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Shear Stress in Bone Marrow has a Dose Dependent Effect on cFos Gene Expression in In Situ Culture

  • Kimberly J. Curtis
  • Thomas R. Coughlin
  • Mary A. Varsanik
  • Glen L. NieburEmail author
Article

Abstract

Introduction

Mechanical stimulation of bone is necessary to maintain its mass and architecture. Osteocytes within the mineralized matrix are sensors of mechanical deformation of the hard tissue, and communicate with cells in the marrow to regulate bone remodeling. However, marrow cells are also subjected to mechanical stress during whole bone loading, and may contribute to mechanically regulated bone physiology. Previous results from our laboratory suggest that mechanotransduction in marrow cells is sufficient to cause bone formation in the absence of osteocyte signaling. In this study, we investigated whether bone formation and altered marrow cell gene expression response to stimulation was dependent on the shear stress imparted on the marrow by our loading regime.

Methods

Porcine trabecular bone explants were cultured in an in situ bioreactor for 5 or 28 days with stimulation twice daily. Gene expression and bone formation were quantified and compared to unstimulated controls. Correlation was used to assess the dependence on shear stress imparted by the loading regime calculated using computational fluid dynamics models.

Results

Vibratory stimulation resulted in a higher trabecular bone formation rate (p = 0.01) and a greater increase in bone volume fraction (p = 0.02) in comparison to control explants. Marrow cell expression of cFos increased with the calculated marrow shear stress in a dose-dependent manner (p = 0.002).

Conclusions

The results suggest that the shear stress due to interactions between marrow cells induces a mechanobiological response. Identification of marrow cell mechanotransduction pathways is essential to understand healthy and pathological bone adaptation and remodeling.

Keywords

Mechanobiology Bone adaptation Trabecular bone Gene regulation Computational modeling 

Notes

Acknowledgments

This material is based on work supported by the National Science Foundation under Grant No. CMMI-1453467 and CMMI-1100207. K.J.C. was supported by the Walther Cancer Foundation through a Notre Dame Harper Cancer Research Institute Interdisciplinary Interface Training Program fellowship and the Notre Dame Advanced Diagnostics and Therapeutics Institute’s Leiva Graduate Fellowship in Precision Medicine.

Conflict of interest

Kimberly J. Curtis, Thomas R. Coughlin, Mary A. Varsanik, and Glen L. Niebur declare no conflicts of interest.

Ethical Standards

No human subjects research was carried out by the authors for the research in this article, and no animal experiments were carried out by the authors for the research in this article.

Supplementary material

12195_2019_594_MOESM1_ESM.eps (3.5 mb)
Supplementary Figure 1 Cell metabolism and proliferation assays indicated that the cultured trabecular bone explants maintain their cell populations and that cells proliferate, although we have not ascertained whether the relative numbers of cell types are maintained. (A) Four trabecular bone explants were harvested and cultured in the bioreactor for 22 days. Three explants were exposed to LMMS at 50 Hz, 0.3 xg in 30 min bouts, twice per day, 5 days per week throughout culture. The remaining three explants were cultured with fluid flow, but no mechanical stimulation. A metabolic assay (CellTiter-Blue, Promega, Madison, WI) was carried out following manufacturer’s instructions. Briefly, on day 4 and day 21, 20% CellTiter-Blue reagent was added to the media and incubated for 24hr at 37 °C. After incubation, fluorescence of the supernatant from each explant was measured in triplicate in a 96-well plate at 570 nm excitation and 600 nm emission. Cell viability was not different between any conditions or time points, indicating that cells in the marrow remained viable during culture (ANOVA; p = 0.08). (B) Eight different trabecular bone explants excised from two pigs were cultured in the bioreactor for 5 days. Four explants were subjected to LMMS at 50 Hz, 0.3 xg in 30 min bouts, twice per day, on days 2 through 5 of culture. The remaining four explants were cultured statically. Twenty-four hours after explants were placed in the bioreactor, 10 µM BrdU (ThermoFisher) was added to the media for the remaining culture period. After culture, explants were fixed in 10% formalin, demineralized in 10% EDTA, processed, and embedded in paraffin. Explants were sectioned at 6 μm and mounted on microscope slides. Sections were dried at 60 °C, dewaxed in xylene, and hydrated through descending concentrations of ethanol. Incubation in HCl was used to break the DNA structure of the labeled cells followed by neutralization in borate buffer. Sections were blocked before being stained with rat anti-BrdU (ab6326) at a 1:40 dilution overnight at 4 °C. A goat anti-rat secondary antibody conjugated to Alexafluor 488 (ab150165) was applied at a dilution of 1:200 for one hour at room temperature. The nuclei were stained using propidium iodide (Sigma) before being mounted with a coverslip. Two sections per explant were imaged in five regions at 100X and BrdU positive nuclei were counted using ImageJ (NIH) and normalized to total area imaged. A representative image of BrdU-positive cells (white arrows) in the marrow is shown, indicating that marrow cells are proliferating during culture. We did not attempt to identify specific cell lineages. Scale bar is 50 μm. (C) There was no difference in cell proliferation between loaded and control samples after 5 days of bioreactor culture (Student’s t-test; p = 0.12). (EPS 3550 kb)

References

  1. 1.
    Aguirre, J. I., L. I. Plotkin, S. A. Stewart, R. S. Weinstein, A. M. Parfitt, S. C. Manolagas, and T. Bellido. Osteocyte apoptosis is induced by weightlessness in mice and precedes osteoclast recruitment and bone loss. J. Bone Miner. Res. 21:605–615, 2006.CrossRefGoogle Scholar
  2. 2.
    Batra, N., R. Kar, and J. X. Jiang. Gap junctions and hemichannels in signal transmission, function and development of bone. Biochim. Biophys. Acta 1909–1918:2012, 1818.Google Scholar
  3. 3.
    Birmingham, E., T. C. Kreipke, E. B. Dolan, T. R. Coughlin, P. Owens, L. M. McNamara, G. L. Niebur, and P. E. McHugh. Mechanical stimulation of bone marrow in situ induces bone formation in trabecular explants. Ann. Biomed. Eng. 43:1036–1050, 2014.CrossRefGoogle Scholar
  4. 4.
    Bonewald, L. F. The amazing osteocyte. J. Bone Miner. Res. 26:229–238, 2011.CrossRefGoogle Scholar
  5. 5.
    Castillo, A. B., and C. R. Jacobs. Mesenchymal stem cell mechanobiology. Curr. Osteoporos. Rep. 8:98–104, 2010.CrossRefGoogle Scholar
  6. 6.
    Chen, J. C., D. A. Hoey, M. Chua, R. Bellon, and C. R. Jacobs. Mechanical signals promote osteogenic fate through a primary cilia-mediated mechanism. FASEB J. 30:1504–1511, 2016.CrossRefGoogle Scholar
  7. 7.
    Chen, N. X., K. D. Ryder, F. M. Pavalko, C. H. Turner, D. B. Burr, J. Qiu, and R. L. Duncan. Ca(2+) regulates fluid shear-induced cytoskeletal reorganization and gene expression in osteoblasts. Am. J. Physiol. Cell Physiol. 278:C989–C997, 2000.CrossRefGoogle Scholar
  8. 8.
    Choi, J. S., and B. A. C. Harley. Marrow-inspired matrix cues rapidly affect early fate decisions of hematopoietic stem and progenitor cells. Sci. Adv. 3:e1600455, 2017.CrossRefGoogle Scholar
  9. 9.
    Coughlin, T. R., and G. L. Niebur. Fluid shear stress in trabecular bone marrow due to low-magnitude high-frequency vibration. J. Biomech. 45:2222–2229, 2012.CrossRefGoogle Scholar
  10. 10.
    Coughlin, T. R., J. Schiavi, M. Alyssa-Varsanik, M. Voisin, E. Birmingham, M. G. Haugh, L. M. McNamara, and G. L. Niebur. Primary cilia expression in bone marrow in response to mechanical stimulation in explant bioreactor culture. Eur. Cell. Mater. 32:111–122, 2016.CrossRefGoogle Scholar
  11. 11.
    Curtis, K. J., T. R. Coughlin, D. E. Mason, J. D. Boerckel, and G. L. Niebur. Bone marrow mechanotransduction in porcine explants alters kinase activation and enhances trabecular bone formation in the absence of osteocyte signaling. Bone 107:78–87, 2018.CrossRefGoogle Scholar
  12. 12.
    Eimori, K., N. Endo, S. Uchiyama, Y. Takahashi, H. Kawashima, and K. Watanabe. Disrupted bone metabolism in long-term bedridden patients. PLoS ONE 11:e0156991, 2016.CrossRefGoogle Scholar
  13. 13.
    Ezura, Y., J. Nagata, M. Nagao, H. Hemmi, T. Hayata, S. Rittling, D. T. Denhardt, and M. Noda. Hindlimb-unloading suppresses B cell population in the bone marrow and peripheral circulation associated with OPN expression in circulating blood cells. J. Bone Miner. Metab. 33:48–54, 2014.CrossRefGoogle Scholar
  14. 14.
    Fritton, J. C., E. R. Myers, T. M. Wright, and M. C. H. van der Meulen. Loading induces site-specific increases in mineral content assessed by microcomputed tomography of the mouse tibia. Bone 36:1030–1038, 2005.CrossRefGoogle Scholar
  15. 15.
    Frost, H. M. Bone’s mechanostat: a 2003 update. Anat. Rec. A 275:1081–1101, 2003.CrossRefGoogle Scholar
  16. 16.
    Haudenschild, A. K., A. H. Hsieh, S. Kapila, and J. C. Lotz. Pressure and distortion regulate human mesenchymal stem cell gene expression. Ann. Biomed. Eng. 37:492–502, 2009.CrossRefGoogle Scholar
  17. 17.
    Jansen, L. E., N. P. Birch, J. D. Schiffman, A. J. Crosby, and S. R. Peyton. Mechanics of intact bone marrow. J. Mech. Behav. Biomed. Mater. 50:299–307, 2015.CrossRefGoogle Scholar
  18. 18.
    Kelly, N. H., J. C. Schimenti, F. Patrick Ross, and M. C. H. van der Meulen. A method for isolating high quality RNA from mouse cortical and cancellous bone. Bone 68:1–5, 2014.CrossRefGoogle Scholar
  19. 19.
    Klein-Nulend, J., A. D. Bakker, R. G. Bacabac, A. Vatsa, and S. Weinbaum. Mechanosensation and transduction in osteocytes. Bone 54:182–190, 2013.CrossRefGoogle Scholar
  20. 20.
    Klein-Nulend, J., P. J. Nijweide, and E. H. Burger. Osteocyte and bone structure. Curr. Osteoporos. Rep. 1:5–10, 2003.CrossRefGoogle Scholar
  21. 21.
    Knothe Tate, M. L. Whither flows the fluid in bone? An osteocyte’s perspective. J. Biomech. 36:1409–1424, 2003.CrossRefGoogle Scholar
  22. 22.
    Kreipke, T. C., and G. L. Niebur. Anisotropic permeability of trabecular bone and its relationship to fabric and architecture: a computational study. Ann. Biomed. Eng. 45:1543–1554, 2017.CrossRefGoogle Scholar
  23. 23.
    Krølner, B., B. Toft, S. Pors Nielsen, and E. Tøndevold. Physical exercise as prophylaxis against involutional vertebral bone loss: a controlled trial. Clin. Sci. Lond. Engl. 64:541–546, 1983.CrossRefGoogle Scholar
  24. 24.
    Lan, S., S. Luo, B. K. Huh, A. Chandra, A. R. Altman, L. Qin, and X. S. Liu. 3D image registration is critical to ensure accurate detection of longitudinal changes in trabecular bone density, microstructure, and stiffness measurements in rat tibiae by in vivo microcomputed tomography (μCT). Bone 56:83–90, 2013.CrossRefGoogle Scholar
  25. 25.
    Leppä, S., M. Eriksson, R. Saffrich, W. Ansorge, and D. Bohmann. Complex functions of AP-1 transcription factors in differentiation and survival of PC12 cells. Mol. Cell. Biol. 21:4369–4378, 2001.CrossRefGoogle Scholar
  26. 26.
    Lin, F.-X., G.-Z. Zheng, B. Chang, R.-C. Chen, Q.-H. Zhang, P. Xie, D. Xie, G.-Y. Yu, Q.-X. Hu, D.-Z. Liu, S.-X. Du, and X.-D. Li. Connexin 43 modulates osteogenic differentiation of bone marrow stromal cells through GSK-3beta/Beta-catenin signaling pathways. Cell. Physiol. Biochem. 47:161–175, 2018.CrossRefGoogle Scholar
  27. 27.
    Metzger, T. A., T. C. Kreipke, T. J. Vaughan, L. M. McNamara, and G. L. Niebur. The in situ mechanics of trabecular bone marrow: the potential for mechanobiological response. J. Biomech. Eng. 137:011006, 2015.CrossRefGoogle Scholar
  28. 28.
    Metzger, T. A., S. A. Schwaner, A. J. LaNeve, T. C. Kreipke, and G. L. Niebur. Pressure and shear stress in trabecular bone marrow during whole bone loading. J. Biomech. 48:3035–3043, 2015.CrossRefGoogle Scholar
  29. 29.
    Metzger, T. A., T. J. Vaughan, L. M. McNamara, and G. L. Niebur. Altered architecture and cell populations affect bone marrow mechanobiology in the osteoporotic human femur. Biomech. Model. Mechanobiol. 16:841–850, 2017.CrossRefGoogle Scholar
  30. 30.
    Parfitt, A. M., M. K. Drezner, F. H. Glorieux, J. A. Kanis, H. Malluche, P. J. Meunier, S. M. Ott, and R. R. Recker. Bone histomorphometry: standardization of nomenclature, symbols, and units. Report of the ASBMR Histomorphometry Nomenclature Committee. J. Bone Miner. Res. 2:595–610, 1987.CrossRefGoogle Scholar
  31. 31.
    Pavalko, F. M., N. X. Chen, C. H. Turner, D. B. Burr, S. Atkinson, Y. F. Hsieh, J. Qiu, and R. L. Duncan. Fluid shear-induced mechanical signaling in MC3T3-E1 osteoblasts requires cytoskeleton-integrin interactions. Am. J. Physiol. 275:C1591–1601, 1998.CrossRefGoogle Scholar
  32. 32.
    Pavalko, F. M., S. M. Norvell, D. B. Burr, C. H. Turner, R. L. Duncan, and J. P. Bidwell. A model for mechanotransduction in bone cells: the load-bearing mechanosomes. J. Cell. Biochem. 88:104–112, 2003.CrossRefGoogle Scholar
  33. 33.
    Robling, A. G., P. J. Niziolek, L. A. Baldridge, K. W. Condon, M. R. Allen, I. Alam, S. M. Mantila, J. Gluhak-Heinrich, T. M. Bellido, S. E. Harris, and C. H. Turner. Mechanical stimulation of bone in vivo reduces osteocyte expression of Sost/sclerostin. J. Biol. Chem. 283:5866–5875, 2008.CrossRefGoogle Scholar
  34. 34.
    Rubin, C. T., E. Capilla, Y. K. Luu, B. Busa, H. Crawford, D. J. Nolan, V. Mittal, C. J. Rosen, J. E. Pessin, and S. Judex. Adipogenesis is inhibited by brief, daily exposure to high-frequency, extremely low-magnitude mechanical signals. Proc. Natl. Acad. Sci. USA 104:17879–17884, 2007.CrossRefGoogle Scholar
  35. 35.
    Rubin, C., G. Xu, and S. Judex. The anabolic activity of bone tissue, suppressed by disuse, is normalized by brief exposure to extremely low-magnitude mechanical stimuli. FASEB J. 15:2225–2229, 2001.CrossRefGoogle Scholar
  36. 36.
    Schaffler, M. B., W.-Y. Cheung, R. Majeska, and O. Kennedy. Osteocytes: master orchestrators of bone. Calcif. Tissue Int. 94:5–24, 2014.CrossRefGoogle Scholar
  37. 37.
    Schulte, F. A., F. M. Lambers, G. Kuhn, and R. Müller. In vivo micro-computed tomography allows direct three-dimensional quantification of both bone formation and bone resorption parameters using time-lapsed imaging. Bone 48:433–442, 2011.CrossRefGoogle Scholar
  38. 38.
    Sen, B., Z. Xie, N. Case, M. Ma, C. Rubin, and J. Rubin. Mechanical strain inhibits adipogenesis in mesenchymal stem cells by stimulating a durable beta-catenin signal. Endocrinology 149:6065–6075, 2008.CrossRefGoogle Scholar
  39. 39.
    Soves, C. P., J. D. Miller, D. L. Begun, R. S. Taichman, K. D. Hankenson, and S. A. Goldstein. Megakaryocytes are mechanically responsive and influence osteoblast proliferation and differentiation. Bone 66:111–120, 2014.CrossRefGoogle Scholar
  40. 40.
    Stavenschi, E., M. A. Corrigan, G. P. Johnson, M. Riffault, and D. A. Hoey. Physiological cyclic hydrostatic pressure induces osteogenic lineage commitment of human bone marrow stem cells: a systematic study. Stem Cell Res. Ther. 9:276, 2018.CrossRefGoogle Scholar
  41. 41.
    Stavenschi, E., M.-N. Labour, and D. A. Hoey. Oscillatory fluid flow induces the osteogenic lineage commitment of mesenchymal stem cells: The effect of shear stress magnitude, frequency, and duration. J. Biomech. 55:99–106, 2017.CrossRefGoogle Scholar
  42. 42.
    van der Meulen, M. C. H., X. Yang, T. G. Morgan, and M. P. G. Bostrom. The effects of loading on cancellous bone in the rabbit. Clin. Orthop. Relat. Res. 467:2000–2006, 2009.CrossRefGoogle Scholar
  43. 43.
    Verbruggen, S. W., T. J. Vaughan, and L. M. McNamara. Strain amplification in bone mechanobiology: a computational investigation of the in vivo mechanics of osteocytes. J. R. Soc. Interface 9:2735–2744, 2012.CrossRefGoogle Scholar
  44. 44.
    Waarsing, J. H., J. S. Day, J. C. van der Linden, A. G. Ederveen, C. Spanjers, N. De Clerck, A. Sasov, J. A. N. Verhaar, and H. Weinans. Detecting and tracking local changes in the tibiae of individual rats: a novel method to analyse longitudinal in vivo micro-CT data. Bone 34:163–169, 2004.CrossRefGoogle Scholar
  45. 45.
    Wagner, D. R., D. P. Lindsey, K. W. Li, P. Tummala, S. E. Chandran, R. L. Smith, M. T. Longaker, D. R. Carter, and G. S. Beaupre. Hydrostatic pressure enhances chondrogenic differentiation of human bone marrow stromal cells in osteochondrogenic medium. Ann. Biomed. Eng. 36:813–820, 2008.CrossRefGoogle Scholar
  46. 46.
    Webster, D., F. A. Schulte, F. M. Lambers, G. Kuhn, and R. Müller. Strain energy density gradients in bone marrow predict osteoblast and osteoclast activity: a finite element study. J. Biomech. 48:866–874, 2015.CrossRefGoogle Scholar
  47. 47.
    Weinbaum, S., S. C. Cowin, and Y. Zeng. A model for the excitation of osteocytes by mechanical loading-induced bone fluid shear stresses. J. Biomech. 27:339–360, 1994.CrossRefGoogle Scholar
  48. 48.
    Wuchter, P., J. Boda-Heggemann, B. K. Straub, C. Grund, C. Kuhn, U. Krause, A. Seckinger, W. K. Peitsch, H. Spring, A. D. Ho, and W. W. Franke. Processus and recessus adhaerentes: giant adherens cell junction systems connect and attract human mesenchymal stem cells. Cell Tissue Res. 328:499–514, 2007.CrossRefGoogle Scholar
  49. 49.
    Zerwekh, J. E., L. A. Ruml, F. Gottschalk, and C. Y. Pak. The effects of twelve weeks of bed rest on bone histology, biochemical markers of bone turnover, and calcium homeostasis in eleven normal subjects. J. Bone Miner. Res. 13:1594–1601, 1998.CrossRefGoogle Scholar

Copyright information

© Biomedical Engineering Society 2019

Authors and Affiliations

  1. 1.Tissue Mechanics LaboratoryUniversity of Notre DameNotre DameUSA
  2. 2.Bioengineering Graduate ProgramUniversity of Notre DameNotre DameUSA
  3. 3.Department of Aerospace and Mechanical EngineeringUniversity of Notre DameNotre DameUSA

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