C3 exoenzyme impairs cell proliferation and apoptosis by altering the activity of transcription factors
C3 exoenzyme from C. botulinum is an ADP-ribosyltransferase that inactivates selectively RhoA, B, and C by coupling an ADP-ribose moiety. Rho-GTPases are involved in various cellular processes, such as regulation of actin cytoskeleton, cell proliferation, and apoptosis. Previous studies of our group with the murine hippocampal cell line HT22 revealed a C3-mediated inhibition of cell proliferation after 48 h and a prevention of serum-starved cells from apoptosis. For both effects, alterations of various signaling pathways are already known, including also changes on the transcriptional level. Investigations on the transcriptional activity in HT22 cells treated with C3 for 48 h identified five out of 48 transcription factors namely Sp1, ATF2, E2F-1, CBF, and Stat6 with a significantly regulated activity. For validation of identified transcription factors, studies on the protein level of certain target genes were performed. Western blot analyses exhibited an enhanced abundance of Sp1 target genes p21 and COX-2 as well as an increase in phosphorylation of c-Jun. In contrast, the level of p53 and apoptosis-inducing GADD153, a target gene of ATF2, was decreased. Our results reveal that C3 regulates the transcriptional activity of Sp1 and ATF2 resulting downstream in an altered protein abundance of various target genes. As the affected proteins are involved in the regulation of cell proliferation and apoptosis, thus the C3-mediated anti-proliferative and anti-apoptotic effects are consequences of the Rho-dependent alterations of the activity of certain transcriptional factors.
KeywordsC3 exoenzyme RhoA Proliferation Apoptosis Transcription factors
C3 exoenzyme from Clostridium botulinum
Specificity protein 1
Activating transcription factor 2
E2F transcription factor 1
CCAAT/enhancer binding protein (C/EBP), zeta
Signal transducer and activator of transcription 6
Growth Arrest and DNA Damage-inducible protein 153
C3 exoenzyme from Clostridium botulinum (C3) belongs to the group of eight bacterial ADP-ribosyltransferases including C3lim from Clostridium limosum, C3stau from Staphylococcus aureus, C3cer from Bacillus cereus, and C3larvin from Paenibacillus larvae that possess low molecular weight Rho-GTPases as substrates (Aktories and Frevert 1987; Just et al. 1992a; Just et al. 1992b; Wilde et al. 2001; Krska et al. 2015). C3 selectively inactivates the Rho-GTPases RhoA, B, and C by transferring an ADP-ribose moiety from NAD+ onto asparagine 41 of Rho (Chardin et al. 1989; Sekine et al. 1989). This resulting loss of functional Rho causes cellular consequences such as disorganization of the actin cytoskeleton, morphological changes, and impaired formation of contractile ring (Wiegers et al. 1991; Kishi et al. 1993). Because of its specificity, C3 is often applied as a selective Rho inhibitor in studying cellular RhoA signaling. Furthermore, the treatment of murine primary hippocampal neurons with C3 reveals an increased axonal growth as well as branching independently of the enzyme activity and an additional dendritotrophic effect of the C3 wild type (Ahnert-Hilger et al. 2004). Moreover, previous studies demonstrated that Rho inactivation by C3 inhibits cell growth in various cell types (Nishiki et al. 1990; Yamamoto et al. 1993; Zuckerbraun et al. 2003; Rohrbeck et al. 2012). RhoA is associated with the regulation of various proteins involved in the control of cell cycle progression like cyclin D1 and p21 (Adnane et al. 1998; Watts et al. 2006). Additionally, RhoA modulates the activity of certain transcription factors known to play a major role in the regulation of cell proliferation. For example, the overexpression of constitutively active RhoAQ63L increases the transcriptional activity of AP-1 and E2F in NIH3T3 cells (Berenjeno et al. 2007). Interestingly, in murine hippocampal HT22 cells, both C3 and enzyme-deficient C3-E174Q mediate inhibition of proliferation that was accompanied by a reduced level of cyclin D1 and increased expression of negative cell cycle regulator RhoB (Du and Prendergast 1999; Rohrbeck et al. 2012).
Besides the inhibition of cell proliferation, previous studies described an influence of C3 on apoptosis in various cell types. Depending on the cell type, C3 is able to trigger apoptosis in EL4 T lymphoma, HUVEC, and hepatic stellate cells (Moorman et al. 1996; Li et al. 2002; Ikeda et al. 2003). Contrary, treatment of astrocytes with C3 after induction of apoptosis with thrombin increases the amount of surviving cells (Donovan et al. 1997). Furthermore, the in vivo application of C3 protects retinal ganglion cells from apoptosis induced either after optic nerve injury or by injection of NMDA (Bertrand et al. 2005; Wang et al. 2014). The injection of C3 on the lesion site decreases the number of apoptotic cells after a spinal cord injury in rodents (Dubreuil et al. 2003). Rohrbeck et al. reported that the prevention of serum-starved and staurosporin-treated HT22 cells from apoptosis is accompanied by the C3-mediated reduction of pro-apoptotic proteins and of the activity of various caspases. Indeed, this anti-apoptotic effect depends on Rho because enzyme-deficient C3-E174Q is without effect (Rohrbeck et al. 2012).
In the present study, we investigated the impact of C3 on the transcriptional level and downstream proteins in HT22 cells. These conditions were selected due to the appearance of C3-mediated inhibition of cell proliferation after 48 h. We demonstrated that C3 Rho-dependently modulated the activity of transcription factors as well as the protein abundance of certain target genes that were associated with the regulation of cell proliferation and apoptosis. Thus, these results strongly indicate that the C3-mediated anti-proliferative and anti-apoptotic effects are mediated by alterations of transcriptional and protein level as a consequence of Rho inactivation by C3.
Materials and methods
The murine hippocampal cell line HT22 was cultivated in Dulbecco’s modified essential medium ((Gibco, Life Technologies, Paisley, UK), 10 % fetal bovine serum (PAN Biotech GmbH, Aidenbach, Germany), 1 % penicillin, 1 % streptomycin (PAA Laboratories GmbH, Pasching, Austria), and 1 mM sodium pyruvate (Biochrom AG, Berlin, Germany)) at 37 °C and 5 % CO2. When the cells reached confluence, they were passaged.
For growth kinetics experiments, 30,000 cells∙mL−1 were seeded onto 3.5-cm plates. After 24 h, the cells were treated with 500 nM C3, C3-E174Q, or 20 nM skepinone-L. Every 48 h the medium was replaced including C3 or C3-E174Q. The determination of cell number was performed as described previously (Rohrbeck et al. 2012).
The isolation of RNA, primer design, and determination of gene expression level of p21 by the use of real-time qRT-PCR measurements were accomplished as described prior (Rohrbeck et al. 2012). The following primer pairs were applied for qRT-PCR: p21/Cdkn1 (NM_007669.4) forward: GTACTTCCTCTGCCCTGCTG; reverse: GGCACTTCAGGGTTTTCTC, B2M (NM_009735.3) forward: ATTCACCCCCACTGAGACTG; reverse: GCTATTTCTTTCTGCGTGCAT. PCR primers were acquired by Eurofins (Ebersberg, Germany).
Western blot analysis
The cells were seeded onto 3.5-cm plates with a concentration of 150,000 cells∙mL−1. The next day, cells were treated with 500 nM C3, C3-E174Q, or indicated concentrations of inhibitors NSC23766 and skepinone-L (Calbiochem, Merck KGaA, Darmstadt, Germany) for various incubation times. After termination of incubation, the cells were washed with ice-cold PBS and frozen at −20 °C. Preparation of cell lysates and Western blot analysis was performed as described previously (Rohrbeck et al. 2012). For the analysis of phosphorylated proteins, 1 mM sodium-ortho-vanadate (Sigma-Aldrich Chemie GmbH, Munich, Germany) was applied in lysis buffer. The following primary antibodies were applied for immunoblotting: α-RhoA, α-p38, α-JNK1, α-p21, and α-GADD153 (Santa Cruz Biotechnology, CA, USA); α-β-Actin (Sigma-Aldrich, St. Louis, MO, USA); and α-pp38 Thr180/182, α-p-c-Jun Ser63, α-COX-2, and α-p53 (Cell Signaling Technology, Beverly, MA, USA). The chemiluminescence reaction was performed by ECL Femto (Pierce, Thermo Fisher Scientific Inc., Rockford, IL, USA), and the signals were detected and analyzed densitometrical by Kodak 1D software (KODAK GmbH, Stuttgart, Germany).
TF activation profiling plate array
For screening the transcriptional activity of 48 different transcription factors after treatment with C3 for 48 h, the TF Activation Profiling Plate Array I (Signosis Inc., Santa Clara, CA, USA) was performed. HT22 cells were incubated with 500 nM C3 or medium for control conditions. After 48 h, the nuclear extraction (Signosis Inc., Santa Clara, CA, USA) was performed according to manufacturers’ instructions. The protein concentration was determined by Bradford assay, and 5 μg of nuclear extracts per condition were applied in TF Activation Profiling Plate Array I according to manufacturers’ instructions. Both conditions were measured on one 96-well plate containing two sets for each 48 transcription factors. The luminescence was detected at Synergy4 microplate reader (BioTek Instruments Inc., Winooski, VT, USA). For each condition, the relative light units of the transcription factors were normalized to the value of the non-regulated SATB1 as internal control. The relative regulation was calculated by the ratio of C3-treatment in comparison to control condition. Significant regulations were estimated in a twofold increase or decrease of transcriptional activity. Transcription factors whose activity was altered in all three experiments significantly in the same direction were defined as regulated.
Luciferase reporter experiments
The dual-luciferase reporter experiments were performed with the Cignal Reporter Assay Kit (Qiagen, Hilden, Germany). The reporter system consists of a firefly luciferase reporter under the control of an inducible basal TATA box promotor, with upstream tandem repeat elements (TRE)-sequences for Sp1, and as an internal control, a construct that constitutively expressed Renilla luciferase under the control of a CMV immediate early enhancer/promotor in a ratio of 40:1. For detection of background signals, a negative control construct that encodes the firefly luciferase under a non-inducible basal TATA box promotor and a constitutively expressed Renilla luciferase (in a ratio of 40:1) were applied. 7500 HT22 cells per well were seeded into 96-well plates. The cells were transfected with 1 μg DNA construct of either transcription factor reporter or negative control by the use of jetPrime Polyplus transfection system (Polyplus transfection S.A., Illkirch, France) according to manufacturers’ instructions. After 4 h, the cells were treated with 500 nM C3, 500 nM C3-E174Q, 20 nM skepinone-L, or 50 μM NSC23766 for 48 h. To attain a stimulation of Sp1 activity, cells were incubated with 100 ng/mL PMA (Sigma-Aldrich Chemie GmbH, Munich, Germany) for 18 h as a positive control. The luciferase activity was determined by Dual-Glo® Luciferase assay system (Promega Corporation, Madison, WI, USA) on Synergy4 microplate reader (BioTek Instruments Inc., Winooski, VT, USA). Data were processed by normalizing the relative light units of firefly to Renilla luciferase, subtracting background signals and calculating the relative regulation of transcriptional activity. To determinate the effectivity of transfection, cells were transfected with a positive control reporter containing a construct that encodes GFP. The cells were visualized by light and fluorescence microscopy (Zeiss Axiovert 200 M; Carl Zeiss GmbH, Göttingen, Germany).
Expression and purification of recombinant C3 proteins
C3 wild type and C3-E174Q were expressed as recombinant fusion proteins with a glutathione S-transferase (GST)-tag into plasmid pGEX-2T (gene of C. botulinum C3, accession no. X59039) that was transferred into E. coli TG1. The purification of recombinant protein was performed as described previously (Rohrbeck et al. 2012).
Reproducibility of the experiments and statistics
All experiments were performed independently at least three times. The figures display results from representative experiments. For graphical and statistical analysis, Microsoft® Excel 2010 version 14.0 (Microsoft Corporation, Redmond, USA) was applied. The values (n ≥ 3) are means ± SEM. The statistical significance of differences between treated compared to untreated cells were calculated by the use of a two-sided unpaired Student’s t test (* = p ≤ 0.05, ** = p ≤ 0.01, and *** = p ≤ 0.001). The statistical differences between treated compared to untreated cells in qRT-PCR experiments were calculated by the use of a one-sided unpaired Student’s t test (* = p ≤ 0.05).
C3 altered the transcriptional activity of ATF2 and Sp1
For validation of the identified transcription factors, the downstream target genes of Sp1 and ATF2 were analyzed. Western blot analyses of the three different Sp1 target genes p21, c-Jun, and cyclooxygenase (COX)-2 harboring at least one Sp1 binding site in their gene promotor were performed (Rozek and Pfeifer 1993; Appleby et al. 1994; Datto et al. 1995; Rozek and Pfeifer 1995; Biggs et al. 1996; Xu et al. 2000). In case of c-Jun, we focused on the activation in terms of a phosphorylation of c-Jun. The enzyme-deficient C3-E174Q was carried along as a negative control, as it did not provoke any alterations in transcriptional activity of Sp1. Additionally, the effects of Rac and p38 inhibitors on the target proteins were determined.
C3-induced increase of p21 and anti-proliferative effect
As p21 is a major regulator of cell cycle progression; the effect of C3 on cell proliferation was examined by growth kinetic experiments in a concentration-dependent manner (Fig. 2d). Incubation with 100 nM of C3 did not impair the proliferation of HT22 cells. A fivefold raise of the concentration of C3 caused an inhibition of cell growth from the second day of treatment. Interestingly, a tenfold increase in concentration to about 5 μM of C3 did not further enhance the observed proliferation inhibition shown in Fig. 2d nor resulted in any signs of cellular toxicity. The temporal delay of 48 h of the C3-mediated inhibition of cell proliferation was independently of the cellular growth phase and only correlated with the incubation time with C3 (Fig. S2). In contrast, the enzyme-deficient C3-E174Q induced a Rho-independent, medium inhibition of cell proliferation in HT22 cells (Rohrbeck et al. 2012). The p38 inhibitor skepinone-L inhibited the proliferation of HT22 cells moderately starting from the second day of incubation (Fig. S3). A combined incubation of C3 and skepinone-L provoked a minimum increased anti-proliferative effect compared to the single C3 treatment confirming the involvement of p38 in the C3-mediated anti-proliferative effect.
C3 induced increase in phosphorylation of c-Jun and reduced the level of p53
C3 modulated the level of COX-2 biphasically
Surprisingly, C3 reduced distinctly and significantly the abundance of COX-2 by more than 60 % after 60 and 72 h. Incubation with skepinone-L from 60 h on lowered slightly the COX-2 level by 20 %, but no effect was detectable after treatment with C3-E174Q for 60 and 72 h. The decrease in COX-2 starting from 60 h revealed a biphasic modulation mediated by C3. Due to the weak signal intensity at Western blot analysis, the quantification of COX-2 after 24 h was not reliable.
Taken collectively, the increased protein abundances of p21, phospho-c-Jun, and COX-2 after treatment with C3 for 48 h indicated an enhanced activity of Sp1.
C3-induced alterations in p38 activity and reduction of GADD153
In a rat spinal cord injury model, the expression of Sp1 target gene COX-2, a marker protein of inflammation, is increased up to 48 h (Appleby et al. 1994; Resnick et al. 1998; Xu et al. 2000). Indeed, the expression of COX-2 is not only induced by inflammation and after injuries but COX-2 is also constitutively expressed on a basal level in neuronal cells of the spinal cord and certain areas of the brain (Yamagata et al. 1993; Resnick et al. 1998). Nevertheless, the enhanced expression after 48 h was mediated enzyme-independently, because both C3 and C3-E174Q induced this effect. The C3-mediated decreased COX-2 abundance after 60 and 72 h is in agreement with previous studies demonstrating that the induction of COX-2 promotor by a constitutively active Gα13-subunit of heteromeric G proteins is blocked in NIH 3T3 cells after transfection with a C3 expression vector for 72 h (Slice et al. 1999). Due to the fact that also the p38 inhibitor skepinone-L slightly reduced COX-2 by 20 %, after 72 h, the p38 signaling seems to play a minor but Sp1-independent role in the regulation of COX-2.
Besides the inhibition of cell proliferation, C3 prevents serum-starved HT22 cells from apoptosis by downregulation of the pro-apoptotic proteins Bax, Bid, p53, and certain caspases at an mRNA and protein level. Moreover, the enzyme activity of caspase-3 and caspase-7 is reduced by C3 treatment for 48 h (Rohrbeck et al. 2012). Among the identified transcriptional factors, especially ATF2 is involved in the transcription of apoptosis-inducing proteins like GADD153, whose increased expression is strongly associated with induction of apoptosis in various cell types (Walton et al. 1998; Bruhat et al. 2000; Maytin et al. 2001; Oh-Hashi et al. 2001; van der Sanden et al. 2004). However, the C3-mediated protection from apoptosis in HT22 cells is in agreement with our findings of a reduced GADD153 abundance. The activation of ATF2 is mediated via phosphorylation by certain kinases such as p38 and JNK (Gupta et al. 1995; Raingeaud et al. 1996). The observed decreased level of phosphorylated p38 supports a connection between ATF2 and p38. The findings are endorsed by a study of Pausawasdi et al. identifying a C3-induced decrease in carbachol-stimulated p38 activity (Pausawasdi et al. 2000). These results imply that the reduced level of phosphorylated p38 may lead downstream to a decreased activity of ATF2. The proposed correlation between p38, ATF2, and GADD153 is further strengthened by the C3-like effects of the p38 inhibitor reducing moderately the level of GADD153 by 20 % after incubation with skepinone-L for 60 and 72 h. In agreement with these findings, prior studies reported that GADD153 transcription is highly associated with p38 in the context of apoptosis induction in various cell types (Oh-Hashi et al. 2001, Wang and Ron 1996). Moreover, GADD153 can also be activated directly by p38 via phosphorylation at serine 78 and 81 (Maytin et al. 2001). Additionally, the missing inhibiting effects of C3-E174Q on phospho-p38 and GADD153 strongly indicate a Rho-dependent reduction of ATF2 activity as a result of the decreased activity of p38 downstream inhibiting the GADD153 abundance (Fig. 6). With regard to the impact of p53 and c-Jun on the C3-mediated anti-apoptotic effect, a prior study in primary hepatocytes demonstrated that c-Jun not only represses the p53 expression via the PF-1 site but also antagonizes p53 activity after apoptosis induction by TNFα (Ginsberg et al. 1990; Schreiber et al. 1999; Eferl et al. 2003). Accordingly, as already mentioned for the C3-induced growth inhibition, also the C3-mediated prevention of apoptosis represents a consequence of the several alterations on the transcriptional and downstream protein level interfering to the anti-apoptotic impact.
In conclusion, we demonstrated that C3-mediated inactivation of Rho-GTPases also influenced transcriptional regulation involved in distinct cellular functions in addition to reorganization of the actin cytoskeleton. We identified a Rho-dependent effect of C3 on transcription factors such as Sp1 and ATF2 and their downstream target genes that were strongly involved in cell proliferation and apoptosis. Thus, these alterations in cell signaling after 48 h result in the C3-mediated anti-proliferative and anti-apoptotic effects.
L.v.E. planned the experiments, performed experiments, analyzed data, and wrote the manuscript. S.H. performed the experiments. I.J. planned the experiments and wrote the manuscript. A.R. planned the experiments, performed experiments, analyzed data, and wrote the manuscript.
Compliance with ethical standards
Competing financial interests’ declaration
The authors declare that they have no conflict of interest.
- Adnane J, Bizouarn FA, Qian Y, et al. (1998) p21(WAF1/CIP1) is upregulated by the geranylgeranyltransferase I inhibitor GGTI-298 through a transforming growth factor beta- and Sp1-responsive element: involvement of the small GTPase rhoA. Mol Cell Biol 18:6962–6970CrossRefPubMedPubMedCentralGoogle Scholar
- Bertrand J, Winton MJ, Rodriguez-Hernandez N, et al. (2005) Application of Rho antagonist to neuronal cell bodies promotes neurite growth in compartmented cultures and regeneration of retinal ganglion cell axons in the optic nerve of adult rats. J Neurosci 25:1113–1121. doi: 10.1523/JNEUROSCI.3931-04.2005 CrossRefPubMedGoogle Scholar
- Bruhat A, Jousse C, Carraro V, et al. (2000) Amino acids control mammalian gene transcription: activating transcription factor 2 is essential for the amino acid responsiveness of the CHOP promoter. Mol Cell Biol 20:7192–7204. doi: 10.1128/MCB.20.19.7192-7204.2000 CrossRefPubMedPubMedCentralGoogle Scholar
- Koeberle SC, Fischer S, Schollmeyer D, et al. (2012a) SI—design, synthesis, and biological evaluation of novel disubstituted dibenzosuberones as highly potent and selective inhibitors of p38 mitogen activated protein kinase. J Med Chem 55:5868–5877. doi: 10.1021/jm300327h CrossRefPubMedGoogle Scholar
- Rivard N, Boucher M-J, Asselin C, L’Allemain G (1999) MAP kinase cascade is required for p27 downregulation and S phase entry in fibroblasts and epithelial cells. Am J Phys 277:C652–C664Google Scholar
- Rohrbeck A, Stahl F, Höltje M, et al (2015a) C3-induced release of neurotrophic factors from Schwann cells—potential mechanism behind its regeneration promoting activity. Neurochem Int 1–14. doi: 10.1016/j.neuint.2015.09.007
- van der Sanden MHM, Meems H, Houweling M, et al. (2004) Induction of CCAAT/enhancer-binding protein (C/EBP)-homologous protein/growth arrest and DNA damage-inducible protein 153 expression during inhibition of phosphatidylcholine synthesis is mediated via activation of a C/EBP-activating transcription factor-r. J Biol Chem 279:52007–52015. doi: 10.1074/jbc.M405577200 CrossRefPubMedGoogle Scholar
Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.