Skip to main content
NCBI home page
Toxicological Sciences logoLink to Toxicological Sciences
. 2021 Apr 15;182(1):96–106. doi: 10.1093/toxsci/kfab044

Mechanistic Evaluation of Black Cohosh Extract-Induced Genotoxicity in Human Cells

Ji-Eun Seo 1, Xiaoqing Guo 1, Dayton M Petibone 1, Sharon D Shelton 1, Ying Chen 1, Xilin Li 1, Volodymyr Tryndyak 2, Stephanie L Smith-Roe 3, Kristine L Witt 3, Nan Mei 1, Mugimane G Manjanatha 1,✉
PMCID: PMC8285013  PMID: 33856461

Abstract

Black cohosh extract (BCE) is marketed to women as an alternative to hormone replacement therapy for alleviating menopausal symptoms. Previous studies by the National Toxicology Program revealed that BCE induced micronuclei (MN) and a nonregenerative macrocytic anemia in rats and mice, likely caused by disruption of the folate metabolism pathway. Additional work using TK6 cells showed that BCE induced aneugenicity by destabilizing microtubules. In the present study, BCE-induced MN were confirmed in TK6 and HepG2 cells. We then evaluated BCE-induced DNA damage using the comet assay at multiple time points (0.5–24 h). Following a 0.5-h exposure, BCE induced significant, concentration-dependent increases in %tail DNA in TK6 cells only. Although DNA damage decreased in TK6 cells over time, likely due to repair, small but statistically significant levels of DNA damage were observed after 2 and 4 h exposures to 250 µg/ml BCE. A G1/S arrest in TK6 cells exposed to 125 µg/ml BCE (24 h) was accompanied by apoptosis and increased expression of γH2A.X, p-Chk1, p-Chk2, p53, and p21. Conditioning TK6 cells to physiological levels of folic acid (120 nM) did not increase the sensitivity of cells to BCE-induced DNA damage. BCE did not alter global DNA methylation in TK6 and HepG2 cells cultured in standard medium. Our results suggest that BCE induces acute DNA strand breaks which are quickly repaired in TK6 cells, whereas DNA damage seen at 4 and 24 h may reflect apoptosis. The present study supports that BCE is genotoxic mainly by inducing MN with an aneugenic mode of action.

Keywords: botanical extract, TK6 cells, in vitro genotoxicity, micronucleus assay, comet assay, DNA damage response

Black cohosh (Actaea racemosa, previously known as Cimicifuga racemosa) is a perennial native to North American. Black cohosh extract (BCE) prepared from roots and rhizomes is among the top 10 best-selling botanical dietary supplements in the United States (Smith et al., 2019). It is marketed to women for relief of symptoms associated with gynecological ailments, including menopausal symptoms (e.g., hot flashes), menstrual cramps, and premenstrual syndrome. Early studies suggested that BCE works via an estrogenic mechanism of action (MOA), but conflicting results, that is, estrogenic, antiestrogenic, or neither estrogenic nor antiestrogenic effects, have been reported in both in vivo and in vitro studies (Mercado-Feliciano et al., 2012; Ruhlen et al., 2007, 2008). Several high-quality randomized controlled trials also demonstrated no significant benefit of BCE over placebo in treating menopausal symptoms (Leach and Moore, 2012; Pockaj et al., 2006; Reed et al., 2008; Shulman et al., 2011; van der Sluijs et al., 2009). Meanwhile, cases of BCE-associated liver toxicity have been reported, including acute and chronic hepatitis, fulminant hepatic failure, cirrhosis, and liver necrosis (Chow et al., 2008; Guzman et al., 2009; Lynch et al., 2006; Muqeet Adnan et al., 2014; Pierard et al., 2009). Thus, the efficacy of BCE in alleviating menopausal symptoms is questionable but the wide consumption of BCE raises safety concerns regarding potential hepatotoxicity and other adverse health effects.

BCE was nominated to the National Toxicology Program (NTP) for general toxicity testing, and reproductive and developmental toxicity and carcinogenicity testing. Results of initial 90-day toxicity studies found that oral administration of BCE to female B6C3F1/N mice and Wistar-Han rats resulted in increased frequencies of micronucleated red blood cells and hematological changes consistent with megaloblastic anemia (Mercado-Feliciano et al., 2012). In a follow-up study, BCE induced hematological and biochemical changes consistent with a cobalamin (vitamin B12), and possibly folate, deficiency in female B6C3F1/N mice (Cora et al., 2017). Deficiencies in cobalamin or folate can induce micronucleus (MN) formation and megaloblastic anemia (Everson et al., 1988; Fenech, 1999; MacGregor et al., 1997; Wickramasinghe, 2006), suggesting that BCE-induced MN and hematological changes in rodents may have resulted from disruption of the 1-carbon folate metabolism pathway. Recently, the effects of folate deficiency on MN induction were evaluated by the NTP using human lymphoblastoid TK6 cells (Smith-Roe et al., 2018). The results showed that although BCE significantly increased MN frequencies in TK6 cells cultured in both standard medium (supraphysiological levels of folic acid [FA], 3000 nM) and medium with more physiologically relevant concentrations of FA (120 nM), the response observed in the low FA medium (120 nM) was significantly higher than the response in high FA medium (3000 nM). A subsequent study demonstrated that BCE destabilized tubulin polymerization in TK6 cells, suggesting an aneugenic MOA for the MN induction (Bernacki et al., 2019).

Genotoxic xenobiotics may act through a variety of mechanisms, inducing single- and double-strand DNA breaks, chromosomal damage, and gene mutations. BCE treatment of TK6 cells has demonstrated the ability to induce MN formation in medium containing physiologically relevant levels of FA (Smith-Roe et al., 2018), suggesting that BCE might disrupt folate metabolism (Cora et al., 2017). The disruption of folate metabolism results in reduced levels of available methyl groups required for conversion of deoxyuridine monophosphate to deoxythymidine monophosphate. This can lead to excess accumulation and incorporation of uracil into DNA in place of thymine, leading to double-strand DNA breaks (Blount et al., 1997; Everson et al., 1988; MacGregor et al., 1997). Given these observations, we sought to determine using the alkaline comet assay whether BCE induces DNA damage and if so, whether folate metabolism is associated with this effect.

The alkaline comet assay is a sensitive method for detecting DNA damage, including DNA strand breaks, alkali-labile sites, DNA-DNA/DNA-protein cross-linking, and single-strand breaks associated with incomplete excision repair sites, at the single-cell level (Azqueta and Collins, 2013; Tice et al., 2000). In the present study, we evaluated BCE-induced DNA damage in human p53-competent TK6 and HepG2 cells at various time points. TK6 cells cultured in standard medium (supraphysiological levels of FA) and medium modified to contain levels of FA closer to physiological levels in humans were used in order to compare results with the previous NTP in vitro MN study (Smith-Roe et al., 2018). BCE is a complex mixture containing more than 100 chemical constituents (Wang et al., 2011). Because the chemical composition of BCE can vary significantly due to plant species, growth conditions, harvesting, and manufacturing processes (Jiang et al., 2006), we tested the same sample of BCE used in previous NTP toxicity studies (Bernacki et al., 2019; Cora et al., 2017; Mercado-Feliciano et al., 2012; Smith-Roe et al., 2018). In addition, mechanisms underlying BCE-induced cytotoxicity and genotoxicity were explored by measuring activation of DNA damage checkpoint signaling cascades, apoptosis, and cell cycle arrest, along with DNA methylation modifications.

MATERIALS AND METHODS

 

 

Cell culture conditions

Human lymphoblastoid TK6 cells and human hepatoblastoma HepG2 cells were obtained from the American Type Culture Collection (ATCC) (Manassas, Virginia). TK6 cells were grown in RPMI-1640 medium containing l-glutamine (Gibco, Gaithersburg, Maryland) and 2268 nM FA, supplemented with 10% fetal bovine serum (FBS) (Atlanta Biologicals, Lawrenceville, Georgia), 50 U/ml penicillin (Gibco), and 50 µg/ml streptomycin (Gibco). TK6 cells were also cultured in a specialized medium containing physiological levels (120 nM) of FA by adding FA (dissolved in a water-based 1 M sodium bicarbonate solution) to FA-free RPMI-1640 medium (Gibco), supplemented with 10% dialyzed FBS (Sigma-Aldrich), 1% pluronic F-68 (Gibco), 1 mM sodium pyruvate (Gibco), 50 U/ml penicillin, and 50 µg/ml streptomycin. The cell culture density was maintained at less than 1.5 × 106 cells/ml. Prior to BCE exposure, TK6 cells were cultured in either the standard culture medium (2268 nM FA) or 120 nM FA medium for 5 days in 6-well plates, with cells subcultured at a seeding concentration of 2 × 105 cells/ml every other day. HepG2 cells were grown in Dulbecco’s modified Eagle medium (DMEM) containing d-glucose and l-glutamine (Gibco), supplemented with 10% FBS and 1% antibiotic-antimycotic (Gibco) agents. All cell cultures were maintained at 37°C in a humidified atmosphere with 5% CO2.

Black cohosh extract treatment

BCE powder (NTP BCE, lot no. 3012782; PlusPharma, Inc., Vista, California) was stored at −20°C and protected from light prior to use. The stock solution (100 mg/ml) was prepared by dissolving the BCE powder in DMSO (Sigma-Aldrich), sonicating for 1 min in an ultrasonic water bath, and mixing overnight at 37°C on a shaker. BCE stock solution was prepared fresh for each experiment. Working solutions were diluted from the stock solution to give final BCE concentrations of 25‒250 μg/ml. DMSO was used as a vehicle control and never exceeded 0.5% (v/v). For the treatment, TK6 cells in the standard culture medium or 120 nM FA medium were adjusted to 6 × 105 cells/well (2 ml) in 24-well plates. HepG2 cells were seeded at a density of 2 × 104 cells/well (100 μl) in 96-well plates and cultured overnight. The cells were exposed to various concentrations of BCE or vehicle (DMSO) for 0.5, 1, 2, 4, or 24 h.

Micronucleus assay

The MN assay was performed using the In Vitro MicroFlow kit (Litron Laboratories, Rochester, New York). Following a 24-h exposure, TK6 cells were harvested and HepG2 cells were cultured for additional 16 h in fresh medium to achieve a 1.5- to 2-fold cell population increase. After harvesting, cells were stained with a nucleic acid dye, ethidium monoazide (EMA), which can cross the compromised cell membranes of apoptotic and necrotic cells and stain DNA. Next, the cells were lysed, DNA was stained with SYTOX Green, and the MN analysis was performed using a FACSCanto II flow cytometer equipped with a High Throughput Sampler (BD Biosciences, San Jose, California). The stopping gate was set at 10,000 intact nuclei per well and threshold parameters were set as recommended in the instruction manual. The MN frequency (%MN) was calculated as the ratio of MN events to the total number of nucleated events. Cytotoxicity was evaluated using relative survival (% of control) by comparing treated cells to the vehicle control using nucleated event count at a specified time point (Guo et al., 2020). The top concentration was selected to achieve a cytotoxicity of 55 ± 5% as recommended by the Organization for Economic Co‐operation and Development (OECD) Test Guideline 487 for the in vitro MN assay (OECD, 2016). The percent of apoptotic/necrotic cells (EMA-positive events) was also obtained from the MN assay. Mitomycin C (50 ng/ml for 24 h) (Sigma-Aldrich) dissolved in water was used as the positive control.

Cytotoxicity assays

Cytotoxicity was determined by the CellTiter-Glo luminescent cell viability assay (ATP) (Promega, Madison, Wisconsin) and the CellTiter-Blue cell viability assay (Promega), as described previously (Guo et al., 2018). For TK6 cells, 10 μl of the CellTiter-Glo or CellTiter-Blue reagents were added into each well of a 96-well flat bottom plate (white or black, respectively), and then 90 μl of treated cells were transferred into each well. For HepG2 cells, after the removal of treatment media, CellTiter-Glo, or CellTiter-Blue reagents were added directly into each well at a ratio of 1:10 in phosphate-buffered saline (PBS, Gibco). Luminescence for the CellTiter-Glo luminescent cell viability assay or fluorescence at 530 nm excitation and 590 nm emission for the CellTiter-Blue cell viability assay was recorded with a multimode microplate reader (Cytation 5 with Gen5 software, BioTek, Winooski, Vermont). The relative cytotoxicity (%) was calculated by comparing the intensity level of the luminescence or fluorescence signals in BCE-treated cells to those of the vehicle control.

Alkaline comet assay

DNA damage was measured in the human cells using the standard comet assay (pH > 13) for BCE treatment at various time points or the 96-well CometChip system for BCE treatment in low FA medium. The standard comet assay was performed according to the manufacturer’s instructions (Trevigen, Gaithersburg, Maryland). Briefly, treated cells were washed with cold PBS (4°C) and combined with 1% agarose (Thermo Fisher Scientific, Waltham, Massachusetts) in PBS (>1.0 × 105 cells/ml) at 37°C. A 50-μl aliquot was pipetted immediately onto the Comet Slides (Trevigen) and the slides were placed flat at 4°C for 10 min in the dark. The slides were then transferred into a pre-chilled CometAssay lysis solution (Trevigen) for 1 h or overnight in the dark, followed by immersion in freshly prepared alkaline solution (pH > 13) for 1 h at 4°C. After electrophoresis at 21 V for 30 min, the slides were stained with SYBR Gold (Invitrogen, Carlsbad, California) and the comet images were acquired using a Leica DMI4000 B fluorescence microscope (Leica Microsystems Inc. Buffalo Grove, Illinois). The CometChip assay (Trevigen) was performed using the method described in our previous study (Seo et al., 2019). Briefly, treated cells (approximately 1.0 × 105 cells/ml) were transferred into each well of a 96-well CometChip which contains approximately 400 microwells per well, each with a diameter of 30 microns. The cells were gravity loaded into the microwells at 37°C for 30 min in a humidified atmosphere with 5% CO2. CometChip was gently rinsed with PBS to wash off excess cells and overlaid with 1% low melting point agarose (Thermo Fisher Scientific) in PBS. The CometChip was immersed in lysis solution at 4°C for 1 h, followed by freshly prepared alkaline solution (pH > 13) for 40 min to unwind and denature the DNA. Comets were developed by electrophoresis at 22 V for 50 min at 4°C. Subsequent staining with SYBR Gold allowed visualization of the comet tail. All comet images were analyzed by Trevigen Comet Analysis Software for calculating the percentage of tail DNA. Methyl methanesulphonate (MMS, 100 μM for a 4-h exposure) (Sigma-Aldrich) dissolved in DMSO was used as the positive control.

Western blotting

Following exposures, a total of 1 × 107 TK6 cells were harvested and lysed in 150 μl radioimmunoprecipitation (RIPA) lysis buffer supplemented with Halt Protease Inhibitor Cocktail (Thermo Fisher Scientific) for protein extraction. The protein concentrations of cell lysates were determined by the Pierce BCA protein assay kit (Thermo Fisher Scientific) (Lin et al., 2014). The protein samples were dissolved in Laemmli sample buffer (Bio-Rad) with 2.8% β-mercaptoethanol (Sigma-Aldrich) and boiled at 95°C for 5 min. Equivalent amounts (20 μg) of protein lysates were loaded and separated using 4%–15% Mini-PROTEAN TGX precast gradient gels (10-well; Bio-Rad), and then transferred onto PVDF membranes (Millipore Corporation, Billerica, Massachusetts). The following primary antibodies were used: phospho-histone H2A.X (γH2A.X; Ser-139), histone H2A.X (H2A.X), cleaved caspase 3 (c-caspase 3), poly (ADP-ribose) polymerase (PARP), phospho-checkpoint kinase1 (p-Chk1; Ser-345), phospho-checkpoint kinase 2 (p-Chk2; Thr-68), p21, p53 (Cell Signaling Technology, Danvers, Massachusetts), and GAPDH (internal control; Santa Cruz Biotechnology, Dallas, Texas). The membranes were then incubated with the secondary antibody conjugated with horseradish peroxidase (HRP) (Santa Cruz Biotechnology, Dallas, Texas) at a dilution of 1:10,000. The protein band signals were determined by FluorChem E System and quantified by AlphaView SA (ProteinSimple, San Jose, California).

Terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling (TUNEL) assay

DNA fragmentation in apoptosis was detected using the TUNEL Assay Kit—BrdU-Red (Abcam) following the manufacturer’s instructions. Briefly, the treated cells were fixed with 4% paraformaldehyde solution in PBS (Santa Cruz Biotechnology, Santa Cruz, California) for 15 min and washed with PBS. The fixed cells were incubated in 70% ethanol at –20°C for 30 min. After washing, the cells were sequentially incubated in DNA labeling solution containing Br-dUTP at 37°C for 60 min, anti-BrdU-Red antibody solution at 25°C for 30 min in the dark, and 7-amino actinomycin D (7-AAD)/RNase A solution for 30 min in the dark. Apoptotic cells were detected and analyzed using a FACSAria IIIu flow cytometer (BD Biosciences) following the manufacturer’s instructions.

Caspase 3/7 activity

The caspase 3/7 enzyme activity was measured using the Caspase-Glo 3/7 assay kit (Promega) according to the manufacturer’s instructions (Guo et al., 2015). Following a 4-h or 24-h treatment with BCE, the treatment medium was removed by centrifugation and cells were resuspended in PBS at a concentration of 1 × 105 cells/ml. Approximately 10,000 cells (100 μl of the cell suspension) were dispensed into each well of a 96-well white flat-bottomed plate and 100 μl/well of reaction reagents were added. The plates were incubated at 25°C for 30 min and luminescence was read with a Cytation 5 microplate reader (BioTek).

Cell cycle analysis

Cell cycle stages were determined using histograms of the nuclear DNA content in the MicroFlow MN assay. DNA content histograms were deconvoluted into G1, S, G2/M, and subG1 populations; the proportion of cells within each stage of the cell cycle was approximated using Flow Jo Software (Tree Star, Inc., Ashland, Oregon).

Methylated DNA immunoprecipitation quantitative PCR analysis of long interspersed nucleotide elements 1

Following a 72-h treatment, TK6 and HepG2 cells treated with BCE at concentrations causing 20% or less cytotoxicity were harvested for performing the Methylated DNA immunoprecipitation (MeDIP) assay to investigate alterations in DNA methylation. Genomic DNA was isolated from these cells using the DNeasy Blood and Tissue kit (Qiagen, Valencia, California). MeDIP was performed using the MethylMiner Methylated DNA Enrichment Kit (Invitrogen). Briefly, 1 µg of genomic DNA was randomly sheared by sonication to obtain fragments with an average range of 0.2–1.0 kb and 90% of the sheared DNA was incubated overnight at 4°C with MBD-Biotin protein coupled to M-280 Streptavidin Dynabeads; the remaining sheared DNA was used as an input. The captured methylated DNA was eluted as a single fraction using a High-Salt Elution Buffer containing 2000 mM NaCl and purified by ethanol precipitation. The methylation status of long interspersed nucleotide elements 1 (LINE-1) DNA repetitive elements was determined by real-time quantitative PCR (qPCR) of immunoprecipitated and input DNA fractions using primer sets shown in Supplementary Table 1. The results were normalized to the amount of input DNA.

Statistical analysis

Data were presented as the mean ± standard deviation (SD) from at least 3 independent experiments. Statistical significance was determined by 1-way analysis of variance followed by Dunnett’s test for pairwise comparisons using SigmaPlot 13.0 (Systat Software, San Jose, California). The significance level was set at p < .05.

RESULTS

 

BCE-Induced MN in TK6 and HepG2 Cells

Concentration-dependent increases in %MN were observed in TK6 cells cultured in the standard medium after 24 h exposure to BCE (Figure 1A). Significant increases in %MN were observed at 75, 100, and 125 µg/ml. BCE induced a 25-fold increase in %MN at the top concentration (125 µg/ml) compared with the vehicle control (10.1 ± 1.9 vs 0.4 ± 0.2), with a 30% reduction in relative survival. In contrast to the concentration-dependent increases in MN formation in TK6 cells, a marginally positive response was observed in HepG2 cells exposed to 200 µg/ml BCE only (Figure 1B), resulting in a 2-fold increase in MN frequencies compared with the vehicle control, with a cytotoxicity of 55%.

Figure 1.

Figure 1.

Open in a new tab

BCE-induced micronuclei formation in TK6 and HepG2 cells. TK6 (A) and HepG2 (B) cells were exposed to various concentrations of BCE for 24 h. Micronuclei formation was detected by the in vitro micronucleus (MN) assay. MN frequency (%MN) is presented as the percentage of micronuclei relative to intact nuclei (left y-axis and black bar). The cytotoxicity is presented as the percentage of relative survival (right y-axis and red line) from treated cells compared with the vehicle control. PC, positive control (50 ng/ml mitomycin C). *p < .05, **p < .01, and ***p < .001, compared with the vehicle control.

Evaluation of BCE-Induced Cytotoxicity and DNA Damage in TK6 and HepG2 Cells

In the standard medium, BCE at concentrations up to 250 µg/ml in TK6 cells (Figure 2A) and up to 500 µg/ml in HepG2 cells (Figure 2B) showed little or no cytotoxicity at 0.5–4 h time points when evaluated by the ATP and CellTilter-Blue assays. However, following a 24-h treatment, significant cytotoxicity was observed in TK6 cells treated with 75 µg/ml (18%), 125 µg/ml (37%‒39%) or 250 µg/ml (77%‒82%) BCE and in HepG2 cells treated with 250 µg/ml (12%‒24%) or 500 µg/ml (46%‒77%) BCE.

Figure 2.

Figure 2.

Open in a new tab

Evaluation of BCE-induced cytotoxicity in TK6 and HepG2 cells. TK6 (A) and HepG2 cells (B) were exposed to 25–250 µg/ml and 50–500 µg/ml BCE, respectively and the relative cytotoxicity (% of control) was measured using the CellTiter-Glo luminescent cell viability (ATP) and CellTiter-Blue cell viability assays at 0.5, 1, 2, 4, and 24 h time points. The data are expressed as the mean ± SD from at least 3 independent experiments. *p < .05, **p < .01, and ***p < .001, compared with the vehicle control.

The DNA damage response (DDR) to BCE exposure was assessed by the alkaline comet assay at various time points (0.5–24 h). A cutoff of <30% cytotoxicity was used to exclude false positive responses. Accordingly, TK6 cells treated with 250 µg/ml BCE and HepG2 cells treated with 500 µg/ml BCE for 24 h were not used for the comet assay due to severe cytotoxicity. Following a 0.5-h exposure, the % tail DNA was significantly increased in a concentration-dependent manner in TK6 cells (Figure 3A); at the top concentration of 250 µg/ml, BCE induced a 14.6-fold increase in DNA damage compared with the vehicle control (p < .001). Subsequently, decreasing levels of DNA damage were observed in TK6 cells with increasing time of exposure to BCE. Although the % tail DNA at 250 µg/ml BCE decreased with increasing time of exposure, it remained significantly elevated following 1, 2, and 4 h exposures, with 7.3-, 3.0-, and 3.1-fold increases over the control, respectively. The 1.9-fold increase in DNA damage observed in TK6 cells treated with 125 µg/ml BCE for 24 h was not statistically significant. In contrast to the significant DNA damage observed in TK6 cells after acute exposure, no DNA damage was observed in HepG2 cells exposed up to 500 µg/ml BCE at any exposure duration (0.5–24 h) (Figure 3B).

Figure 3.

Figure 3.

Open in a new tab

Evaluation of BCE-induced DNA damage in TK6 and HepG2 cells. TK6 (A) and HepG2 cells (B) were exposed to 25–250 µg/ml and 50–500 µg/ml BCE, respectively and DNA damage (% tail DNA) was detected using the comet assay at 0.5, 1, 2, 4, and 24 h time points. PC, positive control (100 µM MMS) for 4 h. The data are expressed as the mean ± SD from at least 3 independent experiments. *p < .05, **p < .01, and ***p < .001, compared with the vehicle control.

Effects of Low Folic Acid Medium on BCE-Induced Cytotoxicity and DNA Damage in TK6 Cells

TK6 cells were cultured in RPMI-1640 medium containing either a supraphysiological concentration of FA (2268 nM; standard medium) or a physiological concentration of FA (120 nM medium) for 5 days prior to the BCE treatment. No significant differences in relative cell viability were seen with either media (Figure 4A). BCE (at 250 µg/ml for 4 h) induced statistically significant higher % tail DNA in TK6 cells cultured in the standard medium using the CometChip assay (a 2.1-fold increase over the control) (Figure 4B). TK6 cells cultured in FA 120 nM medium showed a slight increase in DNA damage following a 4-h treatment (a 1.9-fold increase over the control), but it was not statistically significant. Significant levels of DNA damage were not observed in TK6 cells cultured in either type of media after 24-h exposure to BCE. Overall, no enhancement of BCE-induced cytotoxicity or DNA damage was observed in TK6 cells cultured in medium containing a physiological level (120 nM) of FA.

Figure 4.

Figure 4.

Open in a new tab

Effects of physiological folic acid on BCE-induced cytotoxicity and DNA damage in TK6 cells. TK6 cells were cultured in the standard (2268 nM folic acid [FA]) and low FA (120 nM) RPMI-1640 medium for 5 days. BCE-induced cytotoxicity and DNA damage were evaluated by the CellTiter-Blue cell viability assay (A) and the CometChip assay (B), respectively. Black bars represent TK6 cells cultured in the standard medium whereas gray bars represent TK6 cells cultured in FA 120 nM medium. The data are expressed as the mean ± SD from at least 3 independent experiments. *p < .05, **p < .01, and ***p < .001, compared with the vehicle control.

BCE-Induced Activation of Molecular Biomarkers in DNA Damage Response Pathway in TK6 Cells

Expression of proteins and post-translational modifications indicative of the DDR and cell cycle regulation—γH2A.X, p-Chk1, p-Chk2, p53, and p21—was evaluated in TK6 cells by Western blotting (Figure 5). In the standard medium, BCE induced significant and concentration-dependent increases in the phosphorylation of H2A.X (γH2A.X) following a 4-h or 24-h treatment (p ≤ .02). Phosphorylation of Chk1 and Chk2 was also increased significantly at the highest BCE concentration following a 4-h treatment (p ≤ .03), whereas a significant increase in expression was observed only for p-Chk2 at the 24-h time point (p ≤ .03). In addition, the expression of the DNA damage-induced cell cycle regulators, p53 and p21, was significantly increased after a 24-h exposure to 75  and 125 µg/ml BCE (p ≤ .01).

Figure 5.

Figure 5.

Open in a new tab

Activation of molecular biomarkers in DNA damage response pathway by BCE in TK6 cells. A, Total cellular proteins of TK6 cells cultured in the standard FA (2268 nM) or low FA (120 nM) media were extracted following a 4-h or 24-h treatment with BCE. The expression levels of γH2A.X, H2A.X, p-Chk1, p-Chk2, p53, and p21 were detected by Western blotting analysis. GAPDH was used as a loading control. B, Intensity of γH2A.X was normalized to the intensity of H2A.X. Intensities of the bands on the other bar graphs were normalized to the amount of GAPDH. The bar graphs are the mean ± SD of fold changes compared with the vehicle control from at least 3 independent experiments. *p < .05, **p < .01, and ***p < .001 versus the vehicle control. GAPDH, glyceraldehyde-3-phosphate dehydrogenase; p-Chk1, phosphorylated checkpoint kinase1; p-Chk2, phosphorylated checkpoint kinase2.

Regarding TK6 cells cultured in 120 nM FA medium, expression levels of γH2A.X, p-Chk1, p-Chk2, p53, and p21 were all comparable to levels expressed in cells cultured in the standard medium with one exception. Following a 24-h exposure, the expression of p-Chk1 was significantly increased in TK6 cells cultured in FA 120 nM medium, whereas no such increase was observed in cells cultured in the standard medium (Figs. 5A and 5B).

BCE-Induced Apoptosis in TK6 Cells

A battery of apoptosis tests was employed to evaluate BCE-induced apoptosis in TK6 cells, including nucleic acid staining with EMA, DNA fragmentation during apoptosis by the TUNEL (BrdU-Red) assay, caspase 3/7 activity, detection of c-caspase 3 and cleaved PARP (c-PARP), which is a target of caspases 3 and 7. Following a 24-h treatment, the percentage of apoptotic/necrotic cells was significantly increased at concentrations of 100  and 125 µg/ml BCE (Figure 6A). Compared with the vehicle control, the number of BrdU-positive cells in the TUNEL assay increased by 7-fold when exposed to 125 µg/ml BCE for 24 h (p ≤ .001) (Figure 6B). A 4.7-fold increase in DNA fragmentation over control was observed in cells treated with 250 µg/ml BCE for 4 h, but it was not statistically significant. In addition, caspase 3/7 activities were significantly increased following both 4 h and 24 h exposures in a concentration-dependent manner (Figure 6C). In general, BCE concentrations of 100–250 µg/ml induced markedly stronger responses at the 24-h exposure time point than those observed at the 4-h exposure time point (4.9–19.8- vs 1.7–4.5-fold increases over the control). The activation of caspase 3 was confirmed by increased expression levels of c-caspase 3 and c-PARP in the cells exposed to 250 µg/ml BCE for 4 h and 125 µg/ml BCE for 24 h (Figs. 6D and 6E). Further, the expression levels of c-caspase 3 and c-PARP were comparable between TK6 cells cultured in FA 120 nM and standard FA media.

Figure 6.

Figure 6.

Open in a new tab

BCE-induced apoptosis in TK6 cells. TK6 cells were exposed to various concentrations (25–250 µg/ml) of BCE for 4 or 24 h. BCE-induced apoptosis was evaluated by ethidium monoazide staining from the MN assay (A), the TUNEL assay (B), caspase 3/7 enzyme activity assay (C), and the expression of protein levels of cleaved caspase 3 (c-Cas3) and PARP by Western blotting (D). GAPDH was used as a loading control. (E) The bar graphs represent the densitometric analysis of c-Cas3 and cleaved-PARP (c-PARP, second band at 89 kDa). Intensities of bands were normalized to the amount of GAPDH. The data are expressed as the mean ± SD from at least 3 independent experiments. *p < .05, **p < .01, and ***p < .001 versus the vehicle control. GAPDH, glyceraldehyde-3-phosphate dehydrogenase; PARP, poly (ADP-ribose) polymerase.

BCE-Induced G1 Phase Cell Cycle Arrest in TK6 Cells

The cell cycle distribution analysis by flow cytometry demonstrated that following a 24-h treatment, a significantly altered BCE concentration-dependent cell cycle phase distribution was evident in TK6 cells (Figure 7A). At the 125 µg/ml concentration, the proportion of cells in G1 was markedly increased from 36.7% to 50.1%, accompanied by a concordant decrease in the proportion of cells in S and G2/M phases (Figure 7B), indicating a cell cycle arrest at G1 phase. In addition, a concentration-dependent increase in sub-G1 apoptotic cells was observed.

Figure 7.

Figure 7.

Open in a new tab

BCE-induced cell cycle arrest in TK6 cells. A, DNA histograms representing cell cycle distribution following a 24-h treatment with BCE (25–125 µg/ml) in TK6 cells. B, The proportion of cells (%) in various stages (subG1, G1, S and G2/M) of the cell cycle were analyzed by measuring the nuclear DNA content of treated cells.

DNA Methylation in TK6 and HepG2 Cells After BCE Treatment

Folate deficiency is associated with global DNA hypomethylation (Crider et al., 2012). Therefore, the level (%) of LINE-1 methylation was assessed as a representative marker for global DNA methylation in TK6 and HepG2 cells. The MeDIP qPCR data showed that there were no significant differences, compared with the vehicle control, in LINE1 DNA methylation (%) of region 1 and 2 in TK6 or in HepG2 cells treated with BCE at various concentrations for 72 h, indicating that BCE did not induce significant changes in global DNA methylation (%) in these cells, at least when cells are cultured in supraphysiological levels of FA (Figure 8).

Figure 8.

Figure 8.

Open in a new tab

Evaluation of BCE-induced global DNA methylation in TK6 and HepG2 cells. DNA methylation of long interspersed nucleotide elements (LINE) 1 was assessed in TK6 (A) and HepG2 cells (B) using the methylated DNA immunoprecipitation (MeDIP) assay combined with real-time quantitative PCR (qPCR) for LINE1 region 1 and region 2. The data are expressed as the mean ± SD from at least 3 independent experiments. The values were expressed as the percentage (%) of methylated DNA in total DNA.

DISCUSSION

In the present study, genotoxic and cytotoxic effects of BCE were investigated using two different human cell lines, lymphoblastoid TK6 cells and hepatoma HepG2 cells. We first confirmed that BCE induced concentration-dependent increases in %MN in TK6 cells following a 24-h exposure (Figure 1A), an observation consistent with previous findings (Bernacki et al., 2019; Smith-Roe et al., 2018). A marginal but statistically significant increase in MN induction also was observed in HepG2 cells, only at concentrations of 200 µg/ml (Figure 1B). TK6 cells were also found to be more sensitive to BCE-induced cytotoxicity and DNA damage in the comet assay than HepG2 cells (Figs. 2 and 3). TK6 cells are not metabolically competent, whereas HepG2 cells express low levels of phase I cytochrome P450 enzymes and normal levels of most phase II antioxidant/detoxifying enzymes (Westerink and Schoonen, 2007). It has been reported that BCE-induced MN formation was attenuated in the presence of induced rat S9 (Smith-Roe et al., 2018). Therefore, HepG2 cells may have some capacity for metabolizing BCE and reducing its bioactivity. These differences in metabolic capacity may have contributed to the different levels of BCE-induced cytotoxicity, DNA damage, and MN induction observed between TK6 and HepG2 cells in our study.

The most interesting finding was that BCE induced significant DNA damage after a short 0.5-h exposure in TK6 cells and DNA damage decreased with increasing exposure duration, suggesting active DNA repair (Figure 3A). Although the level of DNA damage seen at the 2-h time point was markedly reduced, statistically significant increases in % tail DNA compared with the vehicle controls were observed in TK6 cells exposed to 250 µg/ml BCE for up to 4 h. In contrast, no significant responses in the comet assay were observed at any of the time points tested with HepG2 cells (Figure 3B). These findings demonstrate that although the comet assay can be conducted using any cell line at any time point, sampling time and cell type are critical factors to consider for interpreting responses in this assay (Sasaki et al., 2007).

As a clear acute DDR was observed in TK6 cells after BCE exposure in the comet assay, the expression of γH2A.X, a sensitive biomarker for DNA double-strand breaks, was examined following a 4-h or 24-h treatment. BCE-induced DNA damage in TK6 cells was confirmed by concentration-dependent increases in the expression of γH2A.X after 4 h and 24 h exposures (Figure 5). It has been reported that aneugens predominantly induce apoptotic γH2A.X (γH2A.X+/caspase 3+) stemming from cell cycle arrest in the M-phase, whereas clastogens mainly induce nonapoptotic γH2A.X (γH2A.X+/caspase 3−) (Harada et al., 2014). In our study, in addition to the elevated level of γH2A.X expression, BCE treatment also significantly increased apoptotic and necrotic cells, caspase 3/7 activity, protein expression of c-caspases 3 and PARP cleavage, and DNA fragmentation during apoptosis as measured by the TUNEL assay (Figure 6). These observations support an aneugenic MOA for BCE, which is in agreement with previous NTP findings showing a significant increase in phospho-histone H3 (p-H3)-positive events and p53 translocation to the nucleus with BCE exposure (Bernacki et al., 2019; Smith-Roe et al., 2018).

Due to the possibility that BCE disrupts folate metabolism (Cora et al., 2017) and the concern that excess FA could mask the toxic effects of BCE, we cultured TK6 cells in media containing physiological (120 nM) and standard (2268 nM) levels of FA and evaluated DDRs. Growth in physiological levels of FA did not appear to potentiate the degree of DNA damage observed in the comet assay at 4 h or 24 h (Figure 4). In addition, FA concentration had no effects on the levels of apoptosis and DDR pathway protein expression tested in the present study, except for p-Chk1, whose expression significantly increased in the cells cultured in 120 nM FA medium compared with the cells in the standard medium (Figs. 5 and 6).

The underlying mechanism of BCE’s toxicity was further explored in TK6 cells. Chk1 and Chk2 are essential genome integrity checkpoint proteins expressed in response to diverse genotoxic insults (Bartek and Lukas, 2003). Phosphorylation of Chk1 and Chk2 is involved in cell cycle arrest at G1/S and G2/M boundaries, DNA repair, or apoptosis in response to DNA damage (Patil et al., 2013; Zannini et al., 2014). In addition, γH2A.X is an initiating event in the activation of the p53/p21 pathway in the DDR, with upregulation of p21 resulting in cell cycle arrest to allow repair of DNA lesions or initiation of apoptosis (Fragkos et al., 2009). Our study demonstrated a G1 phase cell cycle arrest with an increase in the sub-G1 cell population (Figure 7) and significant increases in expression levels of p-Chk1, p-Chk2, p53, and p21 proteins in TK6 cells exposed to BCE (Figure 5). Consistent with our findings, it was previously reported that BCE induced G1 cell cycle arrest in human breast cancer cells (MCF-7) exposed to 30 µg/ml BCE (Einbond et al., 2004). Furthermore, treatment of MCF-7 cells with 15 μg/ml BCE for 24 h induced downregulation of the expression of gene products involved in the transition from G1 to S-phase (Gaube et al., 2007).

As mentioned earlier, BCE was shown to be aneugenic by destabilizing microtubules (Bernacki et al., 2019). Consistent with this observation, a microarray analysis showed that BCE clustered with antiproliferative compounds, specifically tubulin binding vinca alkaloids and DNA alkylators, in rat liver tissue (Einbond et al., 2012). The NTP study demonstrated that BCE had an aneugenic MOA when evaluated by the MultiFlow DNA Damage Assay, as BCE induced marked increases in p-H3 accompanied by p53 activation in TK6 cells whereas no significant γH2A.X induction was observed (Smith-Roe et al., 2018). In the present study, however, we observed that exposure to BCE induced γH2A.X at the 4 h and 24 h time points (Figure 5). Although these results may appear to be at odds with Smith-Roe et al. (2018), we note that increases in γH2A.X at these time points occurred when BCE-induced DNA damage appeared to be repaired (Figure 3), but also when markers of apoptosis were expressed (Figure 5), indicating that the γH2A.X is more likely due to apoptosis as opposed to chemically induced DNA damage. The flow cytometric approach used in Smith-Roe et al. (2018) would have gated out apoptotic cells that expressed γH2A.X (Dertinger et al., 2019; Huang et al., 2004).

Our results support aneugenicity as the main genotoxic MOA for BCE, considering that BCE was negative in the Ames bacterial reverse mutation assay (Smith-Roe et al., 2018) and the observation that significant DDR pathway biomarkers increased at concentrations that induced apoptosis in the present study (Figs. 5 and 6). BCE induced acute DNA damage only in TK6 cells but not in HepG2 cells, which have low levels of metabolic capacity, and the damage was quickly repaired (Figure 3). In addition, BCE at concentrations up to 50 µg/ml in TK6 cells or 150 µg/ml in HepG2 cells exposed for 72 h showed no potential for inducing methylation-based epigenetic modifications, as no alterations in global DNA methylation were observed in either TK6 or HepG2 cells, when cells were grown in media with typical, supraphysiological amounts of FA (Figure 8). Because of the longer exposure time required for this assay, higher concentrations of BCE could not be tested due to severe cytotoxicity induced by BCE in these cell lines (Supplementary Figure 1). As a complex mixture containing more than 100 chemicals, it is possible that multiple mechanisms may be involved in BCE-induced genotoxicity and cytotoxicity, mediated by individual chemicals and chemical interactions. Also, commercially available BCE preparations differ markedly in composition, suggesting the possibility of highly variable bioactivities among these complex commercial products. Fully characterizing the genotoxicity of BCE is important because others have calculated that women could potentially be exposed to doses of BCE that were shown to significantly increase MN in mice (Bernacki et al., 2019). Our next steps are to further understand whether metabolism affects the genotoxicity induced by BCE and its components using metabolically competent human cell models, such as HepaRG human liver cells.

SUPPLEMENTARY DATA

Supplementary data are available at Toxicological Sciences online.

DECLARATION OF CONFLICTING INTERESTS

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Supplementary Material

kfab044_Supplementary_Data
Click here for additional data file. (182.7KB, docx)

ACKNOWLEDGMENTS

This work was supported by the U.S. Food and Drug Administration (FDA), National Center for Toxicological Research (NCTR, project number E00220201), and in part by the U.S. NIH, National Institute of Environmental Health Sciences (NIEHS). J.E.S. and X.L. were supported by appointments to the Postgraduate Research Program at the NCTR administered by the Oak Ridge Institute for Science Education (ORISE) through an interagency agreement between the U.S. Department of Energy and the U.S. FDA. We thank Dr Javier Revollo, Dr Robert Heflich, and Dr Page McKenzie at NCTR, and Dr Sreenivasa Ramaiahgari at NTP, for their critical review of this article.

FUNDING

National Toxicology Program under an Interagency Agreement between FDA (FDA IAG No. 224-17-0502); NIEHS (NIEHS IAG No. AES12013).

 

DISCLAIMER

The information in this article is not a formal dissemination of information by the U.S. FDA or NIEHS and does not represent the agency position or policy.

REFERENCES

  1. Azqueta A., Collins A. R. (2013). The essential comet assay: A comprehensive guide to measuring DNA damage and repair. Arch. Toxicol. 87, 949–968. [DOI] [PubMed] [Google Scholar]
  2. Bartek J., Lukas J. (2003). Chk1 and Chk2 kinases in checkpoint control and cancer. Cancer Cell 3, 421–429. [DOI] [PubMed] [Google Scholar]
  3. Bernacki D. T., Bryce S. M., Bemis J. C., Dertinger S. D., Witt K. L., Smith-Roe S. L. (2019). Evidence for an aneugenic mechanism of action for micronucleus induction by black cohosh extract. Environ. Mol. Mutagen. 60, 845–856. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Blount B. C., Mack M. M., Wehr C. M., MacGregor J. T., Hiatt R. A., Wang G., Wickramasinghe S. N., Everson R. B., Ames B. N. (1997). Folate deficiency causes uracil misincorporation into human DNA and chromosome breakage: Implications for cancer and neuronal damage. Proc. Nat. Acad. Sci. U.S.A. 94, 3290–3295. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Chow E., Teo M., Ring J. A., Chen J. W. (2008). Liver failure associated with the use of black cohosh for menopausal symptoms. Med. J. Aust. 188, 420–422. [DOI] [PubMed] [Google Scholar]
  6. Cora M. C., Gwinn W., Wilson R., King D., Waidyanatha S., Kissling G. E., Brar S. S., Olivera D., Blystone C., Travlos G. (2017). A black cohosh extract causes hematologic and biochemical changes consistent with a functional cobalamin deficiency in female B6C3F1/N mice. Toxicol. Pathol. 45, 614–623. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Crider K. S., Yang T. P., Berry R. J., Bailey L. B. (2012). Folate and DNA methylation: A review of molecular mechanisms and the evidence for folate’s role. Adv. Nutr. 3, 21–38. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Dertinger S. D., Kraynak A. R., Wheeldon R. P., Bernacki D. T., Bryce S. M., Hall N., Bemis J. C., Galloway S. M., Escobar P. A., Johnson G. E. (2019). Predictions of genotoxic potential, mode of action, molecular targets, and potency via a tiered multiflow(r) assay data analysis strategy. Environ. Mol. Mutagen. 60, 513–533. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Einbond L. S., Shimizu M., Xiao D., Nuntanakorn P., Lim J. T., Suzui M., Seter C., Pertel T., Kennelly E. J., Kronenberg F., et al. (2004). Growth inhibitory activity of extracts and purified components of black cohosh on human breast cancer cells. Breast Cancer Res. Treat. 83, 221–231. [DOI] [PubMed] [Google Scholar]
  10. Einbond L. S., Soffritti M., Esposti D. D., Wu H. A., Tibaldi E., Lauriola M., He K., Park T., Su T., Huggins L., et al. (2012). Pharmacological mechanisms of black cohosh in Sprague-Dawley rats. Fitoterapia 83, 461–468. [DOI] [PubMed] [Google Scholar]
  11. Everson R. B., Wehr C. M., Erexson G. L., MacGregor J. T. (1988). Association of marginal folate depletion with increased human chromosomal damage in vivo: Demonstration by analysis of micronucleated erythrocytes. J. Nat. Cancer Inst. 80, 525–529. [DOI] [PubMed] [Google Scholar]
  12. Fenech M. (1999). Micronucleus frequency in human lymphocytes is related to plasma vitamin B12 and homocysteine. Mutat. Res. Fund. Mol. Mech. Mutagen. 428, 299–304. [DOI] [PubMed] [Google Scholar]
  13. Fragkos M., Jurvansuu J., Beard P. (2009). H2AX is required for cell cycle arrest via the p53/p21 pathway. Mol. Cell. Biol. 29, 2828–2840. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Gaube F., Wolfl S., Pusch L., Kroll T. C., Hamburger M. (2007). Gene expression profiling reveals effects of Cimicifuga racemosa (L.) NUTT. (black cohosh) on the estrogen receptor positive human breast cancer cell line MCF-7. BMC Pharmacol. 7, 11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Guo X., Chen S., Zhang Z., Dobrovolsky V. N., Dial S. L., Guo L., Mei N. (2015). Reactive oxygen species and c-Jun N-terminal kinases contribute to TEMPO-induced apoptosis in L5178Y cells. Chem. Biol. Interact. 235, 27–36. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Guo X., Seo J. E., Bryce S. M., Tan J. A., Wu Q., Dial S. L., Moore M. M., Mei N. (2018). Comparative genotoxicity of TEMPO and 3 of its derivatives in mouse lymphoma cells. Toxicol. Sci. 163, 214–225. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Guo X., Seo J. E., Petibone D., Tryndyak V., Lee U. J., Zhou T., Robison T. W., Mei N. (2020). Performance of HepaRG and HepG2 cells in the high-throughput micronucleus assay for in vitro genotoxicity assessment. J. Toxicol. Environ. Health A. 83, 702–717. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Guzman G., Kallwitz E. R., Wojewoda C., Chennuri R., Berkes J., Layden T. J., Cotler S. J. (2009). Liver injury with features mimicking autoimmune hepatitis following the use of black cohosh. Case Rep. Med. 2009, 918156. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Harada A., Matsuzaki K., Takeiri A., Mishima M. (2014). The predominant role of apoptosis in γH2AX formation induced by aneugens is useful for distinguishing aneugens from clastogens. Mutat. Res. Gene. Toxicol. Environ. Mutagen. 771, 23–29. [DOI] [PubMed] [Google Scholar]
  20. Huang X., Okafuji M., Traganos F., Luther E., Holden E., Darzynkiewicz Z. (2004). Assessment of histone H2AX phosphorylation induced by DNA topoisomerase I and II inhibitors topotecan and mitoxantrone and by the DNA cross-linking agent cisplatin. Cytometry A 58, 99–110. [DOI] [PubMed] [Google Scholar]
  21. Jiang B., Kronenberg F., Nuntanakorn P., Qiu M. H., Kennelly E. J. (2006). Evaluation of the botanical authenticity and phytochemical profile of black cohosh products by high-performance liquid chromatography with selected ion monitoring liquid chromatography-mass spectrometry. J. Agric Food Chem. 54, 3242–3253. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Leach M. J., Moore V. (2012). Black cohosh (Cimicifuga spp.) for menopausal symptoms. Cochrane Database Syst. Rev. CD007244. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Lin H., Guo X., Zhang S., Dial S. L., Guo L., Manjanatha M. G., Moore M. M., Mei N. (2014). Mechanistic evaluation of ginkgo biloba leaf extract-induced genotoxicity in L5178Y cells. Toxicol. Sci. 139, 338–349. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Lynch C. R., Folkers M. E., Hutson W. R. (2006). Fulminant hepatic failure associated with the use of black cohosh: A case report. Liver Transpl. 12, 989–992. [DOI] [PubMed] [Google Scholar]
  25. MacGregor J. T., Wehr C. M., Hiatt R. A., Peters B., Tucker J. D., Langlois R. G., Jacob R. A., Jensen R. H., Yager J. W., Shigenaga M. K. (1997). Spontaneous’ genetic damage in man: Evaluation of interindividual variability, relationship among markers of damage, and influence of nutritional status. Mutat. Res. Fund. Mol. Mech. Mutagen. 377, 125–135. [DOI] [PubMed] [Google Scholar]
  26. Mercado-Feliciano M., Cora M. C., Witt K. L., Granville C. A., Hejtmancik M. R., Fomby L., Knostman K. A., Ryan M. J., Newbold R., Smith C., et al. (2012). An ethanolic extract of black cohosh causes hematological changes but not estrogenic effects in female rodents. Toxicol. Appl. Pharmacol. 263, 138–147. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Muqeet Adnan M., Khan M., Hashmi S., Hamza M., AbdulMujeeb S., Amer S. (2014). Black cohosh and liver toxicity: Is there a relationship? Case Rep. Gastrointest. Med. 2014, 1–3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. OECD. (2016). In vitro mammalian cell micronucleus test. OECD guideline for the testing of chemicals. TG 487. https://www.oecd-ilibrary.org/environment/test-no-487-in-vitro-mammalian-cell-micronucleus-test_9789264264861-en. Accessed February 17, 2021.
  29. Patil M., Pabla N., Dong Z. (2013). Checkpoint kinase 1 in DNA damage response and cell cycle regulation. Cell Mol. Life Sci.70, 4009–4021. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Pierard S., Coche J.-C., Lanthier P., Dekoninck X., Lanthier N., Rahier J., Geubel A. P. (2009). Severe hepatitis associated with the use of black cohosh: A report of two cases and an advice for caution. Eur. J. Gastroenterol. Hepatol. 21, 941–945. [DOI] [PubMed] [Google Scholar]
  31. Pockaj B. A., Gallagher J. G., Loprinzi C. L., Stella P. J., Barton D. L., Sloan J. A., Lavasseur B. I., Rao R. M., Fitch T. R., Rowland K. M., et al. (2006). Phase III double-blind, randomized, placebo-controlled crossover trial of black cohosh in the management of hot flashes: NCCTG trial N01CC1. J. Clin. Oncol. 24, 2836–2841. [DOI] [PubMed] [Google Scholar]
  32. Reed S. D., Newton K. M., LaCroix A. Z., Grothaus L. C., Grieco V. S., Ehrlich K. (2008). Vaginal, endometrial, and reproductive hormone findings: Randomized, placebo-controlled trial of black cohosh, multibotanical herbs, and dietary soy for vasomotor symptoms: The herbal alternatives for menopause (halt) study. Menopause 15, 51–58. [PubMed] [Google Scholar]
  33. Ruhlen R. L., Haubner J., Tracy J. K., Zhu W., Ehya H., Lamberson W. R., Rottinghaus G. E., Sauter E. R. (2007). Black cohosh does not exert an estrogenic effect on the breast. Nutr. Cancer 59, 269–277. [DOI] [PubMed] [Google Scholar]
  34. Ruhlen R. L., Sun G. Y., Sauter E. R. (2008). Black cohosh: Insights into its mechanism(s) of action. Integr. Med. Insights 3, 21–32. [PMC free article] [PubMed] [Google Scholar]
  35. Sasaki Y. F., Nakamura T., Kawaguchi S. (2007). What is better experimental design for in vitro comet assay to detect chemical genotoxicity. AATEX 14, 499–504. [Google Scholar]
  36. Seo J.-E., Tryndyak V., Wu Q., Dreval K., Pogribny I., Bryant M., Zhou T., Robison T. W., Mei N., Guo X. (2019). Quantitative comparison of in vitro genotoxicity between metabolically competent HepaRG cells and HepG2 cells using the high-throughput high-content CometChip assay. Arch. Toxicol. 93, 1433–1448. [DOI] [PubMed] [Google Scholar]
  37. Shulman L. P., Banuvar S., Fong H. H., Farnsworth N. R. (2011). Discussion of a well-designed clinical trial which did not demonstrate effectiveness: UIC center for botanical dietary supplements research study of black cohosh and red clover. Fitoterapia 82, 88–91. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Smith-Roe S. L., Swartz C. D., Shepard K. G., Bryce S. M., Dertinger S. D., Waidyanatha S., Kissling G. E., Auerbach S. S., Witt K. L. (2018). Black cohosh extracts and powders induce micronuclei, a biomarker of genetic damage, in human cells. Environ. Mol. Mutagen. 59, 416–426. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Smith T., Gillespie M., Eckl V., Knepper J., Reynolds C. (2019). Herbal supplement sales in us increase by 9.4% in 2018. HerbalGram 123, 62–73. [Google Scholar]
  40. Tice R. R., Agurell E., Anderson D., Burlinson B., Hartmann A., Kobayashi H., Miyamae Y., Rojas E., Ryu J. C., Sasaki Y. F. (2000). Single cell gel/comet assay: Guidelines for in vitro and in vivo genetic toxicology testing. Environ. Mol. Mutagen. 35, 206–221. [DOI] [PubMed] [Google Scholar]
  41. van der Sluijs C. P., Bensoussan A., Chang S., Baber R. (2009). A randomized placebo-controlled trial on the effectiveness of an herbal formula to alleviate menopausal vasomotor symptoms. Menopause 16, 336–344. [DOI] [PubMed] [Google Scholar]
  42. Wang Y. J., Dou J., Cross K. P., Valerio L. G. Jr. (2011). Computational analysis for hepatic safety signals of constituents present in botanical extracts widely used by women in the United States for treatment of menopausal symptoms. Regul. Toxicol. Pharmacol. 59, 111–124. [DOI] [PubMed] [Google Scholar]
  43. Westerink W. M., Schoonen W. G. (2007). Cytochrome p450 enzyme levels in HepG2 cells and cryopreserved primary human hepatocytes and their induction in HepG2 cells. Toxicol. In Vitro 21, 1581–1591. [DOI] [PubMed] [Google Scholar]
  44. Wickramasinghe S. N. (2006). Diagnosis of megaloblastic anaemias. Blood Rev. 20, 299–318. [DOI] [PubMed] [Google Scholar]
  45. Zannini L., Delia D., Buscemi G. (2014). Chk2 kinase in the DNA damage response and beyond. J. Mol. Cell Biol. 6, 442–457. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

kfab044_Supplementary_Data
Click here for additional data file. (182.7KB, docx)

Articles from Toxicological Sciences are provided here courtesy of Oxford University Press

RESOURCES