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. 2014 Aug 9;472(11):3523–3532. doi: 10.1007/s11999-014-3829-x

Fractures in Geriatric Mice Show Decreased Callus Expansion and Bone Volume

Luke A Lopas 1, Nicole S Belkin 1, Patricia L Mutyaba 1,2, Chancellor F Gray 1, Kurt D Hankenson 1,2, Jaimo Ahn 1,✉
PMCID: PMC4182401  PMID: 25106797

Abstract

Background

Poor fracture healing in geriatric populations is a significant source of morbidity, mortality, and cost to individuals and society; however, a fundamental biologic understanding of age-dependent healing remains elusive. The development of an aged-based fracture model system would allow for a mechanistic understanding that could guide future biologic treatments.

Questions/purposes

Using a small animal model of long-bone fracture healing based on chronologic age, we asked how aging affected (1) the amount, density, and proportion of bone formed during healing; (2) the amount of cartilage produced and the progression to bone during healing; (3) the callus structure and timing of the fracture healing; and (4) the behavior of progenitor cells relative to the observed deficiencies of geriatric fracture healing.

Methods

Transverse, traumatic tibial diaphyseal fractures were created in 5-month-old (n = 104; young adult) and 25-month-old (n = 107; which we defined as geriatric, and are approximately equivalent to 70–85 year-old humans) C57BL/6 mice. Fracture calluses were harvested at seven times from 0 to 40 days postfracture for micro-CT analysis (total volume, bone volume, bone volume fraction, connectivity density, structure model index, trabecular number, trabecular thickness, trabecular spacing, total mineral content, bone mineral content, tissue mineral density, bone mineral density, degree of anisotropy, and polar moment of inertia), histomorphometry (total callus area, cartilage area, percent of cartilage, hypertrophic cartilage area, percent of hypertrophic cartilage area, bone and osteoid area, percent of bone and osteoid area), and gene expression quantification (fold change).

Results

The geriatric mice produced a less robust healing response characterized by a pronounced decrease in callus amount (mean total volume at 20 days postfracture, 30.08 ± 11.53 mm3 versus 43.19 ± 18.39 mm3; p = 0.009), density (mean bone mineral density at 20 days postfracture, 171.14 ± 64.20 mg hydroxyapatite [HA]/cm3 versus 210.79 ± 37.60 mg HA/cm3; p = 0.016), and less total cartilage (mean cartilage area at 10 days postfracture, 101,279 ± 46,755 square pixels versus 302,167 ± 137,806 square pixels; p = 0.013) and bone content (mean bone volume at 20 days postfracture, 11.68 ± 3.18 mm3 versus 22.34 ± 10.59 mm3; p < 0.001) compared with the young adult mice. However, the amount of cartilage and bone relative to the total callus size was similar between the adult and geriatric mice (mean bone volume fraction at 25 days postfracture, 0.48 ± 0.10 versus 0.50 ± 0.13; p = 0.793), and the relative expression of chondrogenic (mean fold change in SOX9 at 10 days postfracture, 135 + 25 versus 90 ± 52; p = 0.221) and osteogenic genes (mean fold change in osterix at 20 days postfracture, 22.2 ± 5.3 versus 18.7 ± 5.2; p = 0.324) was similar. Analysis of mesenchymal cell proliferation in the geriatric mice relative to adult mice showed a decrease in proliferation (mean percent of undifferentiated mesenchymal cells staining proliferating cell nuclear antigen [PCNA] positive at 10 days postfracture, 25% ± 6.8% versus 42% ± 14.5%; p = 0.047).

Conclusions

Our findings suggest that the molecular program of fracture healing is intact in geriatric mice, as it is in geriatric humans, but callus expansion is reduced in magnitude.

Clinical Relevance

Our study showed altered healing capacity in a relevant animal model of geriatric fracture healing. The understanding that callus expansion and bone volume are decreased with aging can help guide the development of targeted therapeutics for these difficult to heal fractures.

Electronic supplementary material

The online version of this article (doi:10.1007/s11999-014-3829-x) contains supplementary material, which is available to authorized users.

Introduction

Fragility fractures are a substantial burden for older patients, incurring a mean cost of USD 35,000 in the initial 12-month period per hip fracture treated [10]. A mortality rate as much as 21% has been reported for the first year after hip fracture [36] and an age-adjusted relative risk of mortality of 6.68 for females sustaining a hip fracture [9]. Morphologic and biologic changes in bone with age result in decreased bone strength [21] and in longer bone healing times than in younger counterparts [17]. In the United States alone, it is estimated that there will be greater than three million fragility fractures annually by 2025, costing more than USD 25 billion [7].

Factors such as inferior trabecular structure and decreased vascularity have been linked to compromised bone mass and biomechanics in the elderly which has provided valuable clinical insights into the pathogenesis of fractures in this subgroup [20, 34, 41]. However, cellular and molecular bases for the observed biologic deficiencies in fracture healing in older patients remain elusive. Previous studies have shown an altered balance between bone formation and resorption [8, 15, 29, 30, 37, 38], decreased bone formation [23, 27], or delays in callus consolidation [24] in laboratory models of aging. However, many of these models have not adequately reflected geriatric animal age [2, 15, 23, 24, 27]. Although valuable, these studies represent a pool of models that are difficult to apply to aging humans. We used male C57BL/6 mice from the National Institute of Aging (NIA) aging rodent colonies at 5 and 25 months old. These mice have well-evaluated aging and skeletal characteristics [15, 16] and serve as the basis for choosing 25-month-old mice (defined as geriatric in this study) who have 50% to 75% survival, corresponding to a human age of 70 to 85 years [1].

Using a rationally selected, small animal model of long-bone fracture healing based on chronologic age that is well suited for further genetic and mechanistic investigation, we asked: how aging affected (1) the amount, density, and proportion of bone formed during fracture healing—using total volume, bone volume, bone volume fraction, bone mineral content, tissue mineral content, tissue mineral density, bone mineral density; (2) the amount of cartilage produced and the progression to bone, a critical process in endochondral fracture healing—callus area, total cartilage area, hypertrophic cartilage area, percent cartilage, percent hypertrophic cartilage, bone and osteoid tissue area, percent bone and osteoid tissue, and relative expression of Sox9 and osterix; (3) callus structure and timing of the fracture healing process—trabecular number, trabecular thickness, trabecular spacing, polar moment of inertia; and (4) the behavior of progenitor cells in relation to the observed deficiencies of geriatric fracture healing—proliferating undifferentiated mesenchyme, prehypertrophic chondrocytes, and new bone.

Materials and Methods

Animal Experimentation and Surgical Model

Our Institutional Animal Care and Use Committee approved all procedures. The experimental group (n = 107) consisted of 25-month-old (geriatric) C57BL/6 mice from the NIA [28]; 5-month-old (young adult) mice (n = 104) from the same colonies (maintained to minimize genetic drift by the NIA) comprised the control group. Mice underwent bilateral tibial fractures as described [12, 33]. Briefly, the mice were administered buprenorphine and isoflurane, their legs were prepared aseptically, prestabilized with an intramedullary pin, and a traumatic closed fracture was created with a guillotine (three-point bend mechanism). The mice were allowed to move freely and given buprenorphine/Nutella® (Ferrero USA, Inc, Somerset, NJ, USA) [25] for 3 additional days and euthanized at 5, 10, 15, 20, 25, 30, or 40 days postfracture for analysis.

Tissue Harvest and Preparation

Mice were euthanized via CO2 inhalation and cervical dislocation. Fractured tibias were dissected for bone and fracture tissue and fixed in 4% paraformaldehyde at 4° C for 72 hours. Intramedullary pins were removed and tibias were transferred to 70% ethanol for imaging and then decalcified in 15% formic acid, dehydrated in ethanol, and embedded in paraffin. Sagittal 5-μm sections were prepared for histologic analysis. (Supplemental Table 1. Supplemental materials are available with the online version of CORR®).

Micro-CT

To determine the amount, density, and proportion of bone formed during fracture healing we measured and calculated total volume, bone volume, bone volume fraction, bone mineral content, tissue mineral content, tissue mineral density, and bone mineral density using micro-CT [32]. To determine callus structure and timing of the fracture healing process, trabecular number, trabecular thickness, trabecular spacing, and polar moment of inertia were measured using micro-CT. Tibias were scanned using a vivaCT 40 micro-CT system (Scanco Medical, Brüttisellen, Switzerland) with an isotropic three-dimensional voxel size of 21 µm as described [11]. Briefly, callus and cortical bone sections were identified manually and then spline interpolation was performed between slices no more than 10 slices apart (0.105 mm). Points were reviewed and reinterpolation performed if necessary. A fixed, global threshold of 16% of the maximum gray value, which corresponds to a mineral density of 169.8 mg hydroxyapatite [HA]/cm3 was used to distinguish mineralized from unmineralized tissue. Cortical bone sections were removed, and a global threshold was applied that calculated bone volume, tissue mineral density, callus volume, and trabecular thickness. Day 10 samples could not be analyzed for total callus volume because soft tissue was too similar to early fracture callus, but osseous parameters (bone volume, trabecular thickness) were measured with thresholds applied.

Immunohistochemistry and Histomorphometry

To determine the amount of cartilage produced and the progression to bone, a critical process in endochondral fracture healing, callus area, total cartilage area, hypertrophic cartilage area, percent cartilage, percent hypertrophic cartilage, bone and osteoid tissue area, percent bone and osteoid tissue were measured using histomorphometry and relative expression of Sox9 and osterix using quantitative PCR. Sections were deparaffinized, gradually rehydrated, and stained with safranin O or Masson’s trichrome, then dehydrated, and mounted with Permount™ (Thermo Fisher Scientific, Waltham, MA, USA), and analysis was performed as described [12, 25].

Cartilage analysis was performed on two sections stained with safranin O and Fast Green FCF and averaged. Images were obtained with an Olympus BX51 microscope (Olympus Corp, Tokyo, Japan) using a SPOT RT3™ 2 megapixel camera (Spot Imaging Solutions, Sterling Heights, MI, USA), digitally stitched and loaded in ImageJ (NIH, Bethesda, MD, USA) for analysis. Total callus area was determined with a manual contour around the fracture callus. The color thresholding tool was used to determine the amount of cartilage tissue and relative composition of the callus.

Bone and osteoid tissue analysis was performed using Masson’s trichrome with the same process for safranin O slides, except the color thresholding was performed to identify the amount and relative contribution of bone and osteoid tissue.

In addition, to examine the role of mesenchymal proliferation rates in geriatric fracture healing, we evaluated proliferating cellular nuclear antigen (PCNA) in undifferentiated mesenchyme, prehypertrophic chondrocytes, and new bone. Undifferentiated mesenchyme was identified on sequential slides stained with hematoxylin and safranin O and defined as cells that contained nuclei but clearly had not begun to hypertrophy and differentiate into chondrocytes. Sections were stained with an anti-PCNA antibody (1:1000; Abcam, Cambridge, MA, USA) and counterstained with Gill’s hematoxylin. (Appendix 1. Supplemental materials are available with the online version of CORR®.) Two to four ×40 images of undifferentiated mesenchyme, prehypertrophic chondrocytes, and new bone were analyzed with the ImageJ tool Colour Deconvolution (http://www.dentistry.bham.ac.uk/landinig/software/cdeconv/cdeconv.html) to count cells positive for peroxidase staining. Cells stained with either peroxidase or hematoxylin determined total cell count. The number of proliferating cells was normalized to the amount of bone perimeter.

Real-time Quantitative PCR

Total RNA was harvested from callus using miRNeasy (Qiagen, Valencia, CA, USA) [11]. RNA was reverse transcribed using oligo (dT) primers and superscript III reverse transcriptase (Invitrogen, Carlsbad, CA, USA). cDNA was used for real-time quantitative PCR using Fast SYBR® Green and 7500 Fast Real-time PCR System (both from Applied Biosystems, Foster City, CA, USA). Proper amplicon formation was confirmed by melt-curve analysis, and relative mRNA expression levels were calculated using the double delta-Ct method. (Appendix 1. Supplemental materials are available with the online version of CORR®) All values were normalized to β-actin and to 0 days postfracture values for 5-month-old mice. Fracture calluses were evaluated at 0, 10, and 20 days after fracture for expression of Sox9 (master chondrogenic transcription factor) and osterix (master osteoblastic transcriptional regulator).

Statistical Analysis

A two-tailed Student’s t-test was used to compare the young adult and geriatric mice at equivalent times. A result was considered significant when p values were 0.05 or less. Results are presented as mean ± SD of young adult mice versus mean ± SD of geriatric mice, followed by the p value.

Results

Micro-CT measurements (total volume) showed that young adult mice produced and maintained a callus approximately twice as large as that of geriatric mice at 15 (total volume, 62.24 ± 6.98 versus 29.44 ± 15.13 mm3; p = 0.001) through 40 days after fracture (total volume, 54.64 ± 18.25 versus 23.52 ± 18.01 mm3; p = 0.004) (Fig. 1A). Geriatric mice produced approximately four times less bone volume at 15 (bone volume, 18.28 ± 4.38 versus 4.89 ± 2.71 mm3; p < 0.001) through 40 days postfracture (bone volume, 21.00 ± 9.42 versus 6.29 ± 3.21, mm3; p < 0.001) (Fig. 1B); however, the relative amount of bone (bone volume/total volume) differed at early times, eg, 15 days postfracture (bone volume fraction, 0.30 ± 0.08 versus 0.17 ± 0.04; p = 0.005) and 20 (bone volume fraction, 0.51 ± 0.08 versus 0.42 ± 0.14; p = 0.011), but not thereafter (Fig. 1C). Micro computed tomography reconstructions show decreased bone formation in geriatric mice (Fig. 2). The fractures in the geriatric mice produced less bone mineral content from 15 days postfracture (6625 ± 2011 versus 1835 ± 1015 mg; p < 0.001) to 40 days postfracture (14,378 ± 5681 versus 3679 ± 1697 mg; p < 0.001) with the exception of 30 days (10,257 ± 6765 versus 4318 ± 1803 mg; p = 0.064) (Fig. 3A). The same pattern was observed in tissue mineral content (Fig. 3B) at 15 days postfracture (6798 ± 1811 versus 1855 ± 1,039 mg; p < 0.001) through 40 days postfracture (10,882 ± 4920 versus 2599 ± 1308 mg; p < 0.001). At 30 days postfracture tissue mineral density (573 ± 94 versus 457 ± 35 mg HA/cm3; p = 0.018) (Fig. 3C) in the geriatric mice failed to keep pace with that of young adult mice. This continued at 40 days postfracture for bone mineral density (206 ± 81 versus 123 ± 38 mg HA/cm3; p = 0.020) (Fig. 3D) and tissue mineral density (706 ± 103 versus 589 ± 54 mg HA/cm3; p = 0.013).

Fig. 1A–C.

Fig. 1A–C

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Geriatric (25 months) mice show reduced (A) total callus volume and (B) bone volume but similar (C) bone volume fraction (bone volume/total callus volume) relative to young adult (5 months) mice. Micro-CT results of young adult and geriatric C57Bl/6 mice at 10, 15, 20, 25, 30, and 40 days postfracture show that young adult mice form larger calluses with greater total bone volume geriatric mice. Bone volume fraction is increased in young adult mice at 15 and 20 days postfracture. Values are mean ± SD; BV = bone volume; TV = total volume; *p < 0.05; **p < 0.01; ***p < 0.001.

Fig. 2A–H.

Fig. 2A–H

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Three-dimensional reconstructions and midcoronal views of three-dimensional reconstructions of healing fracture callus show delayed, decreased bone formation in geriatric versus young adult mice at (A) 5 months old 10 days postfracture, (B) 5 months old 15 days after fracture, (C) 5 months old 20 days after fracture, (D) 5 months old 40 days after fracture, (E) 25 months old 10 days after fracture, (F) 25 months old 15 days after fracture, (G) 25 months old 20 days after fracture, and (H) 25 months old 40 days after fracture.

Fig. 3A–D.

Fig. 3A–D

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Geriatric mice produce tissue of lower quality during fracture healing. A comparison of micro-CT results between young adult and geriatric C57Bl/6 mice at 15, 20, 25, 30, and 40 days postfracture shows decreased (A) bone and (B) tissue mineral content in geriatric mice relative to young adult mice. (C) Decreased late tissue mineral density occurs in geriatric mice. (D) Increased bone mineral density occurs at early and late times in young adult mice compared with geriatric mice. Values are mean ± SD; HA = hydroxyapatite; *p < 0.05; **p < 0.01; ***p < 0.001.

Histomorphometry confirmed that young adult mice produce more callus (callus area) than geriatric mice by 10 days postfracture (730,256 ± 257,762 versus 343,107 ± 78,469 square pixels; p = 0.011) (Fig. 4A). At 10 days postfracture, corresponding to peak cartilage, young adult mice produced more total cartilage than geriatric mice (302,167 ± 137,806 versus 101,279 ± 46,755 square pixels; p = 0.013) (Fig. 4B). Although not significant, young adult mice appeared to trend toward producing more hypertrophic cartilage at 10 days postfracture (142,170 ± 99,672 versus 40,367 ± 25,819 square pixels; p = 0.055) (Fig. 4C). No difference was found in the percent cartilage content of the callus at 10 days postfracture (41.17% ± 10.13% versus 28.94% ± 8.74%; p = 0.063) (Fig. 4D) and in the percent of total cartilage that was hypertrophic at 15 days postfracture (88.76% ± 14.56% versus 70.21 ± 16.05%; p = 0.078) (Fig. 4E). Osseous tissue formation did not reach high levels until 20 days postfracture with cartilage resorption nearly complete for both groups (Fig. 4F–G). Geriatric mice exhibit decreased total cartilage production and delayed resorption (Fig. 5). Osterix and Sox9 were upregulated at 10 and 20 days postfracture for young adult and geriatric mice, with much greater upregulation at 10 days postfracture for Sox9 (Supplemental Fig. 1. Supplemental materials are available with the online version of CORR®), and no differences were detected between the callus in geriatric and in young adult mice.

Fig. 4A–G.

Fig. 4A–G

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Geriatric mice showed delayed endochondral ossification and produced less cartilage and bone than young adult mice. Histomorphometric comparisons of cartilage (safranin O) and bone (Masson’s trichrome) formation at 5, 10, 15, 20, 30, and 40 days postfracture are shown for (A) callus area, (B) cartilage area, (C) hypertrophic cartilage area, (D) percent cartilage, (E) percent hypertrophic cartilage, (F) bone and osteoid area, and (G) percent bone and osteoid. Values are mean ± SD; *p < 0.05; **p < 0.01; ***p < 0.001.

Fig. 5A–H.

Fig. 5A–H

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Comparison views of young adult and geriatric mice at 5, 10, and 20 days postfracture (Stain, safranin O; original magnification, ×10) and 40 days postfracture (Stain, Masson’s trichrome; original magnification; ×10) are shown for (A) 5 months old 5 days, (B) 5 months old 10 days, (C) 5 months old 20 days, (D) 5 months old 40 days, (E) 25 months old 5 days, (F) 25 months old 10 days, (G) 25 months old 20 days, and (H) 25 months old 40 days after fracture.

In addition to a delay in peak bone formation (Fig. 1C), geriatric mice showed decreased bone mineral density at 15 days postfracture (110.24 ± 31.15 mg HA/cm3 versus 66.40 ± 21.33 mg HA/cm3; p = 0.015), 20 days postfracture (210.79 ± 37.60 mg HA/cm3 versus 171.14 ± 64.20 mg HA/cm3; p = 0.016), and 40 days postfracture (206.42 ± 81.14 mg HA/cm3 versus 123.72 ± 37.74 mg HA/cm3; p = 0.021) (Fig. 3C). Trabecular number was greater in young adult mice at 15 days postfracture (3.22 ± 1.16, 1/mm versus 2.01 ± 0.64, 1/mm; p = 0.042) and 20 days postfracture (6.07 ± 0.85, 1/mm versus 5.13 ± 1.66, 1/mm; p = 0.022) days postfracture, peaked at 20 days postfracture, and then gradually declined thereafter. Geriatric mice showed a temporal shift in this pattern (Fig. 6A). Geriatric mice produced thinner trabeculae from 10 days postfracture (0.13 ± 0.05 mm versus 0.09 ± 0.03 mm; p = 0.039) to 40 days postfracture (0.18 ± 0.06 mm versus 0.11 ± 0.02 mm; p = 0.005) (Fig. 6B). Trabecular spacing was greater in geriatric mice at 15 days postfracture (0.43 ± 0.13 mm versus 0.62 ± 0.14 mm; p = 0.047) and 20 days postfracture (0.18 ± 0.03 mm versus 0.24 ± 0.09 mm; p = 0.003) (Fig. 6C). Geriatric fracture calluses had a lower polar moment of inertia (important indicator of torsional stability) at 10 (0.35 ± 0.24 mm4 versus 0.18 ± 0.12 mm4; p = 0.042) through 40 days postfracture (7.77 ± 6.04 mm4 versus 2.17 ± 1.46 mm4; p = 0.023) (Fig. 6D). While fractures in young adult mice showed increased polar moment of inertia values with time (approximately threefold from 15 to 40 days postfracture), fractures in geriatric mice showed relatively little increase.

Fig. 6A–D.

Fig. 6A–D

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Geriatric mice develop an inferiorly organized callus consistent with decreased mechanical properties. Callus architecture was analyzed by micro-CT for young adult and geriatric C57Bl/6 mice at 10, 15, 20, 25, 30, and 40 days postfracture for (A) trabecular number, (B) trabecular thickness, (C) trabecular spacing, and (D) polar moment of inertia. Values are mean ± SD; *p < 0.05; **p < 0.01 ***p < 0.001.

At 10 days postfracture, there was a decrease in proliferating cells in the undifferentiated mesenchyme from geriatric mice (41.96% ± 14.55% versus 25.16% ± 6.76%; p = 0.047) (Fig. 7A). No difference was observed in PCNA staining rates in prehypertrophic cartilage (10 days postfracture) or new bone formation (10 and 20 days postfracture) (Fig. 7B–C). A comparison of undifferentiated mesenchyme was performed (Fig. 8).

Fig. 7A–C.

Fig. 7A–C

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Geriatric mice showed a reduction in mesenchymal cell proliferation. Immunohistochemistry was performed for proliferating cellular nuclear antigen in (A) undifferentiated mesenchyme (UDM) at 5 and 10 days postfracture, (B) prehypertrophic cartilage at 10 days postfracture, and (C) newly formed bone at 10 and 20 days postfracture for young adult and geriatric mice. Geriatric mice showed a decreased proportion of proliferating cellular nuclear antigen stained undifferentiated mesenchymal cells at 10 days postfracture. Values are mean ± SD; *p < 0.05; **p < 0.01; ***p < 0.001.

Fig. 8A–B.

Fig. 8A–B

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A comparison of undifferentiated mesenchyme at 10 days postfracture in young adult and geriatric mice is shown. Solid arrows indicate positive (brown) staining cells (Stain, anti-proliferating cellular nuclear antigen; original magnification, ×40). Dashed arrows indicate negative (purple) staining cells (Stain, hematoxylin; original magnification, ×40).

Discussion

Fractures in geriatric individuals are a substantial disease burden at individual and societal levels. Not only is bone in older individuals more susceptible to fracture, it also has reduced healing capacity. Improved understanding of the fundamental biologic mechanisms of decreased fracture healing in older individuals will likely provide a basis for rationally selected therapeutics. Studies designed to evaluate the variable of interest, chronologic age, are needed to elucidate the underlying biology. Therefore, we selected C57BL/6 mice, 5 and 25 months old, as they represent skeletally mature young adult and geriatric populations and make chronologic age the variable of interest. We found that geriatric mice had much smaller fracture healing calluses that progressed at a delayed pace but were capable of undergoing endochondral and intramembranous ossification, as observed histologically and genetically.

There are several potential limitations to our study. We evaluated healing in only two age groups of mice. Inclusion of more age groups might show more clearly how healing capacity changes with age but would have substantially increased the time and cost of the study. In addition, studies on molecular mechanism are warranted. For instance, gene pathway and comprehensive gene expression analysis could further define variation in fracture healing in older individuals. These studies are ongoing. While a strength of our model includes rational use of aging characteristics separating young adult and geriatric mice, the fracture technique that we used has an important intrinsic limitation. The traumatic generation of injury introduces a degree of heterogeneity that cannot be precisely controlled. Surgical osteotomy models would mitigate but not eliminate this particular problem, but at the same time introduce other undesired effects (ie, need for surgical dissection at the injury site, bony injury that does not resemble traumatic fractures, soft tissue injury that does not resemble traumatic fracture). With this in mind, we chose a modified 3-point bend traumatic long bone fracture model [5] that has been validated and has provided consistent, reproducible quantification data in our hands.

In the current study, we observed consistent reduction in the amount of regenerating tissue and total bone but with maintenance of callus proportions. Our results are consistent with published results of fracture healing in rodent models; however, in general, these studies did not use animals with ages that truly reflect geriatric human age [23, 24, 27]. This is an important point because model systems that more faithfully mimic the human condition of interest likely hold the most potential for ultimately improving the clinical care of patients. Other models that have been used to investigate aged bone formation include an accelerated senescence mouse model (SAMP6), which has shown relatively weak and brittle bones but with increased cortical bone mass [31]. However, variable differences in fracture healing have been observed when comparing in vivo with controls [14, 18]. Another popular model has been the use of ovariectomy to simulate an osteoporotic-like state [35]. However, the process of ovariectomy, especially in the popular rat model, does not mimic well osteoporotic disease as observed in humans [13]. The use of systems that best mimic the clinical entity being modeled will improve the applicability of findings to the clinical arena.

Geriatric mice showed reduced total and hypertrophic cartilage production and delays in the peak rates of cartilage resorption and bone formation via endochondral ossification. This is consistent with what was shown with increasing age in another study [23]. However, decreased total cartilage production was not present in the geriatric mice in the study by Lu et al. [23], as was shown in our study.

Geriatric mice also produced a callus with reduced tissue mineral density, smaller trabeculae, and a much smaller polar moment of inertia (suggesting mechanical inferiority) when compared with the young adult mice. Timing data combined with organization and architecture data suggested a delayed bone formation response in the geriatric mice, resulting in lower trabecular number and connectivity and increased spacing, ultimately reflecting a slower expansion process during early to midhealing. As healing progressed, these parameters tended to partially catch up to those of the younger counterparts. The observed increase with time of polar moment of inertia in young adult mice compared with a comparatively stagnant polar moment of inertia in geriatric mice, especially when combined with the increased density of the newly formed bone (higher tissue mineral density at 30 and 40 days postfracture) in young adult mice, suggests that geriatric mice produce a callus that is less suitable for weightbearing. Clinically, this is consistent with younger patients returning to weightbearing activities more quickly than geriatric patients and perhaps avoiding some of the complications associated with prolonged immobilization.

Despite these differences in quantity, time, and quality, the overall cellular and molecular mechanisms of fracture healing appeared to be preserved in the geriatric mice and were parallel to the process observed in the young adult mice. Analysis of phenotypic gene expression early during healing revealed intact molecular programming in geriatric mice (Supplemental Fig. 1. Supplemental material is available with the online version of CORR). Despite similar genetic expression patterns, the differences in callus that are seen by 10 days postfracture lead us to speculate that aged calluses are unable to undergo appropriate callus expansion. One likely explanation for the reduction in callus size is a decrease in mesenchymal stem cells and reduced proliferative capacity of mesenchymal stem cells from the geriatric mice [3, 4, 6]. Mutyaba et al. [26] showed that mesenchymal stem cells explanted from geriatric mice are reduced in number (as determined by colony-forming unit fibroblasts). Mesenchymal stem cells from geriatric mice also show decreased proliferation in vitro, which suggests alterations in proliferative capacity of the mesenchymal stem cell lineage are at least to some degree cell-autonomous. Other studies of aged fracture healing have suggested progenitor cell deficiencies [22], impaired vasculogenesis [23, 27], and an altered inflammatory response [11, 19, 27, 39, 40] as potential mechanistic culprits. The role of these pathways can be further examined in our characterized aged fracture model.

Taken as a whole, our data showed that geriatric mice produced a much less robust fracture-healing response when compared with that of young adult mice. Our data point toward a deficiency in progenitor cell proliferation as an explanation for the observed tissue volume deficiency of geriatric fracture healing. However, as progression through the stages of endochondral healing was observed, the overall ability of the geriatric progenitor cells to differentiate remained intact. These observations may parallel fracture healing in geriatric humans, in whom healing may be delayed or diminished but not mechanistically different. Our model is ideally suited for future studies investigating potential therapeutics because the fundamental variable in our model is chronologic age and not a proxy manipulation designed to simulate increased age. Additional studies investigating differences in this model, especially during the early phases of fracture healing, could help direct and inform future interventional therapeutics.

Electronic supplementary material

Supplementary material 1 (DOC 26 kb) (26KB, doc)
Supplementary material 2 (PPTX 751 kb) (751.3KB, pptx)
Supplementary material 3 (DOC 32 kb) (32.5KB, doc)

Acknowledgments

We thank Michael Karp MS and Brian Horwich MS (micro-CT data acquisition) Department of Bioengineering, University of Pennsylvania; Derek Dopkin BA (colony management) Department of Clinical Studies-New Bolton Center, University of Pennsylvania; Allison Williams BS (quantitative PCR) Department of Clinical Studies-New Bolton Center, University of Pennsylvania; and Michael Dishowitz PhD (experimental design and techniques) Department of Clinical Studies-New Bolton Center, University of Pennsylvania for their technical contributions.

Footnotes

One or more of the authors certify that they have received, during the study period, funding from NIH R03AG040670 (JA, KDH), American Geriatric Society Jahnigan Award (JA), McCabe Foundation Pilot Award (JA, KDH), and Penn Center for Musculoskeletal Disorders Pilot Award (NIH P30AR050950) (JA, KDH). One of the authors (KDH) certifies that he or she, or a member of his or her immediate family, has received or may receive payments or benefits, during the study period, an amount of less than USD 10,000 from Skelegen (Philadelphia, PA, USA). One of the authors (JA) certifies that he or she, or a member of his or her immediate family, has received or may receive payments or benefits, during the study period, an amount less than USD 10,000 from Synthes Inc (West Chester, PA, USA), an amount of USD 10,000 to USD 100,000 from Merck & Co, Inc (Whitehouse Station, NJ, USA), and an amount less than USD 10,000 from U&I Corp (Uijeongbu-si, Gyeonggi-do, Korea).

All ICMJE Conflict of Interest Forms for authors and Clinical Orthopaedics and Related Research ® editors and board members are on file with the publication and can be viewed on request.

Each author certifies that his or her institution approved the animal protocol for this investigation and that all investigations were conducted in conformity with ethical principles of research.

This work was performed at University of Pennsylvania, Philadelphia, PA, USA.

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