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Keywords:

  • melatonin;
  • bone resorption;
  • osteoclast;
  • RANKL;
  • osteoprotegerin;
  • osteoclastogenesis (mouse)

Abstract

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

This study evaluated if melatonin would increase bone mass in mice. Four groups of 4-week-old male ddy mice received daily injections of vehicle or 1, 5, or 50 mg/kg of melatonin, respectively, for 4 weeks. Treatment with 5 mg/kg per day or 50 mg/kg per day of melatonin significantly increased bone mineral density (BMD; by 36%, p < 0.005) and bone mass (bone volume per tissue volume [BV/TV] by 49%, p < 0.01, and trabecular thickness [Tb.Th] by 19%, p < 0.05). This treatment significantly reduced bone resorption parameters (i.e., osteoclast surface [Oc.S/bone surface {BS}] by 74%, p < 0.05, and osteoclast number [N.Oc/BS] by 76%, p < 0.005) but did not increase histomorphometric bone formation parameters (i.e., bone formation rate [BFR/BS], mineral apposition rate [MAR], and osteoid volume [OV/TV]), indicating that melatonin increases bone mass predominantly through suppression of bone resorption. Melatonin (1–500 μM) in vitro caused dose-dependent reduction (p < 0.001 for each) in the number and area of resorption pits formed by osteoclasts derived from bone marrow cells but not those formed by isolated rabbit osteoclasts. Because RANKL increases, while osteoprotegerin (OPG) serves as a soluble decoy receptor for RANKL to inhibit osteoclast formation and activity, the effect of melatonin on the expression of RANKL and OPG in mouse MC3T3-E1 osteoblastic cells was investigated. Melatonin (5–500 μM) increased in a dose-dependent manner and reduced the mRNA level of RANKL and both mRNA and protein levels of OPG in MC3T3-E1 cells (p < 0.001 for each). In summary, these findings indicated for the first time that melatonin at pharmacologic doses in mice causes an inhibition of bone resorption and an increase in bone mass. These skeletal effects probably were caused by the melatonin-mediated down-regulation of the RANKL-mediated osteoclast formation and activation.


INTRODUCTION

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

THE SECRETION of melatonin, the major pineal hormone, is regulated by neuronal inputs from the suprachiasmatic nucleus(1) and displays a distinct daily rhythm that is linked with sleep. The plasma concentration of melatonin at night is 10-50 times higher than that during the daytime.(2) Melatonin plays a critical role in the hypothalamic regulation of circadian rhythms(1,3,4) such as regulation of reproductive function in seasonally breeding species,(5) phase-shifting effects,(4) and body temperature.(6) These circadian rhythms are related closely to sleep initiation and maintenance.(7) Melatonin also has regulatory actions on sexual activity and development,(8,9) immunomodulation,(10) and cardiovascular functions;(11) is a potent antioxidant;(12) and has been suggested to possess antiaging(13) and oncostatic(14) properties. Melatonin also influences the release of growth hormone in the human(15,16) and corticosterone in the rat.(17) Hence, in addition to its role in processing of photoperiodic information and regulation of various biological rhythms, melatonin also may be involved in a wide range of physiological and pathological processes.

There is considerable interest in the potential role of melatonin in osteoporosis.(18–20) Accordingly, the secretion of melatonin declines progressively with increasing age(21) and menopause is time-related with a substantial and sharp decrease in melatonin secretion and an associated increase in the rate of pineal calcification.(21–23) Because osteoporosis is uncommon among blacks, pineal calcification (which reflects the secretory activity of the gland) also is rare in the black population.(24) Melatonin secretion declines with immobilization (which causes loss of bone mass)(25) but increases with physical exercise (which would increase bone mass).(26) Ovariectomy leads to a significant increase in melatonin secretion in rats, which was associated with a large decrease in bone resorption biochemical markers and a much smaller increase in bone formation biochemical markers.(19) A recent report also shows that obese women (who have a low risk for osteoporosis) have a higher daytime secretion of melatonin compared with nonobese women and that the melatonin level in the obese women is associated with a reduction in serum bone turnover markers.(20) In addition, there is evidence that melatonin is an important modulator of calcium and bone metabolism. For instance, phototherapy, which reduced melatonin secretion, caused hypocalcemia in newborn rats(27) and this phototherapy-induced hypocalcemia was prevented completely by melatonin treatment.(28) The effects of melatonin on calcium metabolism presumably are mediated indirectly by influencing activity of parathyroids and secretion of calcitonin as well as prostaglandins.(29–31) However, the findings that bone marrow cells of mice and humans are capable of synthesizing melatonin(32) and that high concentrations of melatonin are found in bone marrow (∼100 times higher than that in serum) in the rat(33) suggest that melatonin may have local regulatory actions in bone. Consistent with the possibility of direct bone actions of melatonin, we, in human bone cells,(34) and others, in mouse bone cells,(35) recently reported that melatonin acts directly on osteoblasts to stimulate cell proliferation, activity (i.e., type I collagen synthesis) and bone nodule formation activity. Thus, there is circumstantial evidence that melatonin may alter bone metabolism.

In this study, we sought to test if daily melatonin administration in young mice would lead to an increase in bone mass and if the effect of melatonin on bone mass is the result of stimulation of bone formation and/or inhibition of bone resorption. We found that a daily injection of 5 mg/kg or 50 mg/kg of melatonin for 4 weeks significantly increased bone mineral density (BMD), bone mass, and trabecular bone volume in mice. Bone histomorphometry reveals that the melatonin treatment led to a significant decrease in the number of active osteoclasts on bone surface (i.e., osteoclastogenesis), without a significant effect on osteoblast number (N.Oc/BS) and bone formation rate (BFR). Because the receptor activator of NF-κB ligand (RANKL) is a potent stimulator of osteoclastogenesis(36) and because osteoprotegerin (OPG), which serves as a soluble decoy receptor for RANKL, is a potent inhibitor of osteoclastogenesis,(37) we also determined the effects of melatonin on expression of RANKL and OPG in mouse osteoblasts in vitro. We found that melatonin induced the expression of OPG and suppressed the expression of RANKL in vitro. These findings suggest that the melatonin-mediated inhibition of bone resorption is, at least in part, the result of the melatonin-induced down-regulation of the RANKL-mediated osteoclast formation and activation.

MATERIALS AND METHODS

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

Materials

DMEM and α-modified essential medium (α-MEM) were purchased from Life Technologies, Inc. (Gaithersburg, MD, USA). FBS and iron-supplemented bovine calf serum (CS) were purchased from Hyclone (Logan, UT, USA). Melatonin and tetracycline were products of Sigma Chemical Co. (St. Louis, MO, USA). Herring sperm DNA was from Kirkegaard & Perry Laboratories (Gaithersburg, MD, USA). Murine OPG cDNA probe (pCR-Blunt, an 890-bp EcoRI restriction fragment) was a generous gift of Dr. X. Qin of Loma Linda University (Loma Linda, CA, USA). Goat anti-OPG, anti-actin, and horseradish peroxidase-conjugated anti-goat immunoglobulin G (IgG) antibodies were obtained from Santa Cruz, Biotechnology, Inc. (Santa Cruz, CA, USA). The sense and antisense oligodeoxynucleotide primers of GAPDH were purchased from Clontech Laboratories, Inc. (Palo Alto, CA, USA). The TRI Reagent was a product of Molecular Research Center, Inc. (Cincinnati, OH, USA). Reagents for reverse-transcription polymerase chain reaction (RT-PCR) were from Life Technologies, Inc. AmpliTaq Gold DNA polymerase was from Roche Molecular Systems, Inc. (Branchburg, NJ, USA). Enhanced chemiluminescence detection reagents, the random primer labeling kit, and ProbeQuant G-50 microcolumns were products of Amersham Pharmacia Biotech, Inc. (Piscataway, NJ, USA). Nitrocellulose, nylon, and polyvinylidene difluoride (PVDF) membranes were purchased from Osmonics (Westborough, MA, USA). The [32P]deoxycytosine triphosphate (dCTP) was obtained from Perkin Elmer (Boston, MA, USA). All other chemicals were purchased from Fisher Co. (Tustin, CA, USA) or Sigma Chemical Co.

Animals

Twenty-four male ddy mice of 4 weeks of age were purchased from Sankyo Lab Service (Tokyo, Japan). The mice were housed in a group of 3 in plastic cages in a temperature- and light-controlled (12 h light/12 h dark) room. The mice were fed ad libitum a standard laboratory rodent dried pellet chow (Oriental Yeast Co., Tokyo, Japan). Deionized water was available to the animals at all times.

Experimental Protocol

The animal use protocol has been reviewed and approved by the Animal Care and Use Committee of the Health Sciences University of Hokkaido. The mice were randomly divided into four treatment groups (n = 6 per group) receiving daily intraperitoneal (ip) injection of solvent vehicle control (saline) or 1, 5, or 50 mg of melatonin/kg body weight, respectively, for 4 weeks. Melatonin or vehicle alone was administered between 3 p.m. and 5 p.m. The body weight of the animals was recorded weekly. At death, the size and weight of vital organs (i.e., heart, liver, spleen, and kidney) were measured to assess potential toxicity. Serum samples were obtained for measurements of total calcium, phosphorus, ALP, and osteocalcin. Serum calcium was measured with an o-cresolphthalein complexone method.(38) Serum phosphorus was assayed with a commercial kit (phosphorus; Yatron, Tokyo, Japan). Serum ALP was assayed with p-nitrophenyl phosphate in the COBAS-FARA II analyzer,(39) and serum osteocalcin was measured with a commercial radioimmunoassay (RIA) kit (CIS Bio International, Stoughton, MA, USA).

For bone histomorphometry, each mouse received an ip injection of tetracycline hydrochloride (30 mg/kg) on 5 days and 3 days before death. At the time of death, the tibias of each animal were removed. The right tibia was fixed in 70% alcohol without decalcification. After fixation, the bones were immersed in Villanueva bone stain and embedded in methylmethacrylate. Frontal sections (7 μm thick) of proximal tibia were cut with a Jung Model K (Reichert-Jung, Heidelberg, Germany) microtome. Bone histomorphometry was performed with a semiautomatic graphic system (System Supply, Nagano, Japan). Trabecular thickness (Tb.Th), bone volume per tissue volume (BV/TV), osteoid volume (OV/TV), osteoid surface (OS/BS), osteoblast surface (Ob.S/BS), osteoclast surface (Oc.S/BS), osteoclast number (N.Oc/BS), bone formation rate (BFR/BS), and mineral apposition rate (MAR) were determined in each tibial metaphyseal bone section.

DXA

The left femur and tibia were dissected free of soft tissues. Total BMD in the tibia as well as BMD at proximal, midshaft, and distal tibia (mg/cm2) were measured with the Hologic, Inc. model 1000W DEXA (Hologic, Inc., Waltham, MA, USA) in an anterior-posterior direction with constant exposure settings using an ultra-high-resolution software program, which increased the number of lines scanned by fourfold compared with a typical scan for humans. The precision of this equipment is <1%.

Isolation of rabbit osteoclasts and resorption pit formation assay

To isolate rabbit osteoclasts, long bones of 10-day-old rabbits (Japanese White; Saitama Experimental Animal, Saitama, Japan) were minced with sterile scissors in α-MEM supplemented with 5% FBS. The bone chips were rigorously agitated with a vortex mixer, and the released bone cells were collected by centrifugation. An aliquot of unfractionated bone cells was seeded on a 100-mm tissue culture dish, which was precoated with a 0.24% collagen gel layer (Nitta Gelatin, Tokyo, Japan). The plated cells were incubated for 4 h and nonadherent cells and small bone fragments were washed off with prewarmed PBS. Multinucleated TRAP+ osteoclasts were attached to the collagen matrix and were removed from the gel matrix by a brief treatment with the 0.01% crude collagenase solution (Wako, Osaka, Japan). The released osteoclasts were washed and collected by low-speed centrifugation and resuspended in α-MEM supplemented with 5% FBS. The purity of rabbit osteoclasts was >95%. The purified osteoclasts were used immediately for this study.

The resorption pit formation assay was carried out as described previously.(40) Briefly, purified rabbit osteoclasts (200 cells) or unfractionated mouse bone marrow cells (7.5 × 104 cells) were plated in α-MEM supplemented with 1% FBS on a dentine slice (150 μm thick) placed in a 48-multiwell dish. Twenty-four hours later, the osteoclasts were treated with either the vehicle control (0.1% DMSO containing 0.01% bovine serum albumin [BSA]) or the indicated concentration of melatonin for 48 h at 37°C. Melatonin stock was dissolved in 100% DMSO and diluted in DMEM containing 0.01% BSA immediately before use. The final DMSO concentration in each assay was 0.1%. The osteoclasts then were removed by sonication and the dentine slices were stained with acid hematoxylin. The number and area of resorption pits created by osteoclasts was measured with the semiautomatic graphic system.

Bone cell cultures

Subconfluent MC3T3-E1 cells (a preosteoblastic cell line) were plated in 100-mm2 culture dishes in DMEM containing 10% CS. Twenty-four hours later, the medium was changed to fresh serum-free DMEM supplemented with 0.01% BSA and the indicated concentration of melatonin or solvent vehicle was added. Then, the mRNA level of RANKL and the protein and mRNA levels of OPG were measured as described in the following section.

Measurement of RANKL mRNA levels

The mRNA levels of RANKL were determined with a semiquantitative RT-PCR assay. Briefly, after appropriate treatments with melatonin or solvent vehicle for 5 h, total RNA of subconfluent cultures of MC3T3-E1 cells (in 100-mm-diameter culture dishes) was isolated with the TRI reagent. An aliquot (1 μg) of total RNA from each sample was used to synthesize cDNA with the ThermoScript RT-PCR kit (Life Technologies). An aliquot (1 μg) of each resulting cDNA product (in 25 μl) then was amplified with the AmpliTaq Gold DNA polymerase. Optimal conditions for each primer pair (to allow for quantitative analysis of the PCR product) were predetermined in preliminary experiments. The PCR conditions for RANKL were as follows: a total of 35 PCR cycles, each consisting of 1 minute of denaturing at 95°C, 1 minute of annealing at 60°C, and 1 minute of extension at 72°C. The MgCl2 concentration in the PCR reaction was 1.5 M. The design of RANKL primer pairs was based on Deyama et al.(41); the RANKL sense and antisense oligo primers were 5′-TAT GAT GGA AGG CTC ATG GT-3′ and 5′-TGT CCT GAA CTT TGA AAG CC-3′, respectively. Each reverse-transcribed sample (1 μg) also was PCR-amplified 24 cycles for GAPDH transcript using the primer set provided by the RT-PCR kit. Each amplified PCR product was fractionated on 2% agarose gel, stained with ethidium bromide, illuminated with UV light, and photographed. The relative density of each band was analyzed with laser densitometry and the relative level of the RANKL PCR transcript was normalized against that of the GAPDH PCR transcript.

Measurement of OPG protein and mRNA levels

The OPG mRNA level was determined with Northern blot analysis. Briefly, after subconfluent MC3T3-E1 cells were treated with melatonin or solvent vehicle for 1 h at 37°C, total RNA was isolated with the TRI reagent. The RNA was fractionated on 1.5% agarose 4-morpholine propanesulfonic acid (MOPS)/formaldehyde gel and transferred to nylon membrane. The OPG mRNA species was hybridized with a murine OPG cDNA probe, which was labeled with [32P]dCTP. Free [32P]dCTP was removed from the labeled probe with the ProbeQuant G-50 microcolumn. The relative density of the OPG mRNA band on the radioautograph was measured with scanning laser densitometry and a computer-aided ChemiImager model 4400 (Alpha Innotech, Inc., Corp., San Leandro, CA, USA) and was normalized against 18S rRNA.

The OPG protein level was assessed with Western blots. After appropriate treatment for 24 h, MC3T3-E1 cells were extracted with the radioimmunoprecipitation assay (RIPA) buffer (50 mM of Tris-HCl [pH 7.4], 1% NP-40, 150 mM of NaCl, 0.5% sodium deoxycholate, 0.1% SDS, 100 μM of sodium vanadate, 50 μM of leupeptin, 1 mM of phenylmethylsulfonyl fluoride [PMSF], and 0.001% aprotinin). An aliquot (20 μg of cellular protein) of cell extract proteins was fractionated on 10% SDS polyacrylamide gels and transferred to a PVDF membrane. The OPG protein band was identified with a polyclonal anti-OPG antibody followed by incubation with horseradish peroxidase-conjugated anti-goat IgG antibodies and visualized with enhanced chemiluminescence detection according to the manufacturer's recommended protocol. To determine protein loading, each blot was striped and reblotted against the polyclonal anti-actin antibody. The relative intensity of the OPG band on X-ray films was determined with scanning laser densitometry. The relative OPG level was normalized against that of actin and is reported as percentage of solvent vehicle-treated controls.

Statistical analysis

All data were shown as means ± SD. Differences between groups were analyzed by ANOVA followed by the Fisher's protected least significant difference (PLSD) post hoc test. Differences at p < 0.05 were considered significant.

RESULTS

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

In vivo skeletal effects of melatonin in mice

The mean body weight of the mice was not significantly different among all treatment groups during the 4 weeks of the melatonin treatment (data not shown). There was no apparent change in the gross anatomy and wet weight of several vital organs (i.e., heart, liver, spleen, and kidney) between the vehicle-treated and the melatonin-treated groups (data not shown), suggesting that daily injection of 1-50 mg/kg of melatonin for 4 weeks did not appear to have significant adverse effects.

To determine whether the melatonin treatment would increase bone density, we measured total BMD as well as BMD at proximal, midshaft, and distal regions of the tibia of the mice (Table 1). The 4-week melatonin treatment significantly increased total BMD (p < 0.01) as well as BMD at each test region of the tibia (p < 0.001 for each). However, only the increases with 5 mg/kg per day and 50 mg/kg per day but not 1 mg/kg per day of melatonin were significant (p < 0.05 for each). The two higher test doses of melatonin also significantly (p < 0.05) increased BMD in the femur (data not shown). The treatment had no effect on the length of tibia of the mouse (Table 1), indicating that melatonin did not affect the longitudinal bone growth of the growing mice.

Table Table 1.. Effect of 4 Weeks of Daily ip Administration of Melatonin on BMD and Tibial Length (Mean ± SD)
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To evaluate whether the increase in BMD was caused by an increase in bone formation and/or a decrease in bone resorption, we measured bone histomorphometric parameters on the tibial metaphysis. Table 2 shows that the melatonin treatment increased trabecular bone volume (BV/TV) and Tb.Th in a dose-dependent manner (p < 0.05 for each). As is with BMD, significant (p < 0.05 for each) increases in BV/TV and Tb.Th were only seen with the two higher doses. With respect to bone formation, the melatonin treatment at any test dose did not significantly affect BFR/BS, MAR, and OV/TV. The treatment also did not increase serum levels of two bone formation biochemical markers (ALP and osteocalcin; data not shown), supporting the contention that melatonin treatment does not significantly stimulate bone formation in vivo. On the other hand, melatonin, at the 5-mg/kg per day dosage but not the other two test doses, significantly increased OS/BS and Ob.S/BS (Table 2), raising the possibility that melatonin might have a small, biphasic effect on osteoblasts, but the effect was too small to produce a statistically significant increase in bone formation. With respect to bone resorption parameters, melatonin at each test dose significantly (p < 0.05 for each) reduced Oc.S/BS and N.Oc/BS (Table 2), indicating that melatonin inhibited osteoclastic resorption in these mice.

Table Table 2.. Effect of 4 Weeks of Daily Melatonin Treatment on Tibial Metaphyseal Bone Histomorphometric Parameters (Mean ± SD)
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Because melatonin treatment could affect calcium and phosphorus homeostasis,(29–31) which could indirectly affect bone resorption, we measured serum levels of calcium and phosphorus in these mice and found that melatonin at the test doses did not significantly affect serum calcium and phosphorus levels (data not shown).

Effect of melatonin on bone resorption activity in vitro

Because melatonin markedly reduced N.Oc/BS and Oc.S/BS in vivo, we tested whether melatonin treatment would inhibit osteoclastogenesis and/or the activity of osteoclasts in vitro with the resorption pit formation assay. The number and the size of resorption pits formed by mouse bone marrow cells (containing precursors of osteoclasts as well as osteoblasts) on dentine slices after a 48-h treatment with 1-500 μM of melatonin or solvent vehicle, respectively, were measured. Because it is likely that each resorption pit in this assay is created by a single osteoclast, the number of resorption pits formed on the dentine slice would be an index of formation of active osteoclasts from bone marrow cells (i.e., osteoclastogenesis) and the average size of the pit (i.e., pit area/pit) represents the average activity of osteoclasts.

There was a dose-dependent (p < 0.001) reduction in the number of resorption pits by the melatonin treatment, and treatment with 25 μM of melatonin or higher concentrations produced significant (p < 0.05) reduction in pit number (Fig. 1A). Treatment of mouse bone marrow cells with melatonin also markedly decreased the total pit area (Fig. 1B) as well as pit area/pit (Fig. 1C). Figure 2 are representative scanning electron micrographs of resorption pits formed by bone marrow cells after 48 h of treatment with solvent vehicle (Fig. 2A), 50 μM of melatonin (Fig. 2B), or 100 μM of melatonin (Fig. 2C), which illustrate that (1) the number of resorption pits in the melatonin-treated cultures was less than that in vehicle-treated cultures and (2) the average pit size of melatonin-treated cells also was smaller than that of vehicle-treated cells. Thus, melatonin appears to inhibit not only the formation, but also the activity of osteoclasts. In contrast, when purified rabbit osteoclasts were treated for 2 days with melatonin, none of the test doses significantly affected the number of pits, total pit area, and area/pit (Figs. 1D–1F) created by rabbit osteoclasts. These findings may suggest that the inhibitory effect of melatonin on bone resorption is not mediated through a direct action on mature osteoclasts. Because bone marrow cells also contain marrow stromal cells (precursors of osteoblasts), which have been shown to produce paracrine factors that regulate osteoclast formation and activity,(42) the inhibitory effects of melatonin on osteoclasts may be mediated through marrow stromal cells or cells of osteoblast lineage within the bone marrow cell preparation.

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Figure FIG. 1. Effects of melatonin on the (A and D) resorption pit number, (B and E) total resorption pit area, and (C and F) resorption area/pit of osteoclasts formed from (A-C) bone marrow cells and (D-F) those of isolated rabbit osteoclasts in vitro. Unfractionated primary mouse bone marrow cells (7.5 × 104 cells) or isolated rabbit osteoclasts (200 cells) were treated with the indicated concentrations of melatonin or solvent vehicle for 48 h. The resorption pit formation assay was performed as described in the Materials and Methods section. Results are shown as mean ± SD (n = 6 for each group).ap < 0.05,bp < 0.01, andcp < 0.001 compared with solvent vehicle controls (by Fisher PLSD post hoc test).

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Figure FIG. 2. Scanning electron micrographs of resorption pits created by osteoclasts formed from unfractionated primary mouse bone marrow cells after 48 h of treatment with solvent vehicle, 50 μM of melatonin, or 100 μM of melatonin, respectively.

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Effect of melatonin on RANKL and OPG expression in MC3T3-E1 cells in vitro

Because RANKL and OPG are two potent osteoblast-derived regulators of osteoclastogenesis and osteoclast activity,(36,37,43) we investigated the effects of melatonin on the expression of RANKL and OPG in a mouse osteoblast cell line MC3T3-E1 cells. To monitor the RANKL expression in MC3T3-E1 cells, we measured RANKL transcript levels with a semiquantitative RT-PCR assay after the cells were treated for 5 h with 5-500 μM of melatonin or solvent vehicle. Figure 3 shows that melatonin decreased RANKL transcript level in a dose-dependent manner by 50% and that significant reduction was observed with 10 μM or higher concentrations of melatonin. To assess the effect of melatonin on the production of OPG in MC3T3-E1 cells, we first measured OPG mRNA levels with Northern analysis, after a 1-h treatment with melatonin or solvent vehicle. Figure 4 shows that melatonin >10 μM significantly increased OPG mRNA levels in MC3T3-E1 cells. We then measured the production of OPG protein in MC3T3-E1 cells after 24-h treatment with melatonin or solvent vehicle by Western blot analysis (Fig. 5). Again, doses of melatonin of 10 μM or higher concentrations markedly increased cellular OPG protein level (to as high as 550% of solvent vehicle controls) in MC3T3-E1 cells. We also obtained similar results with normal untransformed mouse calvarial cells (data not shown), suggesting that the effects of melatonin on RANKL and OPG were not unique to transformed cells.

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Figure FIG. 3. Effect of melatonin on RANKL expression in mouse MC3T3-E1 cells in vitro. MC3T3-E1 preosteoblasts were treated with solvent vehicle or the indicated concentrations of melatonin for 5 h. RANKL expression was assessed with a semiquantitative RT-PCR assay as described in the Materials and Methods section. Top panel shows representative ethidium bromide-stained RANKL and GAPDH RT-PCR products, respectively. The bottom panel summarizes the average (from two separate experiments) relative level of the RANKL PCR product normalized against that of the GAPDH PCR (in % of solvent vehicle controls, mean ± SD)ap < 0.05,bp < 0.01, andcp < 0.001 compared with solvent vehicle controls (by Fisher PLSD post hoc test).

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Figure FIG. 4. Effect of melatonin on OPG mRNA transcript levels in mouse MC3T3-E1 cells in vitro. MC3T3-E1 preosteoblasts were treated with solvent vehicle or the indicated concentrations of melatonin for 1 h. The OPG mRNA level was measured with Northern analysis and normalized against 18S rRNA as described in the Materials and Methods section. Top panel shows the representative OPG mRNA transcript and 18S rRNA, respectively. The bottom panel summarizes the average (from three separate experiments) relative level of the OPG mRNA normalized against that of the 18S rRNA (in % of solvent vehicle controls, mean ± SD)ap < 0.05,bp < 0.01, andcp < 0.001 compared with solvent vehicle controls (by Fisher PLSD post hoc test).

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Figure FIG. 5. Effect of melatonin on OPG protein level in mouse MC3T3-E1 cells in vitro. MC3T3-E1 preosteoblasts were treated with solvent vehicle or the indicated concentrations of melatonin for 24 h. OPG protein levels were measured with Western immunoblots as described in the Materials and Methods section. Top panel shows the representative OPG and actin protein bands, respectively. The bottom panel summarizes the average (from three separate experiments) relative level of the OPG protein normalized against that of actin (in % of solvent vehicle controls, mean ± SD)ap < 0.05,bp < 0.01, andcp < 0.001 compared with solvent vehicle controls (by Fisher PLSD post hoc test).

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DISCUSSION

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

There has been circumstantial evidence that melatonin may have beneficial effects in bone.(18–20) However, although it has been reported that women taking M-Oval preparation, containing 75 mg of melatonin and a trace amount of estrogen, for 3 years showed an increase in BMD(44); definitive evidence for an increase in BMD and bone mass by melatonin treatment alone has been lacking. In this study, we demonstrated that a daily injection of young mice with 5 mg/kg or 50 mg/kg of melatonin for 4 weeks increased significantly BMD and the thickness and volume of trabeculae. Consequently, these findings provide the first conclusive evidence that melatonin treatment alone could lead to an increase in BMD and bone mass in the mouse in vivo.

Our interest in the potential role of melatonin in osteoporosis began with our discovery that melatonin stimulates human osteoblast proliferation and differentiation in vitro.(34) Hence, we originally had postulated that melatonin would increase bone formation in vivo, which then would lead to an increase in bone mass and BMD. Contrary to our original hypothesis, we found that daily administration of melatonin for 4 weeks did not have an appreciable effect on bone formation histomorphometric parameters (i.e., OV/TV, BFR/BS, and MAR). In addition, the lack of a significant increase in serum ALP and osteocalcin in response to the melatonin treatment in these mice is consistent with the premise that melatonin does not significantly stimulate bone formation in vivo. Thus, despite the fact that melatonin distinctively increases the proliferation, differentiation, and bone nodule formation activity of osteoblasts in vitro,(34,35) these findings indicate that melatonin is not osteogenic in vivo, at least in young growing mice. The lack of a stimulatory response in bone formation to melatonin in mice probably is not caused by species difference, because melatonin also stimulates the differentiation and bone nodule formation in mouse MC3T3-E1 osteoblasts in vitro.(35) On the other hand, it should be noted that melatonin, at 5 mg/kg per day, significantly increased Ob.S/BS and OS/BS. These findings are consistent with the previous in vitro findings(34,35) and raise the possibility that melatonin indeed could have a small biphasic, stimulatory effect on osteoblast proliferation or recruitment in vivo. However, because there was no corresponding increase in bone formation, we conclude that the apparent biphasic increases in Ob.S/BS and OS/BS in response to melatonin probably are too small to produce a biological consequence with respect to an increase in bone formation in vivo.

Conversely, this study clearly showed that melatonin in vivo caused a marked and dose-dependent reduction in bone resorption histomorphometric parameters (i.e., N.Oc/BS and Oc.S/BS) in mice. The conclusion that melatonin would inhibit bone resorption is supported by the findings of two recent studies, which showed that the melatonin treatment in ovariectomized rats(19) as well as in obese women(20) led to an acute and marked decrease in bone resorption markers, that is, serum cross-linked carboxyterminal telopeptide of type I collagen (ICTP) and urinary hydroxyproline. Consequently, these findings led us to conclude that the melatonin-induced increase in BMD and bone mass in mice is caused by predominantly an inhibition of bone resorption rather than an increase in bone formation.

Three noteworthy observations in this study are relevant to the molecular mechanism(s) whereby melatonin inhibits osteoclastic resorption. First, melatonin in vitro markedly reduced the number of resorption pits (an index of osteoclast formation) as well as average pit area/pit (an index of osteoclast activity) created by osteoclasts formed from bone marrow cells in a dose-dependent manner. Thus, melatonin inhibits not only osteoclastogenesis, but also the activity of osteoclasts. Second, because melatonin had no effect on the number of resorption pits and pit area/pit created by isolated rabbit osteoclasts, it follows that melatonin probably does not act directly on mature osteoclasts to inhibit their bone resorption activity nor does it appear to alter the life span of mature osteoclasts (e.g., apoptosis of osteoclasts). We are mindful that the purified osteoclasts were of the rabbit origin and, thus, we cannot rule out the possibility that the lack of an inhibition by melatonin on the activity of purified rabbit osteoclasts might be caused by species differences. However, we believe that the possibility of a species difference in the response to melatonin with respect to inhibition of bone resorption probably is remote because there is circumstantial evidence that the melatonin treatment also led to an inhibition of bone resorption in rats(19) as well as in humans.(20) Because there is evidence that the regulation of osteoclastogenesis and osteoclast activation by a number of hormones and effectors of osteoclastic resorption [e.g., parathyroid hormone (PTH), 1,25-dihydroxyvitamin D3 1,25(OH)2D3, interleukin (IL)-1, and IL-6] is mediated through indirect interactions with cells of osteoblast lineage, including marrow stromal cells,(43) these findings raise the intriguing possibility that the inhibitory effect of melatonin on osteoclastogenesis and osteoclast activity is not mediated through direct actions on mature osteoclasts, but rather, it most probably acts through interaction with other cell types in bone marrow cells such as marrow stromal cells within the bone marrow cell preparation.

The third intriguing observation relevant to the molecular mechanism of melatonin on bone resorption is that melatonin in vitro up-regulated the expression of OPG and down-regulated the expression of RANKL in mouse osteoblast line cells MC3T3-E1 cells. In this regard, RANKL, a membrane-associated member of the TNF receptor-ligand family, is expressed by cells of osteoblast origin, including marrow stromal cells,(36) and has been shown to stimulate the formation and activity of osteoclasts by binding to its receptor RANK, on the cell surface of osteoclast precursors.(43) A number of bone resorption activators such as 1,25(OH)2D3, PTH, prostaglandin E2 (PGE2), IL-1, and IL-11 have been shown to up-regulate the expression of RANKL in osteoblastic cells,(45) and, thus, it generally has been assumed that these resorption activators stimulate osteoclastic resorption through up-regulation of RANKL expression in osteoblastic cells.(36,43) Conversely, OPG is a soluble protein also secreted by cells of osteoblast lineage,(37) is structurally similar to RANK,(46) and has a high affinity for RANKL.(37,46) OPG inhibits osteoclast formation and activity by acting as a soluble decoy receptor for RANKL, rendering RANKL incapable of interacting with RANK on osteoclast precursors.(37) Hence, RANKL is a potent activator and OPG is a potent inhibitor of not only osteoclastogenesis, but also osteoclast activity.(36) Thus, a reduction in RANKL expression and/or an increase in OPG expression would lead to a reduction in both the formation and the activity of osteoclasts. Consequently, our findings that (1) melatonin inhibits osteoclastogenesis and osteoclast activity, (2) melatonin acts through marrow stromal cells to inhibit osteoclastogenesis and osteoclast activity, and (3) melatonin reduced RANKL expression and increased OPG production in osteoblasts, led us to postulate that melatonin acts primarily on cells of osteoblast origin, including bone marrow stromal cells, resulting in suppression of RANKL expression and enhancement of OPG production in osteoblastic cells. The reduced RANKL expression, along with the increased OPG production, in osteoblastic cells then would lead to a decrease in osteoclastogenesis and osteoclast activity, which results in an inhibition of osteoclastic bone resorption and eventually an increase in bone mass and BMD.

However, we should be reminded that melatonin also increased the proliferation, differentiation, and activity of osteoblasts in vitro,(34,35) but yet these in vitro anabolic effects of melatonin on osteoblasts did not translate into a significant stimulation in bone formation in vivo. Accordingly, until we can show that the melatonin treatment also down-regulated the RANKL regulatory system in vivo and also until we can establish a direct relationship between the melatonin-induced down-regulation of the RANKL regulation and the inhibition of bone resorption in vivo, the physiological significance of our in vitro findings on the melatonin-induced down-regulation of the RANKL system cannot be ascertained. Nonetheless, because our in vitro observations are entirely consistent with the general belief that down-regulation of the RANKL regulatory system generally would result in a reduction in osteoclastogenesis and osteoclast activity, we tentatively conclude that the observed inhibitory action of melatonin on bone resorption in vivo might be mediated, at least in part, through down-regulation of the RANKL regulatory system.

It has been shown that melatonin may be involved in the homeostatic regulation of calcium and phosphate metabolism. Previously, it has been postulated that a decline of melatonin secretion (e.g., associated with menopause and/or aging) would result initially in reduced plasma calcium levels, which then would lead to a compensatory increase in PTH secretion and suppression of calcitonin release. The resulting increase in circulating level of PTH and the decrease in serum level of calcitonin subsequently would lead to an increase in bone resorption.(18) Accordingly, the possibility that melatonin treatment may inhibit bone resorption through indirect actions of melatonin on PTH and calcitonin secretion via alteration in serum calcium and phosphate levels cannot be overlooked. On the other hand, although we did not measure serum PTH and calcitonin levels (because of insufficient amounts of serum) to determine whether the melatonin treatment altered the circulating levels of the two hormones, we do not favor this alternative possibility because (1) the melatonin treatment did not significantly affect serum calcium and phosphate levels and (2) melatonin acts directly on bone marrow cells in vitro to suppress osteoclast formation and resorptive activity on dentine slices.

The in vivo effective doses of melatonin (5-50 mg/kg body weight per day) to inhibit bone resorption and to increase bone mass and BMD in these mice were several orders of magnitude higher than the physiological doses of melatonin (i.e., 0.3-3 mg/day) that are usually given to patients to restore nighttime plasma melatonin levels to what they were in adult life.(47) Therefore, the observed effects of melatonin on bone resorption and BMD are not physiologically but rather pharmacologically relevant. On the other hand, inasmuch as the in vitro effective concentrations of melatonin on osteoclastogenesis and osteoclast activity (>1 μM) also are very high (also at least three orders of magnitude higher than the physiological nighttime serum levels of melatonin, which is between 0.4 and 0.9 nM(48)), it has been shown that oral administration of pharmacologic doses of melatonin (80-240 mg/day) in humans could raise the plasma concentrations of melatonin 350- to 10,000-fold to micromolar levels.(49,50) Moreover, we should note that the bone marrow concentrations of melatonin of treated animals were 100 times higher than that in serum(33); it is conceivable that the local bone concentrations of melatonin in the treated animals could be very high and reached the in vitro effective concentrations (i.e., micromolar concentrations). More importantly, although the doses of melatonin used in this study may be considered high, we should emphasize that these high doses of melatonin did not appear to cause significant harmful serious side effects to the animals in vivo, indicating that these pharmacologic doses of melatonin probably are relatively safe.

Finally, we should emphasize that the mice in this study were young (4 weeks old) and were undergoing rapid growth at the time of the study. Because the bone turnover in these rapidly growing young mice is high and favors bone formation, an inhibition of bone resorption, even without an effect on bone formation, could have a more profound effect on the overall bone mass. Hence, the possibility that the enhancing effect of melatonin on bone mass in the fast-growing young animals is much higher than that in older adult animals cannot be overlooked entirely. On the other hand, a previous clinical study has provided circumstantial evidence that daily oral intakes of 75 mg of melatonin along with trace amounts of estrogen for 3 years could lead to a significant increase in BMD in adult women,(44) suggesting that the melatonin treatment also might be effective in increasing bone mass (presumably through an inhibition of bone resorption) in adults as well as in the fast-growing young animals. Further work is needed to confirm this supposition.

In summary, this study shows, for the first time, that (1) administration of pharmacologic doses of melatonin in growing young mice increased trabecular bone mass and BMD, (2) the effects are primarily mediated through an inhibition of bone resorption, and (3) administration of pharmacologic doses of melatonin did not appear to have significant toxic side effects. These findings are clinically relevant because if these observations are confirmed in adult animals and extended to humans, it is conceivable that the melatonin therapy could be developed into an effective and safe treatment for postmenopausal osteoporosis and/or other related bone-wasting diseases.

Acknowledgements

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

This work was supported in part by research supports from the School of Dentistry of the Health Sciences University of Hokkaido and the Muscuoloskeletal Disease Center of the Jerry L. Pettis Memorial Veterans Affairs (VA) Medical Center and also by a research grant from the National Institute of Dental and Craniofacial Research (RO1 DE13097 to K.-H.W. Lau). Parts of the work were performed in facilities provided by the Department of Veterans Affairs.

The authors have no conflict of interest.

REFERENCES

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