Bone is a complex rigid organ composed of several types of cells undergoing a continuous remodeling process of bone formation and bone resorption followed by the replacement of bone to regulate calcium homeostasis and repair damaged bones (Feng and McDonald, 2011).
There are two different types of bone cells: osteoblasts, which form new bone, and osteoclasts, which break bone down, and they both play an important role in the regulation of bone metabolism (Feng and McDonald, 2011). The action of osteoblasts and osteoclasts, coupled via paracrine signaling, is controlled by a number of factors that either enhance or inhibit bone remodeling cells. The subtle balances between osteoblast and osteoclast numbers and activities mediated bone remodeling process (Kapinas and Delany, 2011) (Figure 1). Some cytokines at the remodeling site recruits osteoclasts to the bone surface (Rachner et al., 2011). A boarder formed by osteoclasts allows their tight adherence to the bone surface. The bone remodeling phase begins with mononuclear cells preparing the bone surface for new osteoblasts and providing signals to recruit them. When the osteoblasts differentiate, the matrix matures and is mineralized (Kapinas and Delany, 2011). However, the imbalanced regulation of bone remodeling between two processes—bone resorption and bone formation—leads many metabolic bone diseases, including osteoporosis (Rachner et al., 2011).
Osteoporosis is a common skeletal disease that can lead to an increased risk of fracture (caused by decreased bone mass) and an enhanced risk of bone fragility and susceptibility to
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-fracture (Sambrook and Cooper, 2006; Rachner et al., 2011). In osteoporosis, the bone mineral density (BMD) is reduced, bone microarchitecture deteriorates, and the amount and variety of proteins in bone are altered. Osteoporosis is defined as a bone mineral density of 2.5 standard deviations or more below the mean peak bone mass (average of young, healthy adults) as measured by dual-energy X-ray absorptiometry; the term "established osteoporosis" includes the presence of a fragility fracture (Kanis, 1994).
For the pharmacological therapy of postmenopausal osteoporosis, supplements and medications usually include certain ingredients to help maintain bone health: calcium, magnesium, vitamin D, vitamin K, phytoestrogen, and prebiotic fiber (Hanley et al., 2010;
Jose Ramon et al., 2012; Rizzoli et al., 2013). Novel interventions that target regulators of bone remodeling have been suggested to be promising agents for the treatment of osteoporosis. An inhibitor of cathepsin K, odanacatib, is in clinical trials for the treatment of postmenopausal osteoporosis and monoclonal antibodies to sclerostin, such as AMG 785, have osteoanabolic properties with the potential to improve clinical outcomes in patients with osteoporosis (Lewiecki, 2011). However, some of these medications have side effects, including long-lasting ingestion and an increased risk of endometrial and breast cancers (Bonura, 2009). Therefore, the number of ingredients should be minimized in order to decrease the side effects of medications used to treat osteoporosis.
Recently, as an alternative long‐term therapeutic option against osteoporosis, herbal medicine has come to our attention. This is because the advantages of natural plants have shown the potential of having fewer side effects, thereby making them more suitable for long‐term use. As a potential alternative treatment for osteoporosis, the chemopreventive and
chemotherapeutic effects of several natural products derived from plants, including Carthamus tinctorius, Drynaria fortunei, Gardenia jasminoides, Schizandra chinensis, and Ulmus davidiana, have been reported (Ha et al., 2003; Suh et al., 2007; Kim et al., 2008;
Caichompoo et al., 2009; Jeong et al., 2012). Several natural products, such as Curculigoside (Shen et al., 2013), Withania somnifera (Nagareddy and Lakshmana, 2006), Pueraria mirifica, and miroestrol (Chatuphonprasert et al., 2013), have also been reported to prevent bone loss in osteoporosis animal models with an improvement in total body bone mineral density and bone mineral content.
In this study, my aim was to discover alternative herbal therapeutic drugs for effective osteoporosis treatment in vitro and in vivo. Sixty-four ethanol extracts of edible plants native to Korea were screened for their ability to increase the proliferation and differentiation of two osteoblast cell lines. Based on the screening data of both cell lines, Lycii Radicis Cortex (LRC) extract was selected as a potential natural‐source candidate. I carried out further in vitro experiments in cell lines and in vivo experiments in the osteoporosis model mice to evaluate the effects of LRC extract on bone formation.
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-Figure 1. Bone remodeling process (Kapinas and Delany, 2011). Bone remodeling begins when osteoclasts resorb bone mineral and matrix. Mononuclear cells prepare the resorbed surface for osteoblasts, which generate newly synthesized matrix as they differentiate.
Matrix mineralization and the differentiation of some osteoblasts into osteocytes completes the remodeling cycle.
II. MATIRIALS AND METHODS
1. Herbal extracts and materials.
The 70% ethanol extracts of 64 plants native to Korea were provided from the Korea Promotion Institute for Traditional Medicine Industry (Gyeongsan, Korea) (http://www.kotmin.kr). The plant materials were added 5-fold volume of 70% ethanol, extracted at 100 °C for 3 h, filtered, and then concentrated. Cell culture media (DMEM and α-MEM), antibiotics (penicillin and streptomycin), and fetal bovine serum (FBS) were purchased from Invitrogen (Carlsbad, USA). An EZ-Cytox Cell Viability Assay Kit was purchased from Daeil (Seoul, Korea), and ascorbic acid and β-glycerophosphate were purchased from Sigma-Aldrich (St. Louis, USA).
2. Cell culture.
The murine mesenchymal stem cell line (C3H10T1/2 cells) was purchased from the Korean Cell Line Bank (Seoul, Korea) and grown in a DMEM medium supplemented with 10% FBS, penicillin (100 U/ml), and streptomycin (100 μg/ml). Mouse MC3T3-E1 pre-osteoblast cells were purchased from the RIKEN Cell Bank (Tsukuba, Japan) and grown in a α-MEM medium supplemented with 10% FBS, penicillin (100 U/ml), and streptomycin (100 μg/ml). Osteoblast differentiation was induced by adding an osteogenic medium containing ascorbic acid (50 μg/ml) and β-glycerophosphate (10 mM) after allowing 24 h for cell
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-adherence (day 0), and the medium was changed every three days. All cultured cells were incubated in a humidified atmosphere at 37 °C and at 5% CO2.
3. Water-soluble tetrazolium salt (WST) assay and alkaline phosphatase (ALP) assay.
To test cell viability, cells (3×103 cells/well) were incubated in a 96-well plate overnight and treated with different concentrations of the plant extracts (10, 50 and 100 μg/ml) in the medium for 48 h. An EZ-Cytox Cell Viability Assay Kit was used for cell viability assay. WST (20 μl, 5 mg/mL in PBS) was added to each well, the cells were incubated for another 4 h, and the media were carefully removed. Absorbance was measured at 450 nm and 655 nm using a microplate reader (BioTek; Winooski, USA). For ALP activity assay, cells (1.5×10⁴cells/well) were incubated in a 48-well plate for 24 h and then treated with different concentrations of the plant extracts (10, 50 and 100 μg/ml) in the medium for 48 h. After washing twice with 1×PBS, the cells were added 200 μl of extraction solution and incubated overnight at 4 °C. ALP activity was measured in total cell lysates after homogenization in a buffer containing 1 mmol/l Tris-HCl (pH 8.8), 0.5% Triton X-100, 10 mmol/l Mg2+, and 5 mmol/L p-nitrophenylphosphate as substrates. The absorbance was read at 405 nm (BioTek).
4. In vitro generation of osteoclasts and tartrate-resistant acid phosphatase (TRAP) assay.
For primary-cultured monocytes, bone marrow cells were flushed from the femoral bones of 6-week-old mice in the presence of ascorbate-2-phosphate (1 mM). Monocyte cells
were identified by immunophenotypic analysis with a CD11b antibody using the FACS Aria III cell sorter (BD Biosciences, San Jose, USA) and FACS Diva software (BD Biosciences).
Monocyte cells were cultured in the presence of 30 ng/ml M-CSF (PeproTech, Rocky Hill, USA) and 50 ng/ml RANKL (PeproTech) to induce differentiation to osteoclasts (Sun et al., 2006). The differentiated osteoclast cells from monocytes were measured by a TRAP activity assay and staining using the Acid-Phosphatase Kit (Sigma-Aldrich).
5. Quantitative reverse-transcription PCR (qRT-PCR).
To determine the mRNA expression levels of the bone metabolic genetic markers, Alp, Runx2, Ocn (Osteocalcin, Bglap), quantitative real-time PCR was performed. Total RNA
was extracted from culture cells using a TRIzol reagent (Invitrogen, Carlsbad, USA) following the manufacturer’s instructions and quantified by a spectrophotometer (Beckman Coulter, Brea, USA). The extracted RNA were subsequently reverse transcribed using the RevertAid™ H Minus First Strand cDNA Synthesis Kit (Fermentas Inc., Hanover, USA) with the oligo(dT)15–18 at a random primer. All real-time PCR measurements were performed using the ABI Prism 7000 Sequence Detection System (Applied Biosystems;
Foster City, USA). All PCR amplifications (40 cycles) were performed in a total volume of 25 μl containing 150 ng cDNA using the SYBR Green I qPCR kit (TaKaRa, Shiga, Japan) according to the manufacturer’s recommendations. The specific primers for osteoblast markers were as follows: 5′-TCCCACGTTTTCACATTCGG-3′ and 5′-GGCCATCCTATA TGGTAACGGG-3′ for mouse Alpl, 5′-TAAAGTGACAGTGGACGGTCCC-3′ and 5′-CCT CAGTGATTTAGGGCGCA-3′ for mouse Runx2, and 5′-TAGTGAACAGACTCCGGCG
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-CTA-3′ and 5′-AT GGCTTGAAGACCGCCTACA-3′ for mouse Bglap. To normalize the efficiency of real-time RT-PCR reactions, the mouse Gapdh gene was used as a relative quantification standard with the following primers: 5′-TGACCACAGTCCATGCCATC-3′
and 5′-GACGGACACATTGGGGGTAG-3′. The following amplification parameters were used: 10 min preincubation for hot start polymerase activation at 95°C, followed by 45 amplification cycles at 95 °C for 20 s, 62 °C for 20 s, 72 °C for 40 s. After the end of the last cycle, the melting curve was generated by starting fluorescence acquisition at 60 °C, and taking measurements every 0.2 °C until 95 °C. By normalizing to Gapdh, a relative quantification of gene expression was performed using the comparative threshold (Ct) method as described by the manufacturer (Applied Biosystems). The values were expressed as fold change over control. Relative gene expression was displayed as 2-ΔCt (ΔCt = Ct target gene − Ct Gapdh). Fold change was calculated as 2-ΔΔCt (ΔΔCt = ΔCt control − Ct treatment).
6. In vivo experiment.
The ovariectomized (OVX, n = 30) and sham-operated (Sham, n = 12) 8-week-old female ddY mice were purchased from Shizuoka Laboratory Center Inc. (Hamamatsu, Japan). The mice were acclimated for 10 days prior to experimentation. They were maintained on a diet of Formula-M07 (5.0 g/day) (Feedlab Co. Ltd., Hanam, Korea) and tap water (15 ml/day). All mice were housed individually in clear plastic cages under controlled temperature (23 ± 2 °C), humidity (55 ± 5%), and illumination (12-hmy light/dark cycle).
The mice were administered different concentrations of LRC extract: either 50, 150 and 300
mg/kg/day for 8 weeks or 150 mg/kg/day for 16 weeks. Each calculated concentration of LRC extract was added to tap water. The LRC extract-containing water was changed for fresh water every three days, and the volume of LRC extract-containing water was measured every three days for the administered LRC amount. The right femur bone mineral density (BMD) and bone mineral content (BMC), as well as the whole body percentage of body fat (% fat) and body weight, were measured before and after the administration of LRC extract.
The animal research protocol was approved by the Animal Care and Use Committee of the Ajou University School of Medicine, and all experiments were conducted in accordance with the institutional guidelines established by the Committee.
7. Measurement of whole body fat (%) and right femur BMD and BMC.
Whole body fat (% fat) and right femur BMD and BMC were measured using a PIXImus bone densitometer (GE Lunar, Madison, USA) and calculated by on-board PIXImus software for small animals, adjusted in relation to body weight. After anesthetization using tiletamine/zolazepam (Zoletil; Virbac Laboratories, Carros, France), the mice were placed on the specimen tray for measurement.
8. Statistical analysis.
A statistical software package (SPSS 11.0 for Windows, SPSS Inc., Chicago, USA) was used to perform the statistical tests. The statistical significance of differences was assessed by Student’s t-test. P < 0.05 values were considered significant. Results were expressed as mean ± SEM.
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III. RESULTS
1. Sixty-four plants native to Korea were screened for cellular proliferation and the differentiation of osteoblastic C3H10T1/2 and MC3T3-E1 cell lines.
To discover the potential natural source(s) having an influence on bone formation augmentation, I screened the ethanol extracts of 64 different plants native to Korea. ALP is a glycoprotein found on the surface of osteoblasts and is a sensitive and reliable indicator of bone metabolism. Hence, I used an ALP assay as the main method for screening the extracts’
effects on bone formation. Three different concentrations (10, 50 and 100 μg/ml) of each plant extract were treated in the osteoblastic cell lines C3H10T1/2 and MC3T3-E1. After three days of incubation with the 64 extracts, cell viability was examined by a WST assay, and osteoblastic differentiation levels were determined using the ALP assay (Table 1).
Based on the ALP activity and cell viability of the two cell lines, Lycii Radicis Cortex (LRC), Lycium Chinense root bark, was selected as a primary candidate plant. LRC is widely used in eastern Asia as a traditional medicine. However, the effect of LRC extract on osteoporosis has not yet been explored.
Table 1. Results of water-soluble tetrazolium salt (WST) assay and alkaline
Symbols: ---, 0～50%; --, 50～70%; -, 70～90%; ., 90～110%; +, 110～130%; ++, 130～150%; +++, 150～ vs. non-treated control.
22 Bombyx Batryticatus 44 Salicornia Herbacea
21 Ulmi Cortex 43 Dipsaci Radix
20 Radix Achyrantis 42 Cortex
Acanthopanacis 64 Luffa Cylindrica
19 Thujae Semen 41 Albizziae Cortex 63 Bulbus Allii
18 Chinemys Reevesii
(Gray) 40 Herba Siegesbeckiae 62 Hovenia Dulcis
17 Astragali Radix 39 Artemisiae Argyi
Folium 61 Prunus Mume
16 Saururus Chinensis 38 Puerariae Radix 60 Vitis Vinifera
15 Cervi Cornu 37 Semen Torreyae 59 Aralia Elata
14 Herba Taraxaci 36 Fagopyrum
Esculentum 58 Hypsizigus
Marmoreus
13 Sorbus Commixta 35 Herba Artemisiae
Annuae 57 Inonotus Obliquus
12 Cornus Officinalis 34 Rubi Fructus 56 Laminaria Japonica
Areschoung
11 Codonopsis Pilosula 33 Radix Sophorae 55 Nelumbinis
Rhizomatis Nodus
10 Rhizoma Cibotii 32 Houttuyniae Herba 54 Aurantii Immatri
Pericarpium
9 Onion 31 Mori Folium 53 Citrus Unshiu Peel
8 Carthamus
Tinctorius L. 30 Ramulus Mori 52 Glycine Semen
Preparatum
7 Aralia Continentalis 29 Herba Cirsii 51 Caragana Sinica
6 Kalopanacis Cortex 28 Rosae Laevigatae
Fructus 50 Fructus Aurantii
Immaturus
5 Eucommia Ulmoides 27 Morus Alba L. 49Curcuma Aromatica
Salisb.
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2. LRC extract increased cellular proliferation and differentiation of osteoblast cell lines.
I further investigated the anti-osteoporotic effects of LRC extract. Treatment using 10 μg/mL and 50 μg/mL of LRC extract, respectively, showed a significantly increased proliferation in both cell lines compared to the control group, indicating that LRC extract has no toxic effects on cell growth (Figure 2A). The highest ALP activity in both C3H10T1/2 and MC3T3-E1 cell lines was observed in 10 μg/ml extracts of LRC (Figure 2B).
Imbalanced bone remodeling is one of the key factors in inducing osteoporosis and many other metabolic bone diseases. Naturally, the daily removal of bone mineral (bone resorption) must be balanced by equal amounts of new mineral deposition, which leads to gradual restructuring of the bone. I further examined whether LRC extract has any effect on the differentiation of osteoclast cell lines. For primary-cultured monocytes, bone marrow cells were flushed from the femoral bones of 6-week-old mice in the presence of ascorbate-2-phosphate. Monocyte cells were identified by immunophenotypic analysis with monocyte-specific surface markers (CD11b antibody) using fluorescence activated cell sorter (FACS) analysis (Figure 3A). The TRAP assay results revealed that osteoclastic differentiation of the primary-cultured monocytes was significantly increased by induction with 30 ng/ml of M-CSF and 50 ng/ml of RANKL, but there was no changes by additional treatment of 10 μg/ml of LRC extract in the M-CSF and RANKL-treated monocytes (Figure 3B, 3C). These results suggested that LRC extract may promote the proliferation and differentiation of osteoblast cells rather than the inhibition of osteoclastic differentiation.
Figure 2. Lycii Radicis Cortex (LRC) extract increased cellular proliferation and differentiation in C3H10T1/2 and MC3T3-E1 osteoblastic cell lines. Cells were cultured with three different concentrations of LRC extract (10, 50 and 100 μg/ml) for three days, and cell viability (A) and ALP activity (B) were analyzed. Control: LRC non-treated cells. *, #: p
< 0.05 vs. Control.
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Figure 3. No changes in osteoclastic differentiation after treatment of Lycii Radicis Cortex (LRC) extract in primary-cultured monocytes. (A) Successfully isolated
monocytes from mouse bone marrows were identified by immunophenotypic analysis with monocyte-specific surface markers (CD11b antibody) using FACS analysis. (B, C) Monocyte cells were cultured in the presence of 30 ng/ml of M-CSF and 50 ng/ml of RANKL (Induction), or M-CSF and RANKL with 10 μg/ml of LRC extract (LRC). The differentiated osteoclast cells from mouse bone marrow monocytes were measured by tartrate-resistant acid phosphatase (TRAP) activity assay (B) and TRAP staining (C).
Control: Monocyte cells cultured without M-CSF and RANKL. *: p < 0.05 vs. Control.
3. LRC extract increased mRNA expression of osteoblastic markers.
To further confirm the effect of LRC on the cellular differentiation of osteoblasts, I investigated any changes in the expression of representative osteoblastic marker genes Alpl (alkaline phosphatase, ALP), Runx2 (runt-related transcription factor 2, Runx2), and Bglap (bone gamma carboxyglutamate protein, Osteocalcin). After treatment with 10 μg/mL of LRC extract for three days in C3H10T1/2 and MC3T3-E1 cell lines, total RNAs were prepared and used as a template for quantitative RT-PCR.
There was significant increased expression of Alpl, Runx2, and Bglap genes in both LRC-treated cell lines compared to the non-treated control cell lines (Figure 4). This result suggested that LRC extract may stimulate osteoblast differentiation by up-regulating osteoblastic-inducing genes such as Alpl, Runx2, and Bglap.
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Figure 4. Lycii Radicis Cortex (LRC) extract increased the mRNA expression of Alpl, Runx2, and Bglap in C3H10T1/2 and E1 cells. C3H10T1/2 cells (A) and
MC3T3-E1 cells (B) were treated with 10 μg/mL of LRC extract for three days. The expression level of mRNA was calculated quantitatively by RT-PCR using targeted gene-specific primers and then normalized to Gapdh mRNA expression. Control: LRC non-treated cells. *: p < 0.05 vs.
Control.
4. LRC extract inhibited BMD loss in ovariectomized (OVX) mice.
Based on my in vitro results, I further investigated the effect of LRC extract in the osteoporosis model animals. It is known that OVX mice present reduced bone mass and quality. In addition, ovariectomy leads to an increase in body weight in mice with significant decreased right femur BMD and BMC. I first investigated the effects of LRC extract on OVX mice for 8 weeks. Thirty of the 8-week-old female ddY mice underwent either ovariectomy or sham surgery (Sham). The mice were then divided into five groups of six mice each: 1) Sham, 2) OVX control, 3) OVX administrated with 50 mg/kg/day of LRC extract, 4) OVX administrated with 150 mg/kg/day of LRC extract, and 5) OVX administrated with 300 mg/kg/day of LRC extract. The mice were housed for 8 weeks and their total body weight, total body fat (%fat), and right femur BMD and BMC were compared.
First, I compared the two non-LRC-administered mice groups: the Sham group and the OVX control group. The OVX group showed increased % body fat and decreased BMC and BMD compared to the Sham group, indicating that the OVX mice were an appropriate menopause-induced osteoporosis model (Figure 5). Next, I compared the OVX control and the LRC-administered groups. While body weight, body fat (% fat), and BMC were not significantly different among OVX control and LRC-administered groups, the BMD of the right femur bone was significantly increased in all of the LRC-administered groups compared to the OVX control group (Figure 5). The highest BMD was observed in the 150 mg/kg/day LRC extract-administered group.
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To investigate the effects of LRC extract in vivo for a longer period of time, six mice from the OVX control group and six mice from the LRC extract-administered group (150 mg/kg/day) were housed for 16 weeks. Total body weights were measured every week, and % body fat, BMC, and BMD of the right femur were measured at 0, 8 and 16 weeks, respectively. Similarly, there were no significant differences between the OVX control and the LRC-administered groups in terms of body weight, % fat, and BMC, although the LRC group showed a slight decrease in body weight and % fat, and a slight increase in BMC (Figure 6). As expected, BMD was significantly higher in the LRC-administered group than the OVX control group (Figure 6).
These results indicated that LRC extract inhibited BMD loss through ovariectomy in mice, and its effect continued for the duration of the test period (16 weeks). No toxic effects were observed in either mouse group.
Figure 5. Lycii Radicis Cortex (LRC) extract inhibited the loss of bone mineral density in ovariectomized (OVX) mice. The OVX mice were administered with different concentrations of LRC extract (50, 150, 300 mg/kg/day) for 8 weeks. OVX control: non-treated mice. Their total body weight (A), percentage of total body fat (% fat) (B), right femur bone mineral content (BMC) (C), and right femur bone mineral density (BMD) (D) were measured before and after treatment of LRC extract. *: p < 0.05 vs. OVX control.
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Figure 6. The long-term effects of Lycii Radicis Cortex (LRC) extract in ovariectomized (OVX) mice. The mice were administered with LRC extract (150 mg/kg/day) for 16 weeks.
OVX control: non-treated mice. Total body weights (A) were measured every week and percentage of total body fat (% fat) (B), right femur bone mineral content (BMC) (C), and right femur bone mineral density (BMD) (D) were measured at 0, 8 and 16 weeks. *: p <
0.05 vs. OVX control.
IV. DISCUSSION
Many studies have suggested pharmacological therapy (with several ingredients) for osteoporosis, but some medications have shown negative effects, including endometrial and breast cancers, when used long term (Bonura, 2009). Recently, natural herbal medicines have been used for the alternative long-term therapeutic treatment of several human diseases.
Screening of herbal extracts for increasing osteoblast differentiation may be an attractive approach to finding new therapeutic osteoporosis drugs that have fewer effects. It has been
Screening of herbal extracts for increasing osteoblast differentiation may be an attractive approach to finding new therapeutic osteoporosis drugs that have fewer effects. It has been