ER/estrogen receptor

Estrogen deficiency results in an imbalance between the levels of bone-resorping osteoclasts and bone-forming osteoblasts, eventually leading to overall bone loss. Dehydrodiconiferyl alcohol (DHCA), a lignan compound originally isolated from Cucurbita moschata, has been shown to bind to estrogen receptor, and indeed exhibits various activities of estrogen, such as anti-inflammatory and anti-oxidative stress effects. In this study, we tested whether synthetic DHCA could affect the BMP-2-induced osteoblastogenesis in vitro. In MC3T3-E1 cells, DHCA promoted BMP-2-induced differentiation of osteoblasts. Consistently, the expression of three osteoblastogenic genes known to be induced by BMP-2, ALP, osteocalcin and OPG, was up-regulated by DHCA treatment. DHCA was also shown to activate the production of RUNX2 by activating Smad1/5/9 and AMPK. Data from transient transfection assays suggested that DHCA might activate the estrogen receptor signaling pathway. Effects of DHCA on BMP-2-induced osteoblastogenesis were reduced when cells were treated with a specific siRNA to ERα or ERβ. Taken together, our results suggest that DHCA may be developed as an efficient therapeutic for osteoporosis by regulating osteoblastogenesis through its estrogenic effects. [1]

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DHCA promotes BMP-2-induced osteoblastogenesis with no cytotoxic effect [1]
MC3T3-E1 is a murine pre-osteoblast cell line that can differentiate into osteoblasts when stimulated with BMP-2. To test the effects of DHCA on osteoblastogenesis, MC3T3-E1 cells were treated with BMP-2 (25 ng/mL) and three different concentrations of DHCA (10, 20 and 40 μM) for 5 days followed by measuring the number of ALP-positive cells and the activity of ALP. As shown in Fig. 1A, the number of ALP-positive cells, as determined by ALP staining, was highly increased by treatment with DHCA in a dose-dependent manner. The effect of DHCA on actual ALP activity was also measured using cellular extracts and pNPP as a substrate. The level of ALP activity was enhanced as DHCA concentration increased (Fig. 1B).

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The effects of DHCA alone, (that is, in the absence of BMP-2), were also measured. MC3T3-E1 cells were treated with 10, 20, 40 and 80 μM of DHCA. Interestingly, neither the number of ALP-positive cells (Fig. 1C) nor the level of ALP activity (Fig. 1D) was changed, suggesting that DHCA works only when cells are differentiated by BMP-2. To be certain, the effects of DHCA on cell viability were measured. MC3T3-E1 cells were cultured with or without BMP-2 in the presence of DHCA followed by MTT assay. As shown in Fig. 1E, DHCA had little effect on cell viability throughout all concentrations used in this study, regardless of the presence of BMP-2 during the 72-h period. Taken together, these data indicated that DHCA might promote the BMP-2-induced osteoblastogenesis without cytotoxic effects.

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DHCA regulated the expression of genes associated with osteoblast differentiation [1]
It has previously been reported that stimulation of MC3T3-E1 cells by BMP-2 up-regulates the expression of ALP, osteocalcin and OPG, which all play important roles in the differentiation and function of osteoblasts. To study the effects of DHCA on the expression of these genes, MC3T3-E1 cells were treated with BMP-2 (25 ng/mL) and DHCA (10, 20 and 40 μM) for 24 h, and the RNA level was determined by quantitative RT-PCR. In all three cases, BMP-2 treatment increased their RNA levels by 2–3 fold. When cells were co-treated with 40 μM of DHCA, their levels were further enhanced by 2 fold in a dose-dependent manner (Fig. 2).

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DHCA promoted BMP-2-induced RUNX2 production via Smad and AMPK activation [1]
During BMP-2-induced osteoblastogenesis, RUNX2 becomes activated by the Smad signaling pathway. This is a critical step in the differentiation of osteoblasts, while AMPK has also been shown to play a positive role in this process. To test the effects of DHCA on the BMP-2-induced expression of RUNX2, MC3T3-E1 cells were co-treated with BMP-2 and three different concentrations of DHCA (10, 20 and 40 μM) for 24 h, and the protein level of RUNX2 was measured by Western blot. When cells were treated with BMP-2, the protein level of RUNX2 was increased (Fig. 3A, compare lanes 1 and 2), and co-treatment with DHCA further enhanced the protein level of RUNX2.
We also measured the effects of DHCA on other signaling proteins involved in the BMP-2-induced signaling pathway. MC3T3-E1 cells were co-treated with BMP-2 and DHCA for 30 min, and the phosphorylation status of Smad1/5/9 and AMPK were each determined by Western blot. When cells were treated with BMP-2, the level of phosphorylated Smad1/5/9 was highly increased (Fig. 3B, compare lanes 1 and 2), and co-treatment with DHCA further enhanced the amount of this phosphorylated protein (Fig. 3B, compare lanes 2 and 5). Similarly, phosphorylation of AMPK was also up-regulated by DHCA treatment (Fig. 3B). However, DHCA alone, namely in the absence of BMP-2, did not have any effect on the level of RUNX2 and phosphorylated Smad1/5/9 (Fig. 3C and D). These data indicated that DHCA could up-regulate the RUNX2-related signaling pathways, but only when cells were already in an activated status by BMP-2.

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Effects of DHCA on osteoblastogenesis were mediated by ERα and ERβ [1]
It is well known that estrogen promotes early osteoblast differentiation. To test the effects of DHCA on the estrogen-induced signaling pathway, MC3T3-E1 cells were transfected with a luciferase reporter plasmid containing the nucleotide sequences for estrogen responsive element (ERE). Twenty-four hours later, transfected cells were treated with estradiol or DHCA for 6 h. Total proteins were extracted, and the relative level of luciferase activity was measured. When cells were treated with DHCA, the level of luciferase activity was increased in a dose-dependent manner (Fig. 4A), by 2.6-fold at 40 μM, indicating that DHCA might interact with the estrogen receptor in MC3T3-E1 cells.
There are two different types of estrogen receptors, ERα and ERβ, and each has different functions due to its difference in their affinity for ligands. To investigate which of the two estrogen receptors interacts with DHCA to exert the observed effects, MC3T3-E1 cells were transfected with siRNA against ERα or ERβ followed by treatment with BMP-2 and DHCA. First, specificity of siRNA was measured. Cells were transfected with 30 pmole of siRNA for each receptor, and the protein level of ERα and ERβ was measured by Western blot. In both cases, the protein level was highly reduced (Fig. 4B). Next, the effect of siRNAs on the osteoblastogenesis was determined by measuring the number of ALP-positive cells and the level of ALP activity when cells were treated with BMP-2 (25 ng/mL) and DHCA (40 μM). As shown in Fig. 4C and D, both parameters were highly decreased when cells were transfected with siRNAs for ERα or ERβ. The effect of siRNAs was also measured on the BMP-2/DHCA-mediated activation of three osteoblastogenic genes (ALP, osteocalcin and OPG), and the RNA levels of all three genes were highly reduced (Fig. 4E–G). Taken together, these data indicated that DHCA might interact with both ERα and ERβ to promote BMP-2-induced osteoblast differentiation.

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The effect ofDHCA on LPS-decreased cell viability and LPS-induced NO and IL-1β production in RAW 264.7 cells [2]
Inflammation exerts a vital role in wound healing process. The invasion of pathogens causes a series of cellular immune responses, in which macrophages are critically involved. Macrophages could recognize pathogens through pattern recognition receptors, which produce various pro-inflammatory mediators to increase inflammatory cells infiltration in wound area. We first examined the effect of DHCA on the viability of RAW 246.7 cells by MTT assay. Results indicated treatment of RAW 246.7 cells with DHCA (0–200 µM) had no effect on cells viability compared with the control group (Figure 5a). DHCA inhibited LPS-induced expression of iNOS and NF-κB in RAW 264.7 cells [2]
To further delineate the anti-inflammatory mechanism of DHCA, the activities of NF-κB and iNOS in RAW 264.7 cells were analysed by the immunofluorescence assay. We found LPS stimulation increased nuclear translocation of NF-κB in RAW 264.7 cells, and the expression of iNOS was also upregulated by LPS administration. However, the treatment of 200 µM DHCA attenuated the activation of NF-κB and expression of iNOS (P < 0.05) induced by LPS (Figure 6). These data indicated DHCA might inhibit production of inflammatory mediator NO in LPS-induced RAW 246.7 cells by inhibiting the NF-κB/iNOS signalling pathway.

DHCA could promote scalp wound healing in mice by enhancing epithelial cell proliferation and collagen formation and reducing inflammatory cells infiltration. Moreover, the NF-κB nuclear translocation was suppressed remarkably by DHCA administration in connective tissue of healing area. DHCA was also shown to inhibit production of nitric oxide (NO) and interleukin (IL)-1β with downregulated inducible nitric oxide synthase (iNOS) expression in LPS-induced RAW 246.7 cells. More importantly, DHCA administration upregulated p-IκBα expression and induced nuclear translocation of NF-κB without affecting its expression. Conclusions: Our study indicated that DHCA exerted anti-inflammatory activity through inactivation of NF-κB pathways in macrophages and subsequently improved wound healing. [2]

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DHCA promoted scalp wound healing in mice [2]
The effect of DHCA on wound healing was studied using a mouse full-thickness scalp wound model. The wound areas began to shrink on Day 1. The contraction percentage of wound in both low dose and high dose groups was increased significantly compared with the control group 3 days after wounding (P < 0.05). More importantly, there was almost no unhealed wound area in the DHCA high dose group on Day 7, while the percentage of wound closure is only 69% in control group (Figure 2a, b). These results demonstrated that DHCA could promote scalp wound healing in mice.

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DHCA promoted epithelial cells proliferation and collagen formation and reduced inflammatory cells infiltration [2]
In order to further explore the mechanism of DHCA to promote skin wound healing, we stained the wound tissue sections with H&E and Masson’s trichrome staining. As shown in Figure 3a,b, the thickness of epithelium was significantly thicker in DHCA low dose and high dose groups compared with the control group (P < 0.05), which suggested that DHCA could promote the proliferation of epithelial cells in the wounds. Additionally, the number of inflammatory cells was reduced by DHCA administration, indicating that DHCA could inhibit infiltration of inflammatory cells in healing area (Figure 3a). Masson's trichrome staining illustrated that the collagen density of connective tissue in both low dose and high dose groups was increased significantly compared with the control group (P < 0.05), illuminating that DHCA could promote collagen formation in healing area (Figure 3c, d).

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DHCA inhibited NF-κB nuclear translocation in connective tissue of healing area [2]
Based on the results mentioned above, we speculated DHCA may promote scalp wound healing in mice by acting on the NF-κB signalling pathway. Therefore, we studied the effect of DHCA on the NF-κB nuclear translocation in wound tissue by the immunofluorescence assay. As shown in Figure 4, the NF-κB nuclear translocation was suppressed remarkably by DHCA administration in connective tissue (P < 0.05) rather than epithelium (P > 0.05) of healing area compared with the control group. These results elucidated that DHCA could restrain NF-κB nuclear translocation in cells in connective tissue to promote wound healing.

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DHCA decreased IL-1β level in mice serum [2]
Subsequently, we investigated the effect of DHCA on the level of pro-inflammatory cytokine IL-1β, the NF-κB downstream factor, in mice serum. As shown in Figure 4d, DHCA could significantly inhibit the production of IL-1β (P < 0.05), reconfirming that DHCA-promoted wound healing might be related to NF-κB signalling pathways.

Osteoblast differentiation in vitro [1]
For the osteoblast differentiation experiments, MC3T3-E1 cells were plated at 2 × 103 cells per well in 96-well culture plates containing α-MEM with 10% FBS. Twenty-four hours later, cells were treated with 25 ng/mL of BMP-2 and various concentrations of DHCA. After 5 days in culture, the cells were subjected to Leukocyte Alkaline Phosphatase (ALP) Kit according to the manufacturer's instructions.

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Measuring ALP activity [1]
After osteoblast differentiation, ALP activities of osteoblasts were measured in the well by incubation for 30 min at 37 °C with 100 μl of Alkaline Phosphatase Yellow (pNPP) Liquid Substrate System for ELISA containing 1% Tween-20. The reaction was terminated by the addition of 50 μl of NaOH (300 mM), and activities were measured at 405 nm.

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MTT assay [1]
MTT assay was performed as described previously. Briefly, MC3T3-E1 cells were treated with BMP-2 (25 ng/mL) or various concentrations of DHCA for 24–72 h. Cells were then incubated with an MTT labeling reagent for 4 h followed by the addition of solubilization solution. After 24 h, cytotoxicity was determined by measuring the OD at 550 nm using an ELISA microplate reader.

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Western blot analysis [1]
MC3T3-E1 cells were plated in 100 mm culture dishes. Twenty-four hours later, cells were treated with BMP-2 (25 ng/mL) and various concentrations of DHCA for 30 min. After treatment, cells were washed with cold PBS and lysed with phosphosafe extraction buffer. Total proteins were separated by SDS-PAGE and electrophoretically transferred to PVDF membranes.

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Luciferase reporter plasmid assay Inducible estrogen responsive element (ERE)-responsive luciferase reporter plasmid was purchased from QIAGEN. Luciferase reporter plasmid assay was performed as described previously. Briefly, MC3T3-E1 cells were transiently transfected with ERE-reporter plasmid and a β-galactosidase plasmid (1 μg), using lipofectamine 2000 according to the manufacturer's protocol. Twenty-four hours after transfection, the cells were treated with 17β-estradiol (10 nM) and various concentrations of DHCA for 24 h. Cell lysates were prepared, and a luciferase activity assay was performed using the Luciferase Reporter kit according to the manufacturer's protocol with a microplate luminometer. Luciferase activity was normalized to β-gal activity.

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MTT assay [2]
RAW 264.7 cells were seeded into 96-well plates at a density of 1 × 104 cells/well and then incubated with different concentrations of DHCA (0, 3.125, 6.25, 12.5, 25, 50, 100 and 200 μmol/l) and LPS (0, 0.5, 1, 2, 4, 8, 16 and 32 μg/ml). After 24 h, 10 μl of MTT (5 mg/ml) was added to each well containing 100 μl culture medium and incubated for another 3 h, and then, the supernatant was discarded. Subsequently, 150 μl of DMSO was added to each well to dissolve the formazan crystal. Finally, the absorbance was determined by Thermo Varioskan Flash Multi-Mode Microplate Reader at 492 nm, and the viability of cells in each group was calculated. All of the experiments were performed in triplicate.

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Measurement of NO and IL-1β production [2]
RAW 264.7 cells were incubated in 96-well plates and pretreated with different concentrations of DHCA (100 and 200 μmol/l) for 4 h, and then, cells were treated with 1 μg/ml of LPS for 24 h. Nitrite level was determined by adding Griess reagent (100 μl) to the culture supernatant (50 μl). And the absorbance was measured at 540 nm with Thermo Varioskan Flash Multi-Mode Microplate Reader according to the literature. Moreover, IL-1β level in the culture supernatant of cells with different treatment was determined using a commercial ELISA kit. All of the experiments were performed in triplicate.

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Immunofluorescence assay [2]
RAW 264.7 cells were seeded into 6-well plates and pretreated with 50, 100 and 200 μmol/l of DHCA for 4 h. After stimulated by 1 μg/ml of LPS for another 24 h, the cells were washed three times with PBS, followed by fixation in 4% paraformaldehyde for 20 min. Then, the cells were permeabilized in 0.15% Triton X-100 for 10 min and blocked by 10% fetal bovine serum for 30 min. After culturing with antibodies targeting iNOS or NF-κB overnight, DAPI was used for the nucleus counterstain. Micrographs were captured using the fluorescence microscope. And image analysis was performed using NIS-Elements AR image analysis software.

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Western blot analysis [2]
RAW 264.7 cells were seeded into 6-well plates, pretreated with 50, 100 and 200 μmol/l of DHCA for 4 h, and stimulated by LPS (1 μg/ml). After 24 h, the cells were collected and suspended in the lysis buffer containing protease inhibitors. Lysates were obtained by centrifugation at 12 000g for 10 min. Total protein concentration was measured with BCA protein content assay kit, and 30 μg proteins were separated by 10% SDS-polyacrylamide gel electrophoresis (SDS-PAGE). After separation, proteins were electrophoretically transferred to a nitrocellulose membrane. Then, the membrane was incubated at 4 °C overnight with tris-glycine protein electrophoresis buffer containing the antibodies against β-actin (1 : 1000 dilution), NF-κB (1 : 1000 dilution), IκBα (1 : 1000 dilution) or p-IκBα (1 : 10 000 dilution). Subsequently, membranes were incubated with HRP-labelled secondary antibody for 1 h at room temperature. Finally, the membranes were washed three times with TBST and visualized by an enhanced chemiluminescence with ECL kit.

Establishment of a wound model [2]
All mice were housed under ambient conditions (12-h light/dark cycle at 25 °C) with water and standard mice chow ad libitum and were randomly assigned to four groups including control group, vehicle group and low dose and high dose groups. Subsequently, all mice were anaesthetized with 1% pentobarbital sodium, and fur of the back and scalp were removed with electric shaver. After shaving and sterilization, two full-thickness dermal excisional wounds were made on the mice scalp with a 2-mm sterile biopsy punch. Then, 30 μl of DHCA solution prepared with acetonitrile and olive oil (v/v 4 : 1) was dripped to each wound at a low dose group of 5 mg/ml and a high dose group of 20 mg/ml daily for 9 days. Mice in vehicle control group received vehicle alone, while no treatments were given to the wounds of mice in control group. Wound healing was macroscopically monitored by taking digital photographs at the first seven days. The wound sizes were measured with Image J software, and the percentage of wound contraction was calculated using the formula: wound contraction% = (wound area day 0 − wound area day n/wound area day 0) × 100. Nine days following injury, mice were anesthetized and serum was obtained for analyzing IL-1β level according to the manufacture’s protocol. Wound tissues of the scalp were harvested using 3-mm biopsy punches and were processed by paraffin embedding and fixation.

[1]. Dehydrodiconiferyl alcohol promotes BMP-2-induced osteoblastogenesis through its agonistic effects on estrogen receptor. Biochem Biophys Res Commun. 2018 Jan 15;495(3):2242-2248.

[2]. Dehydrodiconiferyl alcohol from Silybum marianum (L.) Gaertn accelerates wound healing via inactivating NF-κB pathways in macrophages. J Pharm Pharmacol. 2020 Feb;72(2):305-317.

Dehydrodiconiferyl alcohol is a guaiacyl lignin obtained by cyclodimerisation of coniferol. It has a role as a plant metabolite and an anti-inflammatory agent. It is a member of 1-benzofurans, a primary alcohol, a guaiacyl lignin and a member of guaiacols. It is functionally related to a coniferol.
Dehydrodiconiferyl alcohol has been reported in Codonopsis pilosula, Urtica dioica, and other organisms with data available.

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Our data indicate that DHCA promotes BMP-2-induced osteoblast differentiation by interacting with either ERα or ERβ as an agonist for both receptors. We have recently shown that DHCA inhibits RANKL-induced osteoclast differentiation in vitro and ovariectomy-induced bone loss in vivo (submitted for publication). Taken together, our results indicate that DHCA may be developed as an efficient therapeutic for osteoporosis by controlling the osteoclast/osteoblast ratio through its estrogenic effects.[1]

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In summary, we reported firstly that DHCA could promote wound healing by accelerating proliferation of epithelial cells, increasing collagen formation and inhibiting infiltration of inflammatory cells in a full-thickness scalp wound healing model in mice. And the inactivation of NF-κB pathways was involved in the anti-inflammatory effects of DHCA in macrophages. These findings brought to light preliminarily the promising therapeutic potential of DHCA in skin wound healing. [2]

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Objectives: The aim of this study was to investigate the molecular mechanisms of the efficacy of lignin compound dehydrodiconiferyl alcohol (DHCA) isolated from Silybum marianum (L.) Gaertn in improving wound healing. These findings preliminarily brought to light the promising therapeutic potential of DHCA in skin wound healing. Methods: First, the effect of DHCA on healing in vivo was studied using a full-thickness scalp wound model of mice by topical administration. Histopathological examinations were then conducted by haematoxylin and eosin (H&E), Masson's trichrome staining and the immunofluorescence assay. Second, we further examined the anti-inflammatory mechanism of DHCA in lipopolysaccharide (LPS)-induced RAW 264.7 macrophages by immunofluorescence assay and Western blot analysis. [2]

C20H22O6

358.3851

358.142

4263-87-0

(7R,8S)-Dehydrodiconiferyl alcohol;155836-29-6;(E)-Dehydrodiconiferyl alcohol;528814-97-3

5372367

Typically exists as solid at room temperature

1.292g/cm3

562ºC at 760mmHg

161-162℃

293.7ºC

1.63

2.624

3

6

6

26

469

0

O1C2C(=C([H])C(/C(/[H])=C(\[H])/C([H])([H])O[H])=C([H])C=2[C@]([H])(C([H])([H])O[H])[C@@]1([H])C1C([H])=C([H])C(=C(C=1[H])OC([H])([H])[H])O[H])OC([H])([H])[H]

KUSXBOZNRPQEON-ONEGZZNKSA-N

InChI=1S/C20H22O6/c1-24-17-10-13(5-6-16(17)23)19-15(11-22)14-8-12(4-3-7-21)9-18(25-2)20(14)26-19/h3-6,8-10,15,19,21-23H,7,11H2,1-2H3/b4-3+

4-[3-(hydroxymethyl)-5-[(E)-3-hydroxyprop-1-enyl]-7-methoxy-2,3-dihydro-1-benzofuran-2-yl]-2-methoxyphenol

Dehydrodiconiferyl alcohol; 4263-87-0; dehydrodiconiferol; 4-[3-(hydroxymethyl)-5-[(E)-3-hydroxyprop-1-enyl]-7-methoxy-2,3-dihydro-1-benzofuran-2-yl]-2-methoxyphenol; lignin cw compound-2004; Coniferyl alcohol, dehydrodi-; Diconiferyl alcohol, dehydro-; CHEBI:91184;

2934.99.9001

Powder      -20°C    3 years

                     4°C     2 years

In solvent   -80°C    6 months

                  -20°C    1 month

Room temperature (This product is stable at ambient temperature for a few days during ordinary shipping and time spent in Customs)

May dissolve in DMSO (in most cases), if not, try other solvents such as H2O, Ethanol, or DMF with a minute amount of products to avoid loss of samples

Note: Listed below are some common formulations that may be used to formulate products with low water solubility (e.g. < 1 mg/mL), you may test these formulations using a minute amount of products to avoid loss of samples.

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Injection Formulation 1: DMSO : Tween 80： Saline = 10 : 5 : 85 (i.e. 100 μL DMSO stock solution → 50 μL Tween 80 → 850 μL Saline)
*Preparation of saline: Dissolve 0.9 g of sodium chloride in 100 mL ddH ₂ O to obtain a clear solution.
Injection Formulation 2: DMSO : PEG300 ：Tween 80 : Saline = 10 : 40 : 5 : 45 (i.e. 100 μL DMSO → 400 μLPEG300 → 50 μL Tween 80 → 450 μL Saline)
Injection Formulation 3: DMSO : Corn oil = 10 : 90 (i.e. 100 μL DMSO → 900 μL Corn oil)
Example: Take the Injection Formulation 3 (DMSO : Corn oil = 10 : 90) as an example, if 1 mL of 2.5 mg/mL working solution is to be prepared, you can take 100 μL 25 mg/mL DMSO stock solution and add to 900 μL corn oil, mix well to obtain a clear or suspension solution (2.5 mg/mL, ready for use in animals).

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Injection Formulation 4: DMSO : 20% SBE-β-CD in saline = 10 : 90 [i.e. 100 μL DMSO → 900 μL (20% SBE-β-CD in saline)]
*Preparation of 20% SBE-β-CD in Saline (4°C，1 week): Dissolve 2 g SBE-β-CD in 10 mL saline to obtain a clear solution.
Injection Formulation 5: 2-Hydroxypropyl-β-cyclodextrin : Saline = 50 : 50 (i.e. 500 μL 2-Hydroxypropyl-β-cyclodextrin → 500 μL Saline)
Injection Formulation 6: DMSO : PEG300 : castor oil : Saline = 5 : 10 : 20 : 65 (i.e. 50 μL DMSO → 100 μLPEG300 → 200 μL castor oil → 650 μL Saline)
Injection Formulation 7: Ethanol : Cremophor : Saline = 10： 10 : 80 (i.e. 100 μL Ethanol → 100 μL Cremophor → 800 μL Saline)
Injection Formulation 8: Dissolve in Cremophor/Ethanol (50 : 50), then diluted by Saline
Injection Formulation 9: EtOH : Corn oil = 10 : 90 (i.e. 100 μL EtOH → 900 μL Corn oil)
Injection Formulation 10: EtOH : PEG300：Tween 80 : Saline = 10 : 40 : 5 : 45 (i.e. 100 μL EtOH → 400 μLPEG300 → 50 μL Tween 80 → 450 μL Saline)

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Oral Formulation 1: Suspend in 0.5% CMC Na (carboxymethylcellulose sodium)
Oral Formulation 2: Suspend in 0.5% Carboxymethyl cellulose
Example: Take the Oral Formulation 1 (Suspend in 0.5% CMC Na) as an example, if 100 mL of 2.5 mg/mL working solution is to be prepared, you can first prepare 0.5% CMC Na solution by measuring 0.5 g CMC Na and dissolve it in 100 mL ddH2O to obtain a clear solution; then add 250 mg of the product to 100 mL 0.5% CMC Na solution, to make the suspension solution (2.5 mg/mL, ready for use in animals).

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Oral Formulation 3: Dissolved in PEG400
Oral Formulation 4: Suspend in 0.2% Carboxymethyl cellulose
Oral Formulation 5: Dissolve in 0.25% Tween 80 and 0.5% Carboxymethyl cellulose
Oral Formulation 6: Mixing with food powders

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Note: Please be aware that the above formulations are for reference only. InvivoChem strongly recommends customers to read literature methods/protocols carefully before determining which formulation you should use for in vivo studies, as different compounds have different solubility properties and have to be formulated differently.

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 (Please use freshly prepared in vivo formulations for optimal results.)