|Home | About | Journals | Submit | Contact Us | Français|
Clinical management of critical bone defects remains a major challenge. Despite pre-clinical work demonstrating teriparatide (PTH1–34) effectiveness in small animals, inconclusive data from clinical trials have raised questions of dose and regimen. To address this, we completed a comprehensive study in the murine femoral allograft model, to assess the effects of dose (0.4, 4, and 40 μg/kg/day) and various treatment regimens on radiographic, histologic and biomechanical healing at 2, 4, and 9 weeks. Only the high dose (40 μg/kg) of PTH1–34 demonstrated significant effects when given daily over 9 weeks. Remarkably, equivalent biomechanical results were obtained with delayed, short treatment from 2 to 6 weeks that did not induce a significant increase in endochondral bone formation and callus volume. In contrast, PTH1–34 treatment from 1 to 5 weeks post-op demonstrated similar osteogenic effects as immediate-daily treatment for 9 weeks, but failed to achieve a significant increase in biomechanics at 9 weeks. Micro-CT and histologic analyses demonstrated that the 2-week delay in treatment allowed for timely completion of the endochondral phase, such that the prominent effects of PTH1–34 were enhanced intramembranous bone formation and remodeling at the graft-host junction. These findings support the potential use of PTH1–34 as an adjuvant therapy for massive allograft healing, and suggest that there may be an ideal treatment window in which a short course is administered following the endochondral phase to promote osteoblastic bone formation and remodeling to achieve superior union with modest callus formation.
Although most orthopaedic fractures heal, the clinical management of critical (>3cm) segmental defects continues to face major challenges for both amputation and limb salvage approaches, which have high rates of self-reported disability (40% to 50%) and continue to worsen over time.(1,2) Despite lacking osteogenic and remodeling capacity, devitalized cortical allografts remain a major option for reconstructive surgery because no equivalent alternatives exist. However, the limited new bone formation and lack of remodeling associated with massive allograft healing are directly associated with the 23–43% clinical failure rate caused by non-union (27–34%), late graft fracture (24–27%), and infection (9–16%).(3) Thus, the quest for a practical adjuvant therapy for massive allograft surgery remains a high priority.
Intermittent administration of PTH1–34, an active recombinant human peptide sequence of parathyroid hormone or (PTH1–34), has been proven to increase skeletal bone mass in osteoporotic patients.(4) Based on these anabolic properties, several groups have demonstrated that PTH1–34 is effective in small animal models of fracture healing.(5–10) Moreover, clinical case reports of off label use of PTH1–34 to heal delayed and non-unions have suggested that the drug could be effective as an adjuvant for challenging bone healing situations. (11,12) Unfortunately, a recent phase 2 clinical trial of distal radial fractures in postmenopausal women failed to meet its primary prospective endpoint with 40 μg/day teriparatide, although the study did show that 20 μg/day teriparatide accelerated the time to radiographic healing from 9.1 to 7.4 weeks versus saline controls (p = 0.006).(13) The efficacy of the therapy in this study however was confounded by a fracture model that would normally heal in the saline-treated group irrespective of treatment, which raises several questions that warrant further investigation of PTH1–34 dose and regimen in challenging preclinical models of bone healing.
One preclinical model of challenging bone healing that has emerged as a useful tool to understand the biological issues associated with delayed healing and non-unions is the murine femoral allograft model.(14) Similar to the clinical situation, femoral allografts in mice heal only via creeping fracture callus from the host that is mediated by endochondral ossification, and there is very little remodeling of the necrotic graft due to the absence of osteoclasts on the cortical surface of the allograft. Furthermore, murine allografted femurs achieve <50% of the torsional biomechanical properties of normal, ungrafted controls,(15) which is the primary outcome measure of reconstructive surgery for segmental defects in preclinical models. Live bone graft studies have demonstrated that the lack of intramembranous bone formation is due to the absence of periosteal cells, which are responsible for the angiogenic, osteogenic and remodeling responses during allograft healing.(16,17) Although these signals can be supplied exogenously via recombinant virus mediated gene transfer or cell therapy,(16,18,19) PTH1–34 adjuvant therapy offers the potential for a more practical solution for this problem.
In order to better understand the potential of PTH1–34 as an adjuvant therapy for structural allograft healing several questions about optimal dosing regimen must be addressed. With regards to dose, there is a significant discrepancy (approximately 100 fold) between the doses used in the rodent models (typically 40 μg/kg/day),(5–10) versus the approved clinical dose of 20 μg per day total (which in a 50 to 80kg person translates to 0.25 to 0.4 μg/kg/day). This discrepancy suggests that either the higher metabolic activity in small mammals requires higher dosing (on a per weight basis), and/or that efficacious dosing for bone healing is greater than that of anabolic therapy for generalized osteopenia. Moreover, questions about the ideal duration and timing of PTH1–34 therapy regimen for bone healing following reconstructive surgery have not been addressed. While osteoporosis therapy is indicated up to 18-months, it is unlikely that PTH1–34 treatment beyond a few months could significantly influence the dynamics of fracture healing. Given the potential safety concerns associated with long-term use of the drug, in addition to its high cost and the inconvenience of daily subcutaneous injections, identifying the minimum treatment duration required for efficacy is critical for this indication of PTH1–34 therapy. Finally, there is a question as to the ideal timing of PTH1–34 therapy following surgery. Since PTH has multiple mechanisms of action during bone repair via direct and indirect effects on multiple cell types during the initial inflammatory, intermediate osteogenic and late remodeling stages of bone healing, it is important to elucidate the effective window of therapy that achieves the maximum benefit on allograft healing. Therefore, a systematic study to investigate the appropriate clinically-translatable dosing regimen of PTH1–34 needs to be conducted. To this end, we examined the effects of various doses, timing, and duration of PTH1–34 therapy on radiographic, histologic and biomechanical properties of murine femoral allograft healing.
Devitalized femoral allografts were prepared from femurs of 9 week old female ICR mice as previously described.(14) The donor femurs were cut to 4 mm mid-diaphyseal allografts, bone marrow was flushed using saline from a syringe, then the allografts were bathed in 70% ethanol for 3 hours, washed in phosphate buffered saline (PBS) before storing at −70°C for at least 1 week before they were used.
All animal studies were performed in accordance with protocols approved by the University of Rochester’s Committee on Animal Resources. Femoral allograft surgeries were performed as previously described.(14) Briefly, devitalized ICR allografts were implanted into a 4 mm defect created in the mid-diaphysis of the left femur of recipient 9 week old female C57Bl/6 mouse using a 10mm diameter, 0.15mm thick diamond-sintered rotary saw (Brasseler USA, Inc, Savannah, GA), and secured in place with an intramedullary stainless steel pin (Figure 1A). Daily subcutaneous injections of teriparatide (human PTH1–34) (Forteo™, Eli Lilly and Co., Indianapolis, IN) or saline (control) was performed according to the dosing regimens described in Figure 1. Group names were defined by dose, timing of initial treatment and duration asp follows. Three daily doses of PTH1–34 were investigated (L = low dose of 0.4μg/kg/day; M = middle dose of 4 μg/kg/day; and H = high dose of 40 μg/kg/day). The treatment regimen was defined by the timing of the initial dose (0 or one day after the surgery; and 1, 2 or 4 weeks postop), and the timing of treatment withdrawal (2, 3, 4, 5, 6, 8 or 9 weeks post-op).
Daily doses of PTH1–34 (0.4, 4, and 40 μg/kg/day) or saline were administered to the mice (n=15 per group) via subcutaneous injections from one day after the surgery for 4 weeks (Figure 1B). The mice were then maintained without therapy for another 5 weeks before sacrificed at 9 weeks post-surgery. Healing of grafted femurs was monitored by weekly X-rays (Supplemental Figure 1), and the grafted femurs were harvested and subjected to micro-computed tomography (micro-CT) analysis. Then, 10 femurs were used for torsion testing, and the remaining 5 femurs were used for histology.
Administration of high dose (40 μg/kg/day) PTH1–34 was commenced either 1day after the surgery (immediately), or at 1, 2 or 4 weeks post-surgery. The dosing periods were either 2 or 4 or 9 weeks (Figure 1C). The saline treated group as a control and the H-0-4 group in experiment 1 were included in this analysis. A total of 15 mice in each group were sacrificed at 9 weeks post-surgery and subjected to micro-CT (n=15), biomechanical testing (n=10), and histology (n=5) as depicted in Figure 1C.
To elucidate the early mechanistic differences between immediate and delayed PTH1–34 therapy on allograft healing we evaluated the healing response at 2 or 4 weeks post-surgery. Three PTH1–34 dosing regimens starting either 1day after the surgery (Immediately), 1 week after surgery (1w delayed), or 2 weeks after surgery (2w delayed) were compared to saline treatment control group as depicted in Figure 1D. A total of 10 mice in each group were assessed by micro-CT and histology (n=5 per group per time point). In addition, 3 mice per group were sacrificed at various times (0, 3, 7, 10, 14, 21, 28 days) post-surgery for mRNA expression analysis.
The femurs were harvested from euthanized mice, disarticulated at the hip and knee joints, and the intramedullary pins were carefully removed. The specimens for biomechanical testing were stored at −20°C, and the specimens for histology were fixed in 10% neutral buffered formalin and stored in 70% ethanol, before micro CT imaging. The specimens were scanned at 12.5 microns isotropic resolution using the VivaCT 40 (Scanco Medical AG, Bassersdorf, Switzerland) as previously described.(11) From these 3D images, the callus bone volume was measured by manual segmentation, followed by standardized thresholding at a grayscale corresponding to 750 mgHA/cm3 based on a phantom of known HA concentrations. The Union Ratio, a measure of allograft osseointegration, which is based on the minimum graft surface area fraction upon which mineralized callus had formed, was also calculated as recently described.(11)
Immediately following micro-CT imaging, the torsional biomechanical properties of the grafted femurs were determined as previously described,(20) using an EnduraTec TestBench™ system (200 N.mm torque cell; Bose Corporation, Minnetonka, MN) at a rate of 1°/sec. Yield torque (TYield), ultimate torque (TUlt), torsional rigidity (TR) and work to failure were determined for each specimen. After torsion testing, the specimens were x-rayed to analyze the mode of failure as previously described.(20)
After micro-CT, femurs were decalcified in EDTA. At least, two nonconsecutive 3-μm paraffin embedded mid-sagittal sections were stained with either alcian blue/hematoxylin/orange G (AB/OG) or tartrate resistant acid phosphatase (TRAP). Cartilage area and osteoclast number (Oc.N) were measured in week 2 and week 4 samples in experiment 3 as described previously.(14) Briefly, cartilage area was measured by alcian blue stained area. Oc.N was determined from sections stained for TRAP by counting the number of TRAP positive cells. Oc.N inside the callus area (Oc.Ncallus) and on the surface of allograft (Oc.Ngraft) was evaluated separately. The mean values from the two nonconsecutive mid-sagittal sections represented the value for one mouse.
The mRNA expression levels of various genes within the tissue at graft-host junction were measured using real time, reverse transcription-polymerase chain reaction (RT-PCR). The 3mm long callus tissue that formed at graft-host junctions was harvested at postoperative days 3, 5, 7, 10, 14, 21, and 28 as shown in Figure 1D. Tissues were flushed with DEPC treated phosphate-buffered saline to wash out bone marrow cells. The samples were frozen in liquid nitrogen and homogenized using a dismembrator. Total RNA was extracted from the samples using the TRIzol Reagent (Invitrogen Corp., Carlsbad, CA). For cDNA synthesis, 1 μg RNA was reverse-transcribed using iScript cDNA synthesis kit (Bio-Rad Laboratories, Hercules, CA). Real-time PCR was performed using a 20-μL reaction volume. Signals were detected using the PerfeCTa SYBR Green FastMix (Quanta BioSciences, Inc., Gaithersburg, MD) with gene-specific primers. Gene-specific primers of type II collagen (colIIa1), type X collagen (colXa1), vascular endotherial growth factor-a (vegf-a), matrix metalloproteinase 9 (mmp9), type I collagen (colIa1), osteocalcin (oc), cathepsin K (ctsk), tartarate resistant acid phosphatase (trap) and β-actin as a housekeeping gene are shown in supplemental Table 1. The relative mRNA expression of each targeted gene was normalized by the cycle threshold values of β-actin.
One-way analysis of variance (ANOVA) and Tukey’s or Dunnett’s post-hoc multiple comparison tests were performed to assess differences among PTH1–34 treatment groups and compared to the control groups, respectively. P-values ≤ 0.05 were used to detect significant differences.
In order to establish the effective dose of the therapy for murine allograft reconstruction we injected animals with 0 (saline control), 0.4, 4, or 40 μg/kg/day of PTH1–34 for 4 weeks following surgery. As restoration of biomechanical integrity of the injured long bone is the primary goal of structural allografting, we assessed this outcome 9 weeks after surgery to allow for adequate healing for torsion testing.(20) High-dose PTH1–34 treatment (H-0-4) increased the grafted femur yield torque (TYield), ultimate torque (TUlt), and torsional rigidity (TR) to 2.6, 2.2, and 1.9 times those in the control group (Table 1). In contrast, middle- (M-0-4) and low-dose (L-0-4) PTH1–34 showed equivalent values to those in the control group. Assessment of the mode of failure confirmed these results, as control and low dose groups resulted in preunion at a rate of 40% and 50%, respectively, but most importantly never achieved mature union by the end of the study. In contrast, none of the allografts in the high dose group resulted in preunion, whereas 10% of those achieved mature union (Supplemental Table 2)
To assess the differential effects of duration of PTH1–34 therapy on allograft healing in our model, we evaluated high dose treatment regimens commencing immediately out to 2, 4, or 9 weeks post-surgery, and evaluated the biomechanical properties at 9 weeks. The H-0-9 group served as the positive control for this study, as all tested parameters of healing were significantly (p<0.05) greater than saline treated controls. Contrary to our expectation, we found that immediate treatment for only 2 weeks (H-0-2) was sufficient to significantly increase the grafted femur TUlt to 2.4 times that in the control group (p<0.05), and was not significantly different than the H-0-9 group (Table 1). We also observed a trend in which H-0-2 group demonstrated a 3- fold increase in TYield and 1.6-fold increase in TR of grafted femur over saline control, although these differences were not statistically significant. The 4-week (H-0-4) and 9-week (H-0-9) treatment groups were equivalent in biomechanical properties to 2 weeks-long (H-0-2) treatment, indicating that longer treatments do not further improve allograft healing in terms of bone strength. The conclusion was also supported by the failure mode analysis, which demonstrated that the additional 5 weeks of PTH1–34 treatment in the H-0-9 group resulted in only a 20% increase in the incidence of mature union compared the H-0-4 group (Supplemental Table 2).
To assess the efficacy of delayed PTH1–34 treatment on allograft healing in our model, we evaluated the effects of commencing the treatment after 1, 2, or 4 weeks post-reconstruction for 2 or 4 weeks thereafter. We found the treatment 4 weeks after surgery (H-4-8) to be completely ineffective, as no differences were observed compared to saline-treated controls. Interestingly, 1 or 2 week-delayed treatments resulted in equivalent biomechanical properties to the positive control H-0-9, regardless of the duration of treatment. However, the most remarkable results from this study came from the H-2-6 treatment group, which demonstrated significant increases in all parameters tested compared to saline controls, and achieved equivalent biomechanical outcomes to the H-0-9 treatment group.
While 9 weeks post-surgery is ideal for assessing drug effects on torsional biomechanics in this model (11), this time point proved to be too challenging for quantitative radiology and histomorphometry, due to the extensive remodeling that prohibits accurate segmentation of the graft from the host bone, and remodeling of callus and new periosteal bone. Additionally, unsegmented micro-CT analyses of the grafted femurs demonstrated that no differences in bone volume or bone mineral density at 9 weeks could be observed among the groups (data not shown). However, the 3D micro-CT rendering of the grafted femurs demonstrated gaps and irregular cortical surfaces at the graft-host junction of the control, L-0-4, and M-0-4 groups (Figure 2) that were not observed in the H-0-4 group, demonstrating that only the high dose has effects on bone healing in this model.
In order to better understand the mechanism responsible for these drug effects on biomechanical properties, we analyzed micro-CT images and histology at 2 and 4 weeks post-surgery (Figure 1D). These analyses were limited to only the different high dose treatment regimens, since the low and middle dose treatments were not biomechanically efficacious. Interestingly, while immediate PTH1–34 treatment significantly increased callus volume at 2 weeks (Figure 3C and H) compared to controls (Figure 3A and H), this increase was not observed in the 1 week delayed treatment group (Figure 3E and H), and was not significant at 4 weeks due to remodeling. No treatment related effects on the callus BMD were detected at either 2 or 4 weeks post surgery (Figure 3I). However, both the immediate and 1 week delayed treatment groups displayed significant increases in their Union Ratios by 4 weeks (Figure 3J), which demonstrates that this parameter of osseointegration is not related to callus volume, as we have previously reported.(11,15) The results of this experiment also demonstrated that while a 2-week delay in the PTH1–34 treatment did not increase the Union Ratio per se compared to saline controls (Figure 3J), micro CT data at 9 weeks showed remarkable end to end integration in the treated specimens, regardless of the delay, which underscores that the long term healing is independent of callus size.
Histological analyses of the healing allografts harvested 2 and 4 weeks post-surgery also provided evidence for PTH1–34-induced osseointegration independent of callus size (Figure 4). At 2 weeks, there was a small, albeit statistically insignificant, increase in the cartilage area with the immediate PTH1–34 treatment compared to the saline controls and 1 week delayed treatment (Figure 4A–C; Table 2). Moreover, there was no significant evidence of persistent cartilage at 4 weeks (Figure 4D–G; Table 2), regardless of the treatment. Furthermore, there were mild increases in the osteoclast number density at the surface of the treated allografts at 2 weeks, which increased by 4 weeks in all PTH1–34 treatment groups (Figure 4, Table 2). Furthermore, the osteoclast number density in the callus was significantly higher than that on the allograft surface at 2 weeks, regardless of the treatment. At 4 weeks, the osteoclast number density remained increased in the treated groups, but decreased significantly in the saline treated controls (Table 2).
To further elucidate the mechanism responsible for delayed PTH1–34 effects on allograft healing, we performed a time course experiment to assess changes in mRNA expression of markers of chondrocytes, angiogenesis, osteoblasts and osteoclasts over 28 days of healing (Figure 5). The results of these experiments corroborated several of the histologic findings. First, only the immediate PTH1–34 treatment had significant effects on chondrogenic (colIIa1) and hypertrophic (colXa1) gene expression that disappeared by 10 to 14 days, which is consistent with the small increases in cartilage area at 2 weeks and the absence of persistent cartilage at 4 weeks. Immediate PTH1–34 treatment also increased the expression of mmp9 and vegf-a, which are associated with cartilage removal and angiogenesis during fracture healing, at 14 days post surgery (17,21). Most importantly, we found that all regimens of high dose PTH1–34 significantly (p<0.05) increased markers of bone formation (colIa1, and oc) on days 21 and 28 post-surgery, which is consistent with the sustained bone formation at 4 weeks (Supplemental Figures 2 and 3).
Despite its clinical effectiveness in the treatment of osteoporosis (22,23), reports of clinical off-label use to resolve fibrotic nonunion fractures (24), and a growing body of preclinical data from various animal studies that raised expectations of its efficacy in accelerating bone repair (5–7,9,10,25–28), parathyroid hormone (PTH) therapy has yet to be widely clinically adopted for the management of nonoperative fractures or the more challenging cases of fibrotic nonunions. There are several reasons that contribute to the slow translation of PTH to the clinical or surgical management these indications. First, there have been safety concerns regarding the range of effective PTH1–34 doses in the animal studies of fracture repair (10–200 μg/kg/day), which correspond to tens or hundreds of multiples of the FDA approved doses of 20 or 40 μg/day (0.25 to 0.5 μg/kg/day for an 80 kg person) for treatment of osteoporosis. The high doses, in combination with long treatments, have been linked to high incidence of osteosarcomas in rats (23,29), which despite evidence of safety from surveillance studies of patients previously treated with PTH1–34, led the FDA to approve the drug with strong recommendations to limit its prescription to women and men with substantial osteoporosis and susceptibility to fracture, and placed a black box exclusion warning for patients at high risk for osteosarcoma (23). This regulatory barrier is compounded by the absence of long-term patent protection, and hence the lack of commercial incentive to perform expensive safety and efficacy clinical trials for challenging fractures (23). Moreover, there is no consensus on the outcome variables that could be used to definitively establish efficacy in human patients (30). A case-in-point is a recent study that indicated that the clinical value of PTH (teriparatide) treatment in distal radial fractures (assessed by the time to healing based on radiographic evidence of cortical union, patient-related wrist evaluation score, and early callus formation) was limited to marginal acceleration of healing that did not provide strong cost-risk-benefit justification (13,30). The inconclusive findings of this human trial in contrast to the efficacy evidence in animal studies underscore its limitations: i) the lack of validated thresholds of bone healing based on non- or minimally-invasive assessment, and ii) the fact that these fractures still healed with placebo in just weeks, which makes establishing the case for treatment efficacy difficult. Finally, there remain many unanswered questions about the appropriate timing of commencement and withdrawal of the treatment, which have not been thoroughly investigated in preclinical studies. Hence, the motivation for the current study was to address the latter questions of dose, timing, and duration of treatment, in hopes of identifying in a preclinical model a short treatment window that could help abate the more serious concerns about osteosarcoma risk in non-life threatening indications such as fracture repair, which could then incentivize definitive large animal studies and clinical trials of safety and efficacy.
Since allografts are typically complicated by nonunion and therefore are more challenging than osteotomy or closed fracture models, we elected to investigate PTH1–34 treatment regimens in a previously established mouse femoral reconstruction model (11,14,20). We have recently shown in this model that daily PTH1–34 (40 μg/kg/day) commencing immediately after devitalized allograft reconstruction for 6 weeks induced trabeculated bone callus formation and remarkable graft-host integration, which were correlated with more than doubling of the biomechanical parameters of torsional rigidity and yield torque compared to saline treated controls (15). In the current study, we first asked whether lower doses of PTH1–34 could be as effective as higher dose in enhancing allograft osseointegration and biomechanics. Consistent with several previous studies in rodent models of fracture repair (5–7,9,10,25–28), our findings demonstrated that only the high daily dose (40 μg/kg/day) showed remarkable effects on torsional properties of grafted femurs when administered for 9 weeks, while the lower doses of 0.4 and 4 μg/kg/day did not provide any benefits compared to controls. Rodent models typically report the use of doses as high as 10 to 200 μg/kg/day. While the difference between the effective clinical dose for osteoporosis and the much higher doses for rodent models of fracture repair are thought to arise from differences in metabolism and clearance of PTH, and typically require treatment periods amounting to >50% of the animal life span (31), conclusive statements about the effectiveness of lower dose might still require investigation in larger animal species that might better approximate the metabolism of the drug in humans, and a clinically relevant treatment window.
Our studies showed that immediate PTH treatment substantially improved early cartilaginous template formation in the fracture callus and subsequent mineralization, leading to improved healing at the host-allograft junction. This finding is supported by several previous reports that describe the main effects of intermittent PTH treatment on fracture repair to be through enhancement of the early proliferative response of chondroprogenitor and osteoprogenitor cells (9,10,31).
We also observed that short-term treatments (2 weeks), commencing immediately or with a 1- or 2-week delay following reconstruction, were just as effective in enhancing the biomechanical properties at 9 weeks as the continuous daily treatment for 9 weeks. This data suggests that short-term treatment during the first 4 weeks after surgery is sufficient to improve graft-host union from fibrous union to osseous union and that establishing osseous union is a primary mechanism of PTH1–34 induced increase in mechanical strength of grafted femur. These observations have great clinical relevance. The effectiveness of the short treatments could abate concerns related to osteosarcoma risk with prolonged treatment. Furthermore, given that PTH therapy is delivered via inconvenient subcutaneous injections and is very expensive, short treatment could enhance patient compliance and balance cost-benefit concerns. Moreover, since a likely clinical scenario that could benefit from PTH therapy might involve patients in whom bone repair may be delayed (31), the effectiveness of the delayed treatment in our murine model should encourage further investigation in larger species.
The observation that 1- or 2-week delayed short-term treatment was as effective as immediate treatment suggests that PTH1–34 might have different actions depending on the overlap with the different stages of healing and the cells involved. Our results corroborate previous findings that suggest that immediate treatment with PTH1–34 might have multiple effects that include suppressing inflammation upon surgery (32), early stimulation of proliferation of mesenchymal and periosteal stem cells (10), and enhanced early differentiation of chondroprogenitors leading to robust “endochondral ossification” bone repair (9,31), which is consistent with our observations of increased callus size in association with PTH1–34 treatments. On the other hand, 1- or 2-week delayed PTH therapy in which the treatment missed part or all of the endochondral ossification phase still induced persistent osteoblastic and osteoclastic activity up to 4 weeks. This was manifested by remarkable increases in colIa1 and oc gene expression at day 21 which began to decline by day 28. Interestingly, the 4-week delay in the treatment could not improve the biomechanical properties of the allografted femurs compared to controls. These findings suggest that bone repair could still be accomplished, even if treatment is delayed, via intramembranous ossification, and further suggest that the first 3 weeks could be the critical window for PTH1–34 therapy. Given that bone formation peaks at around 2 weeks after surgery and is down-regulated by 4 weeks after surgery in control mice that underwent femoral allograft surgery, PTH1–34 therapy would be effective as long as active bone formation sustains after surgery but becomes ineffective after repair reaction is attenuated. Some clinical case reports of off-label use of PTH1–34 demonstrated that patients with prolonged fracture non-union would benefit from anabolic therapy months to years after fracture (11,33,34), to our knowledge, however, there are no definitive data as to by when after surgery PTH1–34 would be effective for bone repair. Therefore, it should be tested in larger animal models or clinically how long after surgery PTH1–34 could be expected to show positive effect on bone repair. Although we failed to observe significant differences in osteoclast marker gene expression (ctsk and trap) during this critical phase of allograft healing, this was not surprising, as the significant PTH1–34 effects on osteoclasts were on their location rather than total numbers. As the osteoclasts on the graft surface are primarily involved in generating a new marrow space between the necrotic and new bone, while the osteoclasts in the callus are remodeling the osteoid into lamellar bone, this finding suggests that PTH1–34 effects are pertinent for callus remodeling, which was less of a factor in delayed treatment. Thus, it is unlikely that the mechanism responsible for delayed-PTH1–34 treatment effects on structural allografting involves increased bone resorption. Future studies should address the effects of PTH1–34 on the different cellular compartments involved in the repair and remodeling of the allograft.
Our findings provide strong evidence in support of the efficacy of delayed, short PTH1–34 treatment in a mouse model of challenging bone repair. A recent study investigated the effects of different PTH administration regimes applied at different stages of fracture healing in a an osteoporotic (ovariectomized or Ovx) rat model of tibial osteotomy healing (35), and similarly reported that the treatment improved fracture repair compared to untreated controls when PTH was administered either immediately or 7 days post osteotomy, independent of the administration frequency, but not after a 14 day delay. Collectively, our findings and others’ should motivate further investigation of the efficacy of PTH treatment in challenging bone repair scenarios in larger species and clinical studies.
We would like to thank Dr. Christopher Beck for his help with the statistical analysis, Ryan Tierny and the histology core for their excellent technical assistance, Michael Thullen for assistance with micro CT, and Jacy Krystal Bulaon for help with the biomechanical testing. This work was funded in part by grants from the Aircast Foundation and grants from the National Institutes of Health (AR056696, AR054041, DE019902).