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Tumor necrosis factor α and interleukin-1β modulate calcium and nitric oxide signaling in mechanically stimulated osteocytes

  1. A. D. Bakker1,†,
  2. V. C. da Silva2,†,
  3. R. Krishnan3,
  4. R. G. Bacabac1,
  5. M. E. Blaauboer1,
  6. Y.-C. Lin3,
  7. R. A. C. Marcantonio4,
  8. J. A. Cirelli4,
  9. J. Klein-Nulend1,*

Article first published online: 29 OCT 2009

DOI: 10.1002/art.24920

Issue

Arthritis & Rheumatism

Arthritis & Rheumatism

Volume 60, Issue 11, pages 3336–3345, November 2009

Additional Information

How to Cite

Bakker, A. D., da Silva, V. C., Krishnan, R., Bacabac, R. G., Blaauboer, M. E., Lin, Y.-C., Marcantonio, R. A. C., Cirelli, J. A. and Klein-Nulend, J. (2009), Tumor necrosis factor α and interleukin-1β modulate calcium and nitric oxide signaling in mechanically stimulated osteocytes. Arthritis & Rheumatism, 60: 3336–3345. doi: 10.1002/art.24920

Author Information

  1. 1

    ACTA–Universiteit van Amsterdam and Vrije Universiteit, Research Institute MOVE, Amsterdam, The Netherlands

  2. 2

    ACTA–Universiteit van Amsterdam and Vrije Universiteit, Research Institute MOVE, Amsterdam, The Netherlands, and UNESP-São Paulo State University, School of Dentistry at Araraquara, São Paulo, Brazil

  3. 3

    Harvard School of Public Health, Boston, Massachusetts

  4. 4

    UNESP–São Paulo State University, School of Dentistry at Araraquara, São Paulo, Brazil

  1. †

    Drs. Bakker and da Silva contributed equally to this work.

*Department of Oral Cell Biology, ACTA–Vrije Universiteit, Van der Boechorststraat 7, 1081 BT Amsterdam, The Netherlands

Publication History

  1. Issue published online: 29 OCT 2009
  2. Article first published online: 29 OCT 2009
  3. Manuscript Accepted: 27 JUL 2009
  4. Manuscript Received: 1 AUG 2008

Funded by

  • Research Institute MOVE of the Vrije Universiteit Amsterdam
  • Foundation for the Coordination of Higher Education and Graduate Training, Brazil. Grant Number: MEC/CAPES BEX0481/06-8
  • Space Research Organization of The Netherlands. Grant Number: MG-055
  • Netherlands Organization For International Cooperation In Higher Education. Grant Number: Physics Development Project grant PHL-146

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Abstract

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. AUTHOR CONTRIBUTIONS
  7. Acknowledgements
  8. REFERENCES

Objective

Inflammatory diseases often coincide with reduced bone mass. Mechanoresponsive osteocytes regulate bone mass by maintaining the balance between bone formation and resorption. Despite its biologic significance, the effect of inflammation on osteocyte mechanoresponsiveness is not understood. To fill this gap, we investigated whether the inflammatory cytokines tumor necrosis factor α (TNFα) and interleukin-1β (IL-1β) modulate the osteocyte response to mechanical loading.

Methods

MLO-Y4 osteocytes were incubated with TNFα (0.5–30 ng/ml) or IL-1β (0.1–10 ng/ml) for 30 minutes or 24 hours, or with calcium inhibitors for 30 minutes. Cells were subjected to mechanical loading by pulsatile fluid flow (mean ± amplitude 0.7 ± 0.3 Pa, 5 Hz), and the response was quantified by measuring nitric oxide (NO) production using Griess reagent and by measuring intracellular calcium concentration ([Ca2+]i) using Fluo-4/AM. Focal adhesions and filamentous actin (F-actin) were visualized by immunostaining, and apoptosis was quantified by measuring caspase 3/7 activity. Cell-generated tractions were quantified using traction force microscopy, and cytoskeletal stiffness was quantified using optical magnetic twisting cytometry.

Results

Pulsatile fluid flow increased [Ca2+]i within seconds (in 13% of cells) and NO production within 5 minutes (4.7-fold). TNFα and IL-1β inhibited these responses. Calcium inhibitors decreased pulsatile fluid flow–induced NO production. TNFα and IL-1β affected cytoskeletal stiffness, likely because 24 hours of incubation with TNFα and IL-1β decreased the amount of F-actin. Incubation with IL-1β for 24 hours stimulated osteocyte apoptosis.

Conclusion

Our results suggest that TNFα and IL-1β inhibit mechanical loading–induced NO production by osteocytes via abrogation of pulsatile fluid flow–stimulated [Ca2+]i, and that IL-1β stimulates osteocyte apoptosis. Since both NO and osteocyte apoptosis affect osteoclasts, these findings provide a mechanism by which inflammatory cytokines might contribute to bone loss and consequently affect bone mass in rheumatoid arthritis.

The most important task of the skeleton is to provide mechanical support in order to withstand the force of gravity, and to support muscle forces during movement. This mechanical performance is secured by the constant adaptation of bone mass to its mechanical loading environment. Adaptation of bone mass is brought about by the coordinated actions of bone-resorbing osteoclasts and bone-forming osteoblasts, which in turn are orchestrated by the most mechanosensitive cells in bone, the osteocytes.

During inflammatory diseases such as rheumatoid arthritis (RA), the balance between bone formation and resorption is often disturbed, resulting in localized bone loss around the affected joints as well as in generalized osteoporosis (1–4). This bone loss is multifactorial and might be caused by the physiologic adaptation of bone to reduced physical activity that is common in patients with RA, or by the use of corticosteroids by the patients. In addition, it is likely that proinflammatory cytokines such as tumor necrosis factor α (TNFα) and interleukin-1β (IL-1β), which play an important role in the etiology of RA, also contribute to this bone loss (5). Indeed, inhibition of inflammation in RA by administration of antibodies against TNFα has been shown to arrest bone loss in the hip and the spine, underscoring the important role of this proinflammatory cytokine in bone loss in patients with RA (6). While it is now well known that TNFα and IL-1β affect bone metabolism by direct stimulation of bone resorption (7–11) and inhibition of bone formation (7, 12), the effects of these cytokines on the regulation of bone mass by osteocytes are virtually unknown.

Osteocytes are the most prevalent bone cells, and with their cell bodies located in lacunae and their long interconnected cell processes positioned in the canaliculi (13), they are uniquely positioned within the bone matrix to sense mechanical loading. It is well known that lack of mechanical loading results in a rapid loss of bone mass (14, 15) and that osteocytes are essential for this catabolic response of bone to unloading (16). It is generally assumed that loading on bone generates a flow of interstitial fluid through the lacunocanalicular network (17). This flow is sensed by the osteocytes, which respond by the release of signaling factors such as nitric oxide (NO). These signaling factors can alter the recruitment and activity of osteoblasts and osteoclasts (17–20), thereby affecting bone mass. We hypothesized that this mechanosensitive signaling of osteocytes decreases in the presence of cytokines, which could provide a mechanism explaining how cytokines contribute to the generalized bone loss in patients with RA.

MATERIALS AND METHODS

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. AUTHOR CONTRIBUTIONS
  7. Acknowledgements
  8. REFERENCES

Bone cell culture.

MLO-Y4 osteocytes, a cell line derived from mouse long bones (kindly provided by Dr. L. F. Bonewald, University of Missouri, Kansas City, MO) (21), were cultured in α-minimum essential medium (α-MEM) supplemented with 5% fetal bovine serum (FBS; Gibco, Grand Island, NY), 5% calf serum (Gibco), 50 μg/ml gentamicin (Gibco), 1.25 μg/ml Fungizone (Gibco), 60 μg/ml penicillin (Sigma, St. Louis, MO), and 50 μg/ml streptomycin (Sigma) at 37°C and 5% CO2 in air.

Pulsatile fluid flow.

MLO-Y4 osteocytes between passages 30 and 35 were harvested using 0.25% trypsin (Difco, Detroit, MI) and 0.1% EDTA (Sigma) in phosphate buffered saline (PBS) and seeded at 2 × 104 cells/cm2 over an area of 15 cm2 on 50 μg/ml poly-L-lysine hydrobromide (Sigma)–coated glass slides in α-MEM with 5% FBS, 5% calf serum, and antibiotics. They were incubated overnight in preparation for the pulsatile fluid flow experiments. Pulsatile fluid flow (mean ± amplitude 0.7 ± 0.3 Pa, 5 Hz) was generated by pumping 13 ml of culture medium in a pulsatile manner through a parallel-plate flow chamber (65 × 24 × 0.3 mm) containing the bone cells (22, 23). Pulsatile fluid flow was monitored throughout the experiment using a small animal blood flow sensor (T206; Transonic Systems, Ithaca, NY). Control cultures were kept in a petri dish containing 13 ml medium.

Cytokine treatment.

Within both static and pulsatile fluid flow–loaded groups, the cells were incubated with TNFα (0.5, 10, or 30 ng/ml; Sigma) or IL-1β (0.1, 1, or 10 ng/ml; Sigma) for 30 minutes or 24 hours before the start of the experiment, or with calcium inhibitors (described below) for 30 minutes before the start of the experiment. The concentrations of cytokines selected for this study were based on concentrations found in plasma and synovial fluid of patients with RA (24–26).

Intracellular calcium levels.

MLO-Y4 osteocytes between passages 30 and 35 were seeded at 1 × 104 cells/cm2 on 50 μg/ml poly-L-lysine hydrobromide–coated 5-cm2 glass slides in α-MEM with antibiotics and 5% FBS and 5% calf serum. Osteocytes were incubated overnight at 37°C in an atmosphere of 5% CO2 in air to promote cell attachment prior to fluid flow experiments as described below.

One hour before static and pulsatile fluid flow cultures, cells were incubated with 10 μM Fluo-4/AM (Invitrogen, Eugene, OR) in α-MEM without serum and antibiotics at 37°C for 30 minutes. Osteocytes were gently washed with sterile Dulbecco's PBS (Gibco). TNFα (10 ng/ml) or IL-1β (1 ng/ml) was added to α-MEM without serum and antibiotics 30 minutes before static and pulsatile fluid flow cultures. Pulsatile fluid flow (mean ± amplitude 0.7 ± 0.3 Pa, 5 Hz) was generated by pumping 13 ml of CO2-independent medium without L-glutamine (Gibco) in a pulsatile manner through a small parallel-plate flow chamber (20 × 18 × 0.30 mm) containing the bone cells.

Intracellular calcium levels were analyzed using confocal microscopy (TCS-SP2; Leica, Mannheim, Germany). Intracellular calcium concentration ([Ca2+]i) was monitored via fluorescence intensity measurements in single Fluo-4/AM–loaded cells, using a 63× 1.2 water correction objective. The excitation wavelength used was 488 nm, and emission spectra peaked at 516 nm. Fluorescence images were recorded every 7 seconds. The fluorescence intensity was quantified using Leica Confocal Software (Leica Microsystems, Wetzlar, Germany). The background fluorescence intensity was subtracted from the fluorescence intensity in each osteocyte. The number of cells showing an increase in [Ca2+]i were counted and expressed as a percentage of the total number of cells visible. Data were obtained from at least 4 independent experiments. Between 30 and 90 osteocytes were analyzed per treatment group within each experiment.

NO.

The cell medium was collected and assayed for NO concentrations at 5, 15, and 30 minutes after the onset of mechanical loading in the pulsatile fluid flow–exposed cultures and at similar time points in the static cultures. NO was measured as nitrite (NO2–) accumulation in the conditioned medium using Griess reagent (1% sulfanilamide, 0.1% naphthylethylene-diamine-dihydrochloride, and 2.5M H3PO4). Pulsatile fluid flow–induced NO production alters osteoblast and osteoclast activity (18, 20), and cytokine-induced changes in NO production by mechanically stimulated osteocytes can therefore be linked to changes in bone mass. Moreover, NO production is essential for the adaptive response of bone to mechanical loading in vivo, making it a meaningful parameter to quantify for studies on bone cell mechanoresponsiveness (27, 28).

Caspase 3/7 activity and DNA content.

After 30 minutes of culture with or without TNFα or IL-1β, cell lysis–based reagent of the Caspase-Glo 3/7 Assay (Promega, Madison, WI) was added to the cells for the assessment of caspase 3/7 activity with a luminometer (Berthold, Bad Wildbad, Germany), according to the manufacturer's instructions. DNA in the cell lysate was determined by CyQUANT Cell Proliferation Assay (Molecular Probes, Eugene, OR). Apoptosis was expressed in relative light units per ng of DNA.

Calcium inhibitors.

To determine whether NO production is dependent on calcium, calcium inhibitors were used under static and pulsatile fluid flow conditions of MLO-Y4 osteocytes. To inhibit intracellular calcium mobilization from the endoplasmic reticulum, 8-(diethylamino)octyl-3,4,5-trimethoxybenzoate·HCl (TMB-8; Biomol, Plymouth Meeting, PA) was used at a concentration of 20 μM (29). Gadolinium chloride (GdCl3; Sigma) was used at 10 μM to inhibit mechanosensitive calcium channels, and EGTA (Sigma) was used as an extracellular calcium scavenger at a concentration of 2 mM (29).

Staining of paxillin and filamentous actin (F-actin).

MLO-Y4 osteocytes were seeded at 1.2 × 104 cells/cm2 in chamber slides (Lab-Tek; Nunc, Naperville, IL), incubated overnight, and then incubated for an additional 30 minutes with the calcium inhibitors TMB-8, GdCl3, or EGTA, as described above. Osteocytes were fixed with a fixative containing 60 mM PIPES, 25 mM HEPES, 5 nM EGTA, 1 mM MgCl2, 3% sucrose, 0.1% Triton X-100, and 3.7% formaldehyde, washed with PBS, and incubated for 1.5 hours with 5% FBS, 5% glycine, and 0.1% Triton X-100 in PBS. Subsequently, cells were incubated overnight at 4°C with 1:80 monoclonal antipaxillin (Cell Signaling Technology, Danvers, MA) in blocking buffer and incubated for 1.5 hours in a solution containing 1:250 Alexa Fluor 555–conjugated goat anti-rabbit IgG (Invitrogen) in blocking buffer.

For F-actin quantification, MLO-Y4 osteocytes were seeded at 1.2 × 104 cells/cm2 in Lab-Tek chamber slides and incubated overnight, followed by incubation for 30 minutes with the calcium inhibitors TMB-8, GdCl3, or EGTA, as described above, or for 30 minutes or 24 hours with TNFα (10 ng/ml) or IL-1β (1 ng/ml). F-actin was extracted and quantified according to the protocol of Pritchard and Guilak (30).

Traction force measurements in osteocytes.

To determine whether calcium inhibition affects cytoskeletal tension, osteocyte-generated contractile stresses, called tractions, were assessed using traction force microscopy (31). MLO-Y4 osteocytes were seeded at 1 × 103 cells/cm2 on type I collagen–coated deformable polyacrylamide gel substrates of 4 kPa stiffness (32). The cells were incubated overnight in bone cell culture medium at 37°C and 5% CO2 in air. On the day of the experiment, the isolated adherent cell was subjected to GdCl3, TMB-8, or EGTA incubation for 1 hour prior to the measurement. Thereafter, we obtained phase-contrast images of the cell as well as fluorescent images of nanobeads embedded within the substrate immediately underneath the cell. The displacement field was computed by comparing the fluorescent microbead image obtained during the experiment with a reference image obtained at the end of the experiment after detaching the cell from its underlying substrate. The projected cell area was calculated based on a manual trace of the cell contour determined from the phase-contrast image of the cell. From the displacement field we calculated the traction field, and from the traction field we computed a scalar measure of contractility called the contractile moment (31).

Measurements of osteocyte cytoskeletal stiffness.

Osteocyte elastic modulus was measured using optical magnetic twisting cytometry. MLO-Y4 cells were seeded at 2 × 105 cells/well in type I collagen–coated 96-well plates (Immunon Removawells; Dynatech, Chantilly, VA) and left to attach overnight. The cells were incubated with GdCl3, TMB-8, or EGTA for 1 hour and were subsequently incubated with microbeads (4-μm diameter; 68,400 beads/well) that were coated with a peptide containing Arg-Gly-Asp for an additional 20 minutes. The beads were magnetized and twisted by placing the bead-loaded cells in a known magnetic field. Nanoscale displacements of the beads were recorded via a CCD camera. Based on these measurements and the applied magnetic torque, the elastic modulus was determined as previously described (33, 34). For some experiments, MLO-Y4 cells were incubated with TNFα (10 ng/ml) or IL-1β (1 ng/ml) rather than with calcium inhibitors for 24 hours prior to performing optical magnetic twisting cytometry measurements.

Statistical analysis.

Mann-Whitney U tests or Student's unpaired 2-tailed t-tests were used to compare cytokine levels in calcium inhibitor–treated groups with those in their untreated controls. P values less than 0.05 were considered significant.

RESULTS

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. AUTHOR CONTRIBUTIONS
  7. Acknowledgements
  8. REFERENCES

Pulsatile fluid flow and cytokines do not affect cell number.

When we subjected the MLO-Y4 osteocytes to pulsatile fluid flow, TNFα or IL-1β did not affect the total amount of DNA (mean ± SEM) (static 1.8 ± 0.1 μg; static plus 30 ng/ml TNFα 2.1 ± 0.1 μg; static plus 10 ng/ml IL-1β 1.6 ± 0.3 μg; pulsatile fluid flow 1.9 ± 0.7 μg), suggesting that no cells died and that proliferation of MLO-Y4 cells was not significantly affected by the addition of cytokines.

TNFα and IL-1β inhibit pulsatile fluid flow–up-regulated NO production.

We quantified pulsatile fluid flow–induced changes in NO production, which is an essential mediator of the anabolic response of bone to mechanical loading in vivo (27, 28). TNFα (0.5–30 ng/ml) or IL-1β (0.1–10 ng/ml) was added for 30 minutes or 24 hours before as well as during 30 minutes of subjecting MLO-Y4 osteocytes to pulsatile fluid flow or static control conditions. The addition of neither TNFα nor IL-1β for 30 minutes affected NO production under static culture conditions (Figures 1A and B). At 5 minutes, pulsatile fluid flow caused a rapid and significant increase in NO production (4.7-fold) compared with static cultures (Figures 1A–D), consistent with previous observations (18, 20). No such significant effect of pulsatile fluid flow on NO production was observed in the presence of TNFα at 0.5 and 10 ng/ml (Figure 1C).

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Figure 1. Effect of 30 minutes of incubation with tumor necrosis factor α (TNFα) and interleukin-1β (IL-1β) on nitric oxide (NO) production by MLO-Y4 osteocytes. A, TNFα did not change NO production under static culture conditions.B, IL-1β did not change NO production under static culture conditions.C, Pulsatile fluid flow (PFF) stimulated NO production. This response was inhibited by TNFα. D, Pulsatile fluid flow stimulated the NO response in the absence of cytokines. In the presence of IL-1β, no significant effect of pulsatile fluid flow on NO production was observed except for IL-1β at 0.1 ng/ml at 30 minutes. Values are the mean and SEM from a minimum of 4 independent experiments. Stat = static control culture. Dashed line represents no effect of pulsatile fluid flow on NO production (a ratio of 1 for the effect of pulsatile fluid flow to the effect of static culture conditions). ∗ = P < 0.05 versus static control conditions; # = P < 0.05 versus pulsatile fluid flow in the absence of cytokines.

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At later time points, the effect of 30 minutes of incubation with TNFα on pulsatile fluid flow–induced NO production became even clearer. At 15 minutes, pulsatile fluid flow stimulated NO production 6.6-fold, while 30 minutes of TNFα treatment at all concentrations led to a significant decrease in pulsatile fluid flow–stimulated NO production. Importantly, as a result of this decrease, no significant effect of pulsatile fluid flow on NO production was observed in the presence of TNFα. At 30 minutes, the significant effect of pulsatile fluid flow on NO production was inhibited by treatment with 30 ng/ml TNFα and completely abolished in the presence of TNFα at 0.5 and 10 ng/ml. Likewise, no significant effect of pulsatile fluid flow on NO production by MLO-Y4 cells was observed after 30 minutes of IL-1β treatment at any time point, except for 0.1 ng/ml IL-1β at 30 minutes after the start of pulsatile fluid flow.

Neither TNFα nor IL-1β affected NO production under static culture conditions (Figures 2A and B) when applied for 24 hours. Pulsatile fluid flow caused a significant increase in NO production (6.3-fold within 5 minutes) compared with static cultures (Figures 2A–D), but no such significant effect of pulsatile fluid flow on NO production was observed after 24 hours of treatment with TNFα at 30 ng/ml, except at 30 minutes after the start of pulsatile fluid flow (Figure 2C). Pulsatile fluid flow significantly affected NO production after 24 hours of incubation with TNFα at 0.5 and 10 ng/ml at all time points (Figure 2C). Incubation of MLO-Y4 cells with IL-1β (0.1–10 ng/ml) for 24 hours caused a dramatic decrease in the magnitude of the NO response to pulsatile fluid flow at 5 minutes (Figure 2D). After 15 minutes of pulsatile fluid flow treatment, the NO response was significantly reduced by 24 hours of incubation with 10 ng/ml IL-1β and completely absent after 24 hours of incubation with 0.1 ng/ml IL-1β (Figure 2D). Thus, both cytokines negatively affected the increase in NO production that is normally observed after application of mechanical stimulation by pulsatile fluid flow in osteocytes.

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Figure 2. Effect of 24 hours of incubation with TNFα and IL-1β on NO production by MLO-Y4 osteocytes. A, TNFα did not change NO production under static culture conditions.B, IL-1β did not change NO production under static culture conditions.C, Pulsatile fluid flow stimulated NO production. No significant effect of pulsatile fluid flow on NO production was observed in the presence of TNFα at 30 ng/ml 5 and 15 minutes after the start of pulsatile fluid flow. D, Pulsatile fluid flow stimulated the NO response in the absence of cytokines. Five minutes after the start of pulsatile fluid flow, IL-1β at all concentrations reduced the magnitude of the NO response of MLO-Y4 cells compared with that in control cultures. Values are the mean and SEM from a minimum of 5 independent experiments. ∗ = P < 0.05 versus static control conditions; # = P < 0.05 versus pulsatile fluid flow in the absence of cytokines. See Figure 1 for explanations and definitions.

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TNFα or IL-1β inhibits pulsatile fluid flow–stimulated [Ca2+]i.

In addition to NO, we studied the effects of TNFα and IL-1β on the pulsatile fluid flow–induced rise in [Ca2+]i, which is one of the first cellular responses occurring in the cascade of events elicited by subjecting mechanosensitive bone cells to fluid flow (35). TNFα (10 ng/ml) and IL-1β (1 ng/ml) were added 30 minutes or 24 hours before as well as during 30 minutes of subjecting MLO-Y4 osteocytes to pulsatile fluid flow or static control conditions. Pulsatile fluid flow rapidly (within seconds) increased [Ca2+]i in MLO-Y4 osteocytes (increase in fluorescence intensity 125–367%) (Figures 3A and B). In the absence of cytokines, pulsatile fluid flow caused a transient rise in [Ca2+]i in 12.3–13.1% of the cells (Figures 3C and D). TNFα and IL-1β did not affect the mean ± SEM percentage of cells showing a transient increase in [Ca2+]i after the start of pulsatile fluid flow when added for 30 minutes before pulsatile fluid flow application (TNFα 16.6 ± 9.4%; IL-1β 11.4 ± 2.9%) (Figure 3C). After longer incubation (24 hours), TNFα and IL-1β significantly reduced the mean ± SEM percentage of cells showing a transient increase in [Ca2+]i in response to pulsatile fluid flow (TNFα 6.6 ± 2.6%; IL-1β 4.9 ± 2.5%) (Figure 3D). In summary, both cytokines negatively affected the percentage of osteocytes that show an increase in [Ca2+]i after start of pulsatile fluid flow after 24 hours of incubation.

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Figure 3. Effect of pulsatile fluid flow on intracellular calcium concentration ([Ca2+]i) in MLO-Y4 osteocytes. A, Shown is the fluorescence intensity in 2 MLO-Y4 cells before (T [time] = 0 seconds) and after (T = 100 seconds) the start of pulsatile fluid flow (original magnification × 63). B, Fluorescence intensity in the cell body transiently increased within seconds after the application of pulsatile fluid flow (indicated by the dotted line at 70 seconds), indicating a rapid rise in [Ca2+]i. Typical responses of 5 MLO-Y4 cells within 1 control experiment are shown. The second dotted line indicates cessation of pulsatile fluid flow treatment. C, Treatment with IL-1β at 1 ng/ml or TNFα at 10 ng/ml for 30 minutes did not affect the percentage of cells responding to pulsatile fluid flow with an increase in [Ca2+]i. D, Treatment with IL-1β at 1 ng/ml or TNFα at 10 ng/ml for 24 hours significantly reduced the percentage of cells responding to pulsatile fluid flow with an increase in [Ca2+]i. Values in C and D are the mean and SEM from at least 4 independent experiments. ∗ = P < 0.05 versus control. AU = arbitrary units (see Figure 1 for other definitions).

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Cytokines reduce F-actin content and affect cell stiffness of MLO-Y4 osteocytes.

In the experiments described above, we found that TNFα and IL-1β inhibit the increase in NO production and [Ca2+]i that is normally associated with the onset of pulsatile fluid flow. Since cell stiffness, which is largely determined by F-actin content, has been shown to strongly affect the NO response of MLO-Y4 osteocytes to mechanical loading (36), we investigated whether TNFα and IL-1β affect F-actin content and osteocyte stiffness. TNFα at 10 ng/ml and IL-1β at 1 ng/ml did not affect F-actin content after 30 minutes of incubation (Figure 4A). However, after 24 hours of incubation with cytokines, the F-actin content was reduced to 63% of the control values after TNFα treatment and to 68% of the control values after IL-1β treatment (Figure 4A). This large reduction in F-actin content resulted in a reduction of osteocyte stiffness. Elastic moduli, as determined by optical magnetic twisting cytometry measurements, were lower in cells treated with TNFα at 10 ng/ml for 24 hours (mean ± SEM 0.40 ± 0.06 Pa/nm) and in cells treated with IL-1β at 1 ng/ml for 24 hours (mean ± SEM 0.58 ± 0.04 Pa/nm) than in control cells (mean ± SEM 0.71 ± 0.06 Pa/nm) (Figure 4B), indicating that cytokines reduced cell stiffness.

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Figure 4. Effect of cytokine treatment on filamentous actin (F-actin) content and cell stiffness in MLO-Y4 osteocytes. A, TNFα at 10 ng/ml and IL-1β at 1 ng/ml did not affect F-actin content after 30 minutes of incubation but did reduce F-actin content after 24 hours of incubation (n = 5 per group, data obtained from 2 separate experiments). Dashed line indicates control (100%). B, Elastic moduli were lower in cells treated with TNFα at 10 ng/ml for 24 hours and in cells treated with IL-1β at 1 ng/ml for 24 hours than in control cells (n > 204 cells per group). Values are the mean and SEM. ∗ = P < 0.05 versus control. See Figure 1 for other definitions.

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Calcium inhibitors affect pulsatile fluid flow– up-regulated NO production, but do not affect cytoskeletal properties or focal adhesions.

How do cytokines reduce the NO response of MLO-Y4 osteocytes to pulsatile fluid flow if not by reducing the stiffness of the cells? Mechanical loading–induced NO production by bone cells is likely produced by the constitutive endothelial cell NO synthase (eNOS) enzyme (37, 38). The activity of eNOS depends on substrate availability and binding to calmodulin, which is calcium dependent (39). Therefore, we hypothesized that loading-induced NO production is at least partially dependent on [Ca2+]i. If this is true, the effects of TNFα and/or IL-1β on pulsatile fluid flow–induced NO production might be secondary to the effects of TNFα or IL-1β on pulsatile fluid flow–stimulated [Ca2+]i and thereby on eNOS activity. Indeed, we found that subjecting MLO-Y4 osteocytes to the calcium inhibitors TMB-8, GdCl3, and EGTA significantly inhibited the pulsatile fluid flow– up-regulated NO production after 5 minutes (Figure 5A). Maximal inhibition of the stimulatory effect of pulsatile fluid flow on NO production was observed in the presence of GdCl3 (70% decrease) and EGTA (67% decrease) (both P < 0.01), while TMB-8 resulted in a 57% decrease (P < 0.05).

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Figure 5. Effect of calcium inhibitors on focal adhesions, cell traction, and pulsatile fluid flow–induced NO production in MLO-Y4 osteocytes. A, TMB-8 (8-[diethylamino]octyl-3,4,5-trimethoxybenzoate·HCl) (n = 8 experiments), gadolinium chloride (GdCl3) (n = 7 experiments), and EGTA (n = 7 experiments) decreased pulsatile fluid flow–induced NO production by MLO-Y4 cells after 5 minutes. B, TMB-8 (n = 4 experiments), GdCl3 (n = 5 experiments), and EGTA (n = 5 experiments) did not reduce the elastic modulus. C, Shown is a traction map (values in pascals) of an osteocyte under static culture conditions without calcium inhibitors (top). Quantification of cell tractions (bottom) shows that GdCl3 slightly reduced tractions in osteocytes. D, TMB-8, GdCl3, and EGTA did not affect focal adhesion expression by MLO-Y4 cells (top). Nucleus is shown in blue; focal adhesions are shown in red (bottom) (original magnification × 40). Values are the mean and SEM. ∗ = P ≤ 0.05 versus control. See Figure 1 for explanations and other definitions.

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Since many cellular processes, such as cytoskeletal integrity and cell attachment via integrins, are dependent on the availability of calcium, we cannot exclude the possibility that our calcium inhibitors affected more cellular features involved in the NO response to pulsatile fluid flow than just eNOS. Therefore, we investigated whether TMB-8 (20 μM), GdCl3 (10 μM), and/or EGTA (2 mM) affected cell stiffness, cytoskeletal tractions, and the number of focal adhesions. We found that elastic moduli were unaffected following calcium inhibition (Figure 5B). Traction maps showed that traction stresses mainly occurred at the tips of the cell extensions (Figure 5C). Cytoskeletal tractions represented by the contractile moment were unaffected following EGTA or TMB-8 treatment (P > 0.1) (Figure 5C) and were marginally smaller after incubation with GdCl3 (P = 0.05) (Figure 5C). Neither GdCl3, EGTA, nor TMB-8 affected the number of focal adhesions per MLO-Y4 cell (Figure 5D).

Cytokines affect osteocyte apoptosis.

Treatment with TNFα or IL-1β for 30 minutes did not significantly affect caspase 3/7 activity in MLO-Y4 osteocytes (Figure 6A), suggesting that these cytokines did not cause apoptosis. Application of TNFα (0.5–30 ng/ml) for 24 hours did not increase caspase 3/7 activity in osteocytes (Figure 6B), while treatment with IL-1β (0.1–10 ng/ml) for 24 hours increased caspase 3/7 activity in a dose-dependent manner (Figure 6B), indicating that this cytokine can induce apoptosis in osteocytes.

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Figure 6. Effect of cytokine treatment on apoptosis of MLO-Y4 osteocytes. A, Treatment with TNFα or IL-1β for 30 minutes did not significantly affect caspase 3/7 activity in MLO-Y4 osteocytes. B, Application of TNFα (0.5–30 ng/ml) for 24 hours did not increase caspase 3/7 activity in osteocytes, while treatment with IL-1β (0.1– 10 ng/ml) for 24 hours dose-dependently increased caspase 3/7 activity. Values are the mean and SEM (n = 3 independent experiments). ∗ = P < 0.05 versus no treatment. AU = arbitrary units (light units per ng DNA) (see Figure 1 for other definitions).

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DISCUSSION

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. AUTHOR CONTRIBUTIONS
  7. Acknowledgements
  8. REFERENCES

Proinflammatory cytokines such as TNFα and IL-1β have been suggested to be involved in the reduction of bone mass that is observed during inflammatory diseases such as RA, but the exact mechanism is not clear. Osteocytes regulate bone mass by dictating the balance between bone formation and resorption in response to mechanical loading (17). Therefore, any factor altering the response of osteocytes to mechanical loading potentially affects bone mass. We hypothesized that the cytokines TNFα and/or IL-1β alter the mechanotransduction response of osteocytes to fluid flow. We found that both TNFα and IL-1β inhibit the up-regulation of NO production after mechanical stimulation by pulsatile fluid flow. These results are in accordance with the finding that IL-1 also reduces mechanical loading–induced NO production by dynamically compressed chondrocytes (40). The inhibition of pulsatile fluid flow–induced NO production was associated with a prevention of fluid shear stress–induced [Ca2+]i up-regulation, which is consistent with reported data that TNFα decreases [Ca2+]i peaks in contracting adult cardiocytes (41) and in mechanically stimulated brain endothelial cells (42). Our findings provide a novel mechanism by which cytokines might interfere with bone remodeling during inflammatory diseases.

The frequency spectra after loading of the hip bone in living humans have been calculated by Bacabac et al (43). Their findings were based on force measurements in the human hip. The frequency spectra of the measured forces on the hip showed a rich harmonic content ranging between 1 and 3 Hz for walking cycles and reaching 8 to 9 Hz for running cycles. The fluid shear stress amplitude around osteocytes, resulting from daily mechanical loads, has been determined theoretically by applying Biot's theory of poroelasticity to bone. The predicted range of fluid shear stress in vivo ranges from 0.8 to 3 Pa due to physiologic strains in humans (44). Although both the accuracy and the relevance of these calculations are currently under debate, it is noteworthy that bone cells in vitro seem to be highly sensitive to shear stress on this order of magnitude. Our stimulus of 0.7 ± 0.3 Pa at 5 Hz is thus based on the currently available knowledge on physiologic bone-loading stimuli.

Both TNFα and IL-1β are known to promote apoptosis of osteocytes (45, 46). In the present study, the cytokines did not affect apoptosis of MLO-Y4 osteocytes when the cells were incubated with TNFα or IL-1β for a limited time period of 60 minutes. Therefore, the reduction in pulsatile fluid flow–up-regulated NO production by bone cells that were treated for 30 minutes with TNFα or IL-1β was not due to cytokine-mediated osteocyte apoptosis. We did not find an effect of 24 hours of incubation with 10 ng/ml TNFα on caspase 3/7 activity in MLO-Y4 osteocytes, contrary to what was previously shown by Tan et al (46). This might be explained by their use of serum-free culture medium, since MLO-Y4 osteocytes seem extremely sensitive to serum deprivation. In contrast to TNFα, IL-1β dose-dependently increased caspase 3/7 activity after 24 hours of incubation, indicating that longer incubation times with IL-1β induce osteocyte apoptosis. Importantly, this could provide an additional mechanism by which osteocytes contribute to bone loss, since apoptotic osteocytes have been shown to stimulate osteoclastogenesis (47, 48).

Since TNFα did not cause osteocyte apoptosis and IL-1β increased apoptosis only when applied for 24 hours, the question remains of how these cytokines prevented the significant up-regulation of NO production by pulsatile fluid flow. The answer might be sought in the abrogation of the pulsatile fluid flow–induced rise in [Ca2+]i by TNFα and IL-1β. A rise in [Ca2+]i activates many molecules that are calcium/calmodulin dependent. One such enzyme is constitutive eNOS, which requires binding of calcium/calmodulin to achieve maximum NO production (39). Of the different isoforms of NOS (i.e., eNOS, inducible NOS, and neuronal NOS), eNOS is the isoform likely responsible for the pulsatile fluid flow–induced up-regulation of NO production by osteocytes (38). This up-regulation of NO production in response to pulsatile fluid flow might thus be dependent on increased [Ca2+]i. By using GdCl3, TMB-8, and EGTA, we observed that inhibition of [Ca2+]i resulted in a decrease in pulsatile fluid flow–induced NO production by osteocytes. This suggests that the pulsatile fluid flow–induced NO production by osteocytes depends at least partially on Ca2+. Our results are supported by other findings that Ca2+-dependent NO production occurs in whole bones loaded ex vivo, where inhibition of calcium channels by GdCl3 inhibits the loading-induced NO production (49). Ca2+-dependent NO production resulting from mechanical loading has also been observed in osteoblasts (50). Thus, TNFα and IL-1β might affect the pulsatile fluid flow–stimulated NO production at least in part by decreasing [Ca2+]i, thereby inhibiting eNOS activity.

One might argue that inhibition of [Ca2+]i with GdCl3, TMB-8, or EGTA will affect integrins and the cytoskeleton. The cytoskeleton strongly affects the mechanical properties of a cell, which in turn affect the intrinsic ability of the osteocyte to sense mechanical loading (36). Any agent that affects integrins and/or the cytoskeleton would thus potentially affect the NO response of cells to mechanical loading by altering the mechanosensitivity of a cell. Therefore, we tested whether GdCl3, TMB-8, and/or EGTA affects the number of focal adhesions, cell stiffness, and/or basal cell tractions in osteocytes, but we found no such effects in MLO-Y4 cells. This suggests that the inhibition of pulsatile fluid flow–stimulated NO production by calcium inhibitors likely occurs via inhibition of eNOS activity rather than via cytoskeletal changes.

It has been reported that exposure of articular chondrocytes to 10 ng/ml IL-1β for 1 hour slightly increases intracellular F-actin content (30). Using an experimental setup similar to that described by Pritchard and Guilak (30), we found a significant decrease in F-actin content after 24 hours of incubation with TNFα as well as with IL-1β. This reduction in F-actin content was associated with a reduction in elastic modulus. Thus, it is possible that TNFα or IL-1β affected osteocyte stiffness and thereby the intrinsic ability of osteocytes to sense mechanical loading (36).

In conclusion, we have now identified a novel pathway by which cytokines might affect local and systemic bone mass (i.e., by reducing the physiologic response of osteocytes to mechanical loading). One of the consequences of this reduction in mechanoresponsiveness would be a disruption in osteocyte-mediated balance between osteoblast and osteoclast activity. Based on these findings, we propose that reduction of cytokine levels should be a priority in RA, not only to reduce the local deleterious effects of inflammation, but also to reduce the systemic effects on the skeleton.

AUTHOR CONTRIBUTIONS

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. AUTHOR CONTRIBUTIONS
  7. Acknowledgements
  8. REFERENCES

All authors were involved in drafting the article or revising it critically for important intellectual content, and all authors approved the final version to be published. Dr. Klein-Nulend had full access to all of the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

Study conception and design. Bakker, da Silva, Bacabac, Cirelli, Klein-Nulend.

Acquisition of data. Bakker, da Silva, Krishnan, Blaauboer, Lin, Marcantonio.

Analysis and interpretation of data. Bakker, da Silva, Krishnan, Klein-Nulend.

Acknowledgements

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. AUTHOR CONTRIBUTIONS
  7. Acknowledgements
  8. REFERENCES

The authors wish to thank J. M. A. de Blieck-Hogervorst for excellent technical assistance.

REFERENCES

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. AUTHOR CONTRIBUTIONS
  7. Acknowledgements
  8. REFERENCES
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