher in the PTH control (100 nM stock solution) and forskolin (positive control) groups, as well as PTH released from both hydrogels and hydrogels embedded in PPF scaffolds than in vehicle (dPBS) and hydrogel controls (hydrogels containing no PTH; p < 0.009) (Fig. 3). Bioactivity of PTH released from hydrogels at days 3 and 7 was comparable with the forskolin and PTH control groups (p = 0.466).

In vivo degradation

Alexa Fluor 680 signal was detectable in subcutaneous hydrogels between days 8 and 21 (Fig. 4A), when fluorescence could no longer be detected in two rats at day 8, one rat at day 18, and one rat at day 21. Over the first 5 days there was an initial decrease in the amount of fluorescence detected followed by a leveling off at an average radiant efficiency of ∼6E7 (p/s/cm²/sr)/(μW/cm²). Detectable fluorescence remained around that level until fluorescence from the hydrogels was no longer detectable. Upon dissection of the subcutaneous pocket where hydrogels were located, no evidence of the thiol-ene hydrogels could be detected in rats killed at day 18 or 21. In the two rats killed at day 8, there was still a small amount of fluid in the pocket that fluoresced when imaged.

FIG. 4.

(A) In vivo fluorescence of subcutaneous thiol-ene hydrogels (without PPF scaffolds) labeled with Alexa Fluor 680. Each line shows data from one rat/implant. (B) In vivo fluorescence of PPF scaffolds and thiol-ene hydrogels labeled with Alexa Fluor 680. Each line shows data from one rat/implant. (C) Longitudinal IVIS imaging of subcutaneous Alexa Fluor 680 labeled thiol-ene hydrogel and PPF scaffold for 8 weeks. (D) Thiol-ene hydrogel surrounding a PPF scaffold in a subcutaneous implant at day 56. Color images are available online.

Alexa Fluor 680-labeled thiol-ene hydrogels embedded in porous PPF scaffolds remained detectable up to 8 weeks (Fig. 4B). Like the Alexa Fluor 680-labeled hydrogels without PPF scaffolds, an initial drop in fluorescence over the first 5 days was followed by a leveling off at an average radiant efficiency of ∼6E7 (p/s/cm²/sr)/(μW/cm²). This fluorescence level persisted until week 8 when the study was terminated. Upon dissection of the subcutaneous pocket, the implanted hydrogel and PPF scaffold were evident at all timepoints examined (Fig. 4D). Qualitatively, at days 10 and 20, the hydrogel and scaffold were easily removed from the pocket in one piece. However, by days 40 and 56, the hydrogel had substantially degraded, losing some of its mechanical integrity, and fell apart upon attempted removal.

In vivo healing efficacy

In vivo μCT and radiograph measurements detected no statistical difference between doses (vehicle, 1, 3, 10, 30 μg PTH) in bone volume, percent defect bridged, or mineral density at weeks 4 or 8 (p > 0.175). However, full bony bridging (Fig. 5A) was achieved in at least one sample in the 1, 10, and 30 μg treatment groups. At week 12 no significant differences between treatment groups were observed in bone volume in the defect area (p = 0.300) (Fig. 5B). Treatment with 3 μg PTH tended to result in more bridging than vehicle (p = 0.092). Treatment with 30 μg PTH tended to result in less bridging than 3 μg (p = 0.075) (Fig. 5A–C).

FIG. 5.

(A) μCT scans showing healing progression over 12 weeks for a rat in the 10 μg PTH treatment group with full bridging by week 12. (B) Bone volume (n = 7 per group) in the center 3.5 mm of the defect volume at 12 weeks was not different between groups (p = 0.300). (C) Percent defect bridged by bone at 12 weeks (n = 7 per group) was not different between groups (p = 0.37).

Histologically, mineralized area, cartilaginous area, and fibrous area were not different between treatment groups (p > 0.200) (Fig. 6A). However, qualitative observations regarding the type of tissue bridging the defect indicated the spatial distribution of the mineralized and cartilaginous area throughout the defect were different; more defects in the 3 and 10 μg treatment groups were bridged by a combination of bone and cartilage than other groups (Fig. 6B, C). Full bridging by combination of mineralized and cartilaginous tissue was different (p = 0.034) between treatment groups: 5/5 samples in the 3 μg PTH group, 3/5 samples in the 10 μg PTH group, and 1/5 samples in the vehicle, 1 μg PTH, and 30 μg PTH groups (Figs. 6C and 7A, B). In samples not bridged by a combination of mineralized and cartilaginous tissue, loosely organized fibrous tissue was a prominent tissue type present in the defect area. Endochondral ossification was observed in 1/5 of the samples in the vehicle group, 2/5 in the 1 μg treatment group, and 3/5 samples in the 3, 10, and 30 μg treatment groups.

FIG. 6.

(A) Percent of the defect area that was mineralized tissue, cartilaginous tissue, and fibrous tissue were not different between treatment groups. (B) Number of samples with complete bridging of the defect with some combination of bone and cartilage (n = 5 per group). (C) VonKossa MacNeil stained longitudinal sections of femur defect area. The PPF scaffold stains dark blue (indicated by yellow arrows), mineralized tissue stains black (indicated by pink arrow), cartilaginous material stains dark blue/purple (indicated by orange arrow), and red arrows indicate areas of loosely connected fibrous tissue. The image on the left is a defect treated with 10 μg bbPTH 1-84 using thiol-ene hydrogel with a PPF scaffold at 12 weeks. Full bridging of the defect area by either mineralized or cartilaginous material is evident. The image on the right is a sample from the vehicle treatment group. The defect was bridged primarily by loosely organized fibrous tissue. Color images are available online.

FIG. 7.

(A) Representative images of defect sections stained with Goldner's Trichrome. The image on the left is a defect treated with 10 μg bbPTH 1-84 using thiol-ene hydrogel with a PPF scaffold at 12 weeks. The image on the right is a sample from the vehicle treatment group. The PPF scaffold shows up white (indicated by yellow arrows), mature bone tissue stains green (indicated by pink arrow), immature bone tissue stains red, and calcified cartilage stain very pales green. Red arrows indicate areas of loosely connected fibrous tissue. Magnified areas are shown to provide an indication of the healing response on a cellular level. (B) Representative images of defect sections stained with hematoxylin and eosin. The image on the left is a defect treated with 10 μg bbPTH 1-84 using thiol-ene hydrogel with a PPF scaffold at 12 weeks. The image on the right is a sample from the vehicle treatment group. The PPF scaffold shows up white (indicated by yellow arrows), mature bone tissue stains dark pink (indicated by pink arrow). Red arrows indicate areas of loosely connected fibrous tissue. Magnified areas are shown to provide an indication of the healing response on a cellular level. Color images are available online.

Discussion

The impact of the release profile of locally delivered PTH from biomaterial scaffolds on the healing response of bone tissue remains unknown. Thus, characterization of the release kinetics, as well as degradation of the materials used to deliver PTH, are integral to understanding and interpreting the healing response in efficacy studies. Previously we characterized the bone healing response elicited when 80% of PTH was released in the first 72 h. The goal of this study was to modify the thiol-ene hydrogel to prevent a PTH burst release and prolong the time-course of PTH release in the defect. This was accomplished by tethering PTH to the hydrogel matrix using an enzymatically cleavable crosslink. The PTH retained bioactivity after being released from the hydrogel. When subcutaneously implanted with a PPF scaffold, the hydrogel persisted for up to 8 weeks.

Though none of the PTH treatment groups was able to consistently promote full defect bridging by bone, the 3 μg PTH group was able to promote complete bridging by a combination of mineralized and cartilaginous tissue. These findings support the idea that thiol-ene hydrogels can be tailored to alter degradability and PTH release profiles for bone regeneration applications. Future work will continue to explore the effects of PTH release profile and concentrations on bone regeneration.

When PTH was free to diffuse out of the thiol-ene hydrogel, the majority (80%) of the PTH was released by 72 h.¹⁷ Tethering PTH to the hydrogel matrix successfully prevented the burst release of PTH (Fig. 3) at early timepoints, and yielded a release profile comparable with that of PTH tethered to the fibrin hydrogel used to treat drill defects in sheep.¹⁹ Tethering PTH to the thiol-ene hydrogel matrix successfully mitigated the rapid release of the majority of PTH until day 14. Between days 14 and 21, 80% of the tethered PTH was released from the hydrogel. Although it occurs before complete reverse gelation, the rapid in vitro release observed between days 14 and 21 is indicative of the timepoint at which the bonds between PTH and the hydrogel matrix are broken down by hydrolysis and PTH is liberated.

These in vitro release data are representative of a highly controlled environment compared with the complexities of a biological system. In vivo, PTH can be liberated from thiol-ene hydrogels by hydrolysis or MMP activity.²² MMPs were not used in this in vitro work, as it is difficult to define a physiologically relevant value for addition of MMPs to in vitro studies. As such, in an in vivo environment, or if MMPs were introduced in vitro, PTH could be liberated by both hydrolysis and MMP activity; it is likely this release profile would differ. At day 28, protein measurements beyond 100% of the amount of PTH loaded in the thiol-ene gels were observed. This is likely because of the breakdown of the hydrogel. This limitation of the μBCA assay to measure PTH released from scaffolds was addressed by utilizing hydrogels with no PTH as controls to be subtracted from each measurement.

Considering the in vivo degradation of thiol-ene hydrogels together with in vitro release kinetics may lead to greater insight into the anticipated in vivo release profile. Although the in vivo degradation study was performed in a subcutaneous model, it provides useful information for understanding how the in vitro release profiles may be influenced in vivo. Thiol-ene hydrogels labeled with Alexa Fluor 680 were completely broken down in subcutaneous pockets between 8 and 21 days, whereas thiol-ene hydrogels used for in vitro release remained in PBS at 37°C for 28 days with no visible evidence of degradation. Therefore, although in vitro only 80% of the PTH had been released by day 21, in vivo the release profile would likely be accelerated, as all PTH would be released by day 21 if the hydrogel had completely degraded.

In the subcutaneous implant model, an initial drop in detectable fluorescence over the first 3–5 days was observed followed by a leveling off for the duration of the study. The initial drop is likely, in part, because of diffusion of unbound fluorophore out of the matrix. Although an excess of binding sites for the fluorophore were available, some fluorophore likely remained unbound after photo-polymerization. The hydrogel had completely degraded in the subcutaneous pockets of samples where fluorescence was not detected. When the signal was still present, hydrogel was still present. This observation may be indicative of a bulk degradation mechanism rather than a surface degradation mechanism, or may be because of the low removal rate of broken down hydrogel products in the subcutaneous pocket.

Interestingly, when the PPF scaffolds were incorporated, in vivo degradation did not occur as rapidly. The slowed degradation when thiol-ene hydrogels were combined with PPF scaffolds may be owing, in part, to the PPF scaffold playing a role in decreasing fluid flow through the hydrogels or providing a more mechanically protected environment for the hydrogel. Future studies exploring degradation in a bone defect model will likely indicate a more rapid degradation rate than the subcutaneous model as tissue damage, cell migration, MMPs, and fluid are higher in a bone defect.

Bone defect treatment of 3 or 10 μg PTH showed better healing than the other treatment groups when all outcome measurements are considered collectively. All samples in the 3 μg PTH treatment group were fully bridged by a combination of cartilage and bone. Although 12 weeks is a typical timepoint for analyzing healing of a bone defect, had data been collected at a later timepoint, full bony bridging may have been observed in all samples in the 3 or 10 μg treatment groups, especially considering the observation of endochondral ossification in these samples, as endochondral ossification is a normal part of fracture healing and bone formation.^31,32 In contrast, samples bridged by loosely connected fibrous tissue are less likely to continue on to bony union without intervention.³³ These findings support focusing on the 3–10 μg PTH range for future work on optimizing this biomaterial system for bone regeneration applications.

Disclosure Statement

No competing financial interests exist.

Funding Information

This work was supported by funding from the Colorado of Economic Development and International Trade, National Institute of Health/National Center for Research Resources Colorado Clinical and Translational Science Institute Grant No. UL1 RR025780 and National Institute of Dental and Craniofacial Research Grant No. DE016523.