A potential mechanism for fumonisin B1-mediated hepatocarcinogenesis: cyclin D1 stabilization associated with activation of Akt and inhibition of GSK-3ß activity

Danica Ramljak7, Richard J. Calvert1, Paddy W. Wiesenfeld2, Bhalchandra A. Diwan3, Branimir Catipovic4, Walter F.O. Marasas5, Tommie C. Victor6, Lucy M. Anderson and Wentzel C.A. Gelderblom5

Laboratory of Comparative Carcinogenesis, National Cancer Institute, Frederick Cancer Research and Development Center, Building 538, Room 205E, Frederick, MD 21702,
1 Clinical Research and Review Staff and
2 Division of Science and Applied Technology, Office of Special Nutritionals, Center for Food Safety and Applied Nutrition, Food and Drug Administration, Laurel, MD 20708,
3 Intramural Research Support Program, SAIC Frederick, NCI-Frederick Cancer Research and Development Center, Frederick, MD 21702,
4 Asthma and Allergy Center, Johns Hopkins University, Baltimore, MD 21224, USA,
5 Research Institute for Nutritional Diseases, South African Medical Research Council, PO Box 70 and
6 University of Stellenbosch, Medical Research Council Center for Molecular and Cellular Biology, PO Box 19063, Tygerberg 7505, South Africa


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Fumonisin B1 (FB1) is a worldwide corn contaminant and has been epidemiologically linked to the high incidence of human esophageal cancer in South Africa and China. FB1 is hepatocarcinogenic in rats by an unknown mechanism. Inhibition of ceramide synthase and disruption of membrane phospholipids have been shown to be mechanisms of toxicity. Here we show overexpression of cyclin D1 protein in both preneoplastic and neoplastic liver specimens obtained from a long-term feeding study of FB1 in rats. In rats fed FB1 short-term, cyclin D1 protein levels in liver were increased up to five-fold in a dose-responsive manner. Northern blot analysis demonstrated no increase in mRNA levels of cyclin D1. 2D electrophoresis of cyclin D1 protein in FB1-treated samples showed a distinct pattern of migration (presence of less negatively charged form of the protein) that differed from controls. Recently, it has been shown that phosphorylation of cyclin D1 by glycogen synthase kinase 3ß (GSK-3ß) on a single threonine residue (Thr-286) positively regulates proteosomal degradation of cyclin D1. In FB1-treated samples we detected GSK-3ß phosphorylated on serine 9; activated protein kinase B (Akt) appears to be responsible for this activity-inhibiting phosphorylation. These findings suggest that overexpression of cyclin D1 results from stabilization due to a lack of phosphorylation mediated by GSK-3ß. We also observed an increase in cyclin dependent kinase 4 (Cdk4) complexes with cyclin D1 in FB1-treated samples; additionally, elevated Cdk4 activity was shown by increased phosphorylation of the retinoblastoma protein. In summary, the activation of Akt leads to increased survival, inhibition of GSK-3ß activity and post-translational stabilization of cyclin D1, all events responsible for disruption of the cell cycle G1/S restriction point in hepatocytes. This is the first report suggesting the mechanism by which FB1 acts as a carcinogen.

Abbreviations: Akt, protein kinase B; Cdk4, cyclin dependent kinase 4; DEN, N-nitrosodiethylamine; FB1, fumonisin FB1; GSK3ß, glycogen synthase kinase 3ß; HCC, hepatocellular carcinoma; IEF, isoelectric focusing; IHC, immunohistochemistry; MAPK, mitogen-activated protein kinase; PB, phenobarbital; PCNA, proliferating cellular nuclear antigen; PI3K, phosphatidylinositol-3-OH kinase; pRb, retinoblastoma protein; Ser9, serine 9; SSCP, single-strand conformation polymorphism.


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The mycotoxin fumonisin B1 (FB1) is carcinogenic in rat liver, causing primary hepatocellular carcinomas (HCCs) when fed chronically at 50 mg/kg of diet (1). It is produced by the ubiquitous fungus, Fusarium moniliforme, a worldwide contaminant of major grain crops, especially abundant in corn and corn-based products for both animal and human consumption (2). FB1 may be a contributing factor in the development of esophageal cancer in people of South Africa and China (3). It has been shown to be present in corn-based food products in the USA at levels up to 2.7 p.p.m., and in some cases is present together with the hepatocarcinogenic aflatoxin (4).

A mechanism for FB1 toxicity involves disruption of sphingolipid metabolism by inhibition of the enzyme N-acyltransferase (ceramide synthase), responsible for the conversion of sphinganine to sphingosine. This enzymatic inhibition leads to a decrease in cellular ceramide levels (5). In vitro studies in primary hepatocytes (6) and in vivo studies in rat liver (7) suggest that the disruption of phospholipid, cholesterol and fatty acid metabolic pathways might also be important factors in effecting FB1-induced hepatotoxicity. Intracellular effects of FB1 have been studied mainly in non-hepatocyte cell culture and tissue slice models and have included alterations in cell morphology (8), cell–cell interaction (9), mitogen-activated protein kinase (MAPK) activity (10), protein kinase C expression (4) and apoptosis (11). In a cancer target tissue, rat hepatocytes, FB1 was found to inhibit proliferation via G1 arrest and cause apoptosis in the majority of normal hepatocytes, whereas a small minority of hepatocytes survived by escaping from G1 arrest (12). It has been hypothesized that tumor development could be a result of FB1 mimicking genotoxic carcinogens in inducing hepatotoxicity, resulting in compensatory cell proliferation (survival). Clonal outgrowth of initiated cells resistant to the cytotoxic effects of FB1 would then occur amidst the non-proliferative background of surrounding normal cells (13).

Progression through the cell cycle from G1 to S phase is controlled by a restriction point (R) which is limited by the activity of several proteins: the tumor suppressors retinoblastoma (pRb) and p53, the growth factor TGFß, the cyclins D, E and A, their corresponding kinases Cdk2, Cdk4, Cdk5 and Cdk6, and the inhibitors of these cyclin-dependent kinases: the Ink4 family (p15, p16, p18, p19) and Waf/Kip family (p21, p27, p57) (14). The majority of human cancers have been reported to have alterations in the function of one or more of these cell cycle regulatory proteins. Cyclin D1 is a key regulatory protein in the Rb pathway, which controls transition through the restriction point, and is responsible, together with its main catalytic partners, Cdk4/Cdk6, for pRb phosphorylation (15). This process is negatively regulated by the Cdk inhibitors p16INK4, p27KIP1 and p21WAF1. Cyclin D1 is a highly responsive sensor of the growth factor environment of the cell and it is targeted for degradation when mitogenic factors are absent in the cell's milieu (16). When overexpressed due to gene amplification, gene rearrangement, protein stabilization or other mechanisms, cyclin D1 acts as an oncogene by enhancing cell transformation, either alone (17) or in combination with activated ras (18), thereby shortening the G1 phase of the cell cycle. Deregulated function of cyclin D1, often resulting from overexpression of the protein, has been documented in numerous human cancers, including HCC (19,20). Recent evidence indicates that cyclin D1 may also be regulated through the p21 ras/mitogen activated protein kinase (MAPK) pathway (21,22). Interestingly, it has been demonstrated in only a few cell types (e.g. human mammary epithelial cells) that overexpression of cyclin D1 can inhibit cell-cycle progression rather than stimulate growth (23). Cyclin D1 protein expression is regulated during the cell cycle both transcriptionally and post-transcriptionally with differences being cell-type-dependent. Unlike other cyclins (E, A and B) which are regulated by a ubiquitin-dependent proteosomal pathway (24), the regulation of cyclin D1 protein level/activity as a function of cell cycle phase has not yet been clearly defined.

Recently, it was reported that glycogen synthase kinase-3ß (GSK-3ß) phosphorylates cyclin D1 on Thr-286, thereby triggering rapid cyclin D1 turnover (25). It was also shown that a mutant cyclin D1 protein refractory to phosphorylation by GSK-3ß remained present in the nucleus throughout the entire cell cycle. GSK-3ß is a ubiquitously expressed protein-serine/threonine kinase whose activity is inhibited upon phosphorylation of serine 9 (Ser9) by Akt (protein kinase B) (26). Akt is a part of the anti-apoptotic phosphatidylinositol 3 kinase (PI3K)/Akt cell survival pathway (27). It can be activated by different stimuli involved in cellular survival (such as insulin, growth factors, cytokines) or inhibited by the pro-apoptotic lipid molecule ceramide (28,29). Akt is activated by phosphorylation predominantly at two regulatory sites (Thr308 and Ser473) (30). The mechanism by which ceramide inhibits Akt phosphorylation remains unclear.

Although several mechanisms for FB1-associated liver toxicity have been proposed, very little supportive data exist to explain its carcinogenic effects. This investigation was undertaken to determine whether the overexpression of cyclin D1, which disrupts the G1 checkpoint in hepatocytes, may be an early event and a part of the mechanism(s) responsible for FB1-induced hepatocarcinogenesis. Our data indicate that activation of Akt and consequent inhibition of GSK-3ß activity could be responsible for the detected overexpression and post-translational modification of cyclin D1. These molecular events may play a major role in FB1-induced carcinogenesis and provide possible links between the toxic and carcinogenic effects of FB1.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Short-term FB1 study
F344/NCr rats were fed 0, 50, 100 or 250 mg FB1/kg diet over a period of 21 days using a purified (AIN 76) diet as described previously (31). Half of each liver sample was kept at –80°C prior to analyses, while the other half was paraffin-embedded. A total of 14 paraffin-embedded livers were analyzed by immunohistochemistry (IHC) (nine FB1-treated and five controls), whereas a total of eight liver protein lysates were analyzed by both western analysis and immunoprecipitation (six FB1-treated and two controls) and an additional nine liver protein samples were lysed in kinase buffer (six FB1-treated and three controls) for GSK-3ß and Akt kinase activity assays.

Long-term carcinogenic studies
Tumors were generated by chronically feeding FB1 (50 mg/kg of diet) to BD IX rats (1) and nitroglycerin (32) or N-nitrosodiethylamine (DEN)/phenobarbital (PB) to F344/NCr rats (33). A total of 15 paraffin-embedded FB1-treated rat hepatic specimens (10 carcinomas, three adenomas and two livers with preneoplastic foci), seven control livers and 12 HCCs induced by the other two carcinogens were analyzed.

Genomic DNA, RNA and protein isolation
DNA was recovered from seven paraffin-embedded controls and 15 liver specimens from FB1 long-term treated rats as described previously (34), with the exception that only a single wash with acetone was used following octane extraction. One-half of each of the frozen livers from the short-term FB1-feeding study was used for RNA analysis following RNA extraction with TRIZOL (Gibco BRL, Gaithersburg, MD), and protein samples were prepared from the other half of the same livers. For protein analysis (western blot analysis, immunoprecipitations and kinase assays), tissues were frozen in liquid nitrogen. Powders of frozen tissue samples for analysis by both western blot and immunoprecipitation were dissolved in lysis buffer (10 mM Tris pH 7.4, 150 mM NaCl, 5 mM EDTA, 10% glycerol, 1% Triton X-100, 1 mM phenylmethylsulfonyl fluoride, 0.25 TIU/ml aprotinin and 1 mM sodium orthovanadate). Liver tissue samples used for both GSK-3ß and Akt kinase analysis were lysed in a kinase buffer (50 mM HEPES pH 8.0, 150 mM NaCl, 2.5 mM EGTA, 1 mM EDTA, 0.1% Tween-20 and protease and phosphatase inhibitors as described above). The Bio-Rad (Richmond, CA) protein assay was used for protein quantification.

K-ras and H-ras mutation analysis
Genomic DNA, from liver specimens from the long-term study with FB1, was examined for mutations in the K-ras and H-ras genes (exons 1 and 2) by single-strand conformation polymorphism (SSCP) analysis, restriction fragment length polymorphism analysis and sequencing, as described previously (32,35). The PCR conditions for H-ras exon 1 were 92, 58 and 72°C for 30 s each (30–35 cycles) and for H-ras exon 2 were 95, 53 and 72°C for 30 s each (30–35 cycles). The exon 1 primers were: 5'-GCA ACC CCT GTA GAA GC-3' and 5'-TCA TAC TCG TCC ACA AAA TG-3'. The exon 2 primers were: 5'-CCC TTA AGC TGT GTT CTT TTG-3' and 5'-CTG TGC GCA TGT ACT GGT-3'. The sequencing primer for H-ras exon 2 was 5'-CAG GTA GTC ATT GAT GGG GA-3'. `Cold SSCP' analysis for H-ras exon 1 mutation detection was conducted as described (36). The denaturant methyl mercury hydroxide (MeHgOH) was omitted for exon 2 analysis, and additional PCR primers were included (2 µl of 2 µM stock per final 20 µl reaction volume) to allow visualization of mutant mobility shifts caused by primer–single-strand PCR product heteroduplexes. The mixture was heated to 95°C, then the primers were allowed to anneal to the single strands at room temperature for 5 min prior to gel loading. Gels were electrophoresed at 300 V for 3.5 h at 8°C (buffer temperature). Positive mutant controls were used to optimize conditions for mutation detection. In all cases, confirmation of SSCP analyses was made by repeating PCR from the original template for PCR products with mobility shifts on initial SSCP. Mutations were identified by cycle sequencing using {alpha}-32P-labeled dideoxynucleotides (Thermo Sequenase Kit; Amersham Pharmacia Biotech, Cleveland, OH).

Immunohistochemical staining
Paraffin-embedded liver sections (5 µm) from both short-term (nine FB1-treated and five control livers) and long-term FB1-treated rats (10 carcinomas, three adenomas and two livers with neoplastic nodules), and from seven normal livers (corresponding control rats), were used for immunohistochemical analysis. In addition, kidney, prostate, heart, thyroid, intestine, salivary gland, adrenal gland, lung, brain, spleen and testes from the same animals treated with FB1 and from the corresponding control animals were examined. A total of 12 HCCs induced by two other carcinogens (nitroglycerin and DEN/phenobarbital) were also included in this IHC analysis. Freshly cut tissue sections were microwaved twice in 10 mM sodium citrate (pH 6.0) for 5 min to expose the antigen. Goat serum was used to suppress non-specific binding. Tissue sections were incubated at 4°C overnight with a 1:1000 dilution of rabbit polyclonal anti-human cyclin D antibody which recognizes only form [a] of cyclin D1 and has some cross-reactivity with cyclin D2 (Upstate Biotechnology, Lake Placid, NY) (37,38). The sections were then washed in buffer and incubated with biotinylated anti-rabbit secondary antibody for 30 min. Sections for PCNA staining were microwaved in water, blocked with horse serum and incubated with a 1:1600 dilution of mouse monoclonal anti-PCNA antibody (clone PC10, Dako Corp., Carpinteria, CA), followed by anti-mouse IgG. Diaminobenzidine from the Vectastain Elite ABC kit (Vector Laboratories, Burlingame, CA) was used for final detection. As a staining control, primary antibodies were omitted on one slide from each staining series. All the liver tissues were evaluated by a blinded observer. Nuclear staining was graded as follows: zero to very few cells staining (0), weak (1), moderately positive (2) and very strongly positive (3) compared with normal tissues. The tissues were also evaluated for cyclin D1 cytoplasmic staining.

Western blot
An aliquot of 100 µg of total liver protein was used for each analysis. The samples were run on 12% SDS–PAGE gels for analysis of cyclin D, Cdk2 and cyclin E. The samples for pRb analysis were run on 8% SDS–PAGE gels. Proteins were electroblotted to Immobilon-P PVDF membranes (Millipore Corp., Bedford, MA). The membranes were then stained with Ponceau protein stain (Sigma, St Louis, MO) to confirm equal sample loading. Primary antibodies used were: anti-human cyclin D rabbit polyclonal IgG (UBI), rabbit polyclonal cyclin E (M-20), Cdk4 (C-22; Santa Cruz Biotechnology, Santa Cruz, CA), Cdk2 (UBI) and anti-human pRb (PMG3-245) mouse monoclonal (kind gift of Dr Wen-Hwa Lee, University of Texas, San Antonio, TX). The positive control used for cyclin D1 western blot analysis was EGF-stimulated A431 cell lysate (UBI). Membranes first probed for cyclin D were subsequently stripped and probed for Cdk4 (C-22). The secondary antibodies used were horseradish peroxidase-labeled anti-mouse or anti-rabbit antibodies (Amersham Co., Arlington Heights, IL), with an enhanced chemiluminescence detection kit and X-ray film (Amersham) used for detection.

Isoelectric focusing (IEF) and 2D electrophoresis
IEF was performed by using IEF gels pH 3–10 (pI performance range of 3.5–8.5; Novex, San Diego, CA). A total of 100 µg of protein lysate, as used for cyclin D1 expression analysis by western blotting, from all control and FB1-treated livers (short-term experiment) was mixed (1:2) with IEF sample buffer (Novex) and loaded on the gels. Gels were run according to the manufacturer's instructions. Before running in the second dimension, gels were stained with Coomassie blue dye and later destained. IEF gel slices were cut and loaded on 12% 2D gels (Novex) in such a way that in both control and experimental samples the upper part of the IEF gel (more basic pH) was positioned toward the left part of the 2D gel next to the well for the marker. The more acidic part of the slice was positioned toward the right side. The protein marker used was prestained Kaleidoscope marker (Bio-Rad, Richmond, CA). Samples were run in a way that the gel with one FB1-treated protein sample was always parallel to the gel with one control sample as a sandwich in the gel box (Novex) and gels were run according to instructions provided by the manufacturer (Novex). After electroblotting, membranes (Immobilon-P) were blocked with 2% BSA, washed three times with PBST, and probed with a monoclonal antibody (clone DCS-6; Neomarkers, Fremont, CA) that recognizes both forms [a] and [b] of cyclin D1 (38). The secondary antibody used was horseradish peroxidase-labeled anti-mouse (Amersham). Protein detection was performed using an enhanced chemiluminescence kit and X-ray film (Amersham).

Determination of GSK-3ß and Akt kinase activity
Western blots of 25 µg of total protein were performed as described above. The samples were run on 8% SDS–PAGE gels (Novex) for Akt, and 10% SDS–PAGE gels for GSK-3ß. For Akt analysis we used phospho- and non-phospho cell extracts from NIH 3T3 cells prepared following PDGF treatment at 50 ng/ml for 20 min (New England Biolabs, Beverly, MA); these lysates were loaded on Akt gels and served as both positive and negative controls. In order to determine the expression and phosphorylation status/activity of Akt, membranes were probed with a primary antibody recognizing Akt independent of its phosphorylation status, and an antibody which detects Thr308 phosphorylated Akt protein (one of the residues of Akt targeted by activating phosphorylation). Both Akt antibodies were purchased from New England Biolabs. For detection of GSK-3ß protein levels, mouse monoclonal antibody (0011-A) was used (Santa Cruz). The antibody which specifically recognizes the serine 9 phosphorylated form of GSK-3ß, which is the site at which Akt phosphorylation inactivates GSK-3ß, was used to detect the phosphorylated GSK-3ß protein (BioSource International, Camarillo, CA). The secondary antibodies used were horseradish peroxidase-labeled anti-mouse or anti-rabbit IgG (Amersham). Protein detection was performed using an enhanced chemiluminescence kit and X-ray film (Amersham).

Immunoprecipitation and immunoblotting analysis of cyclin D–Cdk4 complexes
An aliquot of 5 µg of anti-cyclin D (UBI) was incubated with 500 µg of total liver protein lysate at 4°C overnight. The immune complexes were then captured with 50 µl of packed protein A–agarose beads (Boehringer Mannheim, Indianapolis, IN) while rocking at 4°C for 2 h. After three washes with cold lysis buffer, the beads were resuspended in 20 µl of 2x Laemmli sample buffer. Samples were run on 12% SDS–PAGE gels and transferred to membranes, which were blocked with 2% BSA and probed with primary antibody for Cdk4 (C-22), followed by secondary anti-rabbit polyclonal antibody (Amersham). Detection was performed as described above.

Cdk4 kinase assay
A total of 500 µg of total protein was immunoprecipitated with 2 µg of Cdk4 antibody (C-22) and kinase activity assay was performed as described previously (39). The plasmid for the fusion protein GST-Rb (amino acids 379–928) was the generous gift of Dr Mark E.Ewan (Dana Farber Cancer Institute, Boston, MA). Labeled products were separated on denaturing 8% polyacrylamide gels (Novex). Gels were fixed in 15% methanol/10% acetic acid, dried and bands with phosphorylated GST-pRb were visualized by autoradiography. In addition, western blot analysis was performed to confirm Cdk4 protein levels using 20 µl of the two controls and two samples from each animal treated with 250 p.p.m. of FB1 from the same aliquots used to assess Cdk4 kinase activity.

Northern blot analysis
An aliquot of 20 µg of total RNA was electrophoresed on a 1% agarose/formaldehyde gel and transferred to a Gene Screen Plus nylon membrane (Du Pont NEN Research Products, Boston, MA). A 1.3 kb fragment of the mouse cyclin D1 cDNA (the kind gift of Dr Charles J.Sherr, St Jude Children's Research Hospital, Memphis, TN), labeled by random priming with [{alpha}-32P]dATP (3000 Ci/mmol) (Lofstrand Labs Limited, Gaithersburg, MD), was used to probe the membranes. After stripping, the membranes were probed with a rat 18S ribosomal RNA probe (DNA oligomer, RAT 18SHP) (40). The intensity of the bands detected by both northern and western blot analysis was quantified using a Molecular Dynamics enhanced-laser densitometer.


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 Abstract
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 Materials and methods
 Results
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 References
 
Early cyclin D1 overexpression caused by short-term exposure to FB1
We hypothesized that FB1 acts as a carcinogen by disrupting the G1/S checkpoint in rat hepatocytes resistant to its mitoinhibitory effects, due to an overexpression of cyclin D1 protein. To test whether cyclin D1 overexpression is an early event during FB1 hepatocarcinogenesis, rats were fed FB1 doses for 21 days. Half of each liver was frozen for immunoblot analysis, while the other half was paraffin-embedded for IHC. Cyclin D1 was found to be overexpressed in FB1-treated livers, as detected by IHC, compared with control livers where little staining was observed (Figure 1Go). About two-thirds of all hepatocyte nuclei in treated rats showed some staining for cyclin D1, whereas ~5–10% were intensively stained. Consecutive sections from the same blocks were stained for PCNA, a marker for proliferating cells. In control liver sections, only a few cells were stained, whereas in experimental liver sections the number of PCNA positive cells exceeded those positive for cyclin D1 (data not shown). In addition, we confirmed the overexpression of cyclin D1 by western analysis, detecting an overexpression of ~36 kDa cyclin D1 protein in a FB1 dose-dependent manner (up to 5-fold) in experimental total protein lysates compared with control (Figure 2Go). Furthermore, in western analysis a 34 kDa protein was detected and was considered to be cyclin D2, since the antibody used for detection of cyclin D1 cross reacts with cyclin D2 and this molecular weight has been reported previously for cyclin D2 (37). Cyclin D2 levels were somewhat higher in most FB1 samples compared with non-treated controls, but did not show a dose–response.



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Fig. 1. Immunohistochemical staining for cyclin D1 in control and FB1-treated livers from short-term exposure study. In control livers only a few cells were stained (A), whereas in FB1-treated (100 p.p.m.) livers many nuclei were stained (B).

 


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Fig. 2. Western immunoblot analysis for cyclins D1, D2 and E, Cdk4, Cdk2 and pRb in livers from control rats and rats fed FB1 for 21 days. The loading sequence for all gels was, in each case (right to left): two controls, two at 50 p.p.m., two at 100 p.p.m. and two at 250 p.p.m. FB1 (see label at bottom of figure). The positions of relevant molecular weight markers, in kDa, are indicated at the right and the positions of bands of interest with arrows at the left. These results are representative of three identical, independent assays using the same lysates.

 
Other proteins of the pRb pathway (Cdk4, pRb, cyclin E and Cdk2) were analyzed by immunoblot (Figure 2Go). Levels of Cdk4 were not altered by FB1 treatment. The pRb antibody recognized several bands ranging from ~105 to ~115 kDa. pRb was present in a slower migrating (hyperphosphorylated) form in some of the livers from animals treated with high doses of FB1 (100 or 250 p.p.m.); it is noteworthy that these were livers in which levels of cyclin D1 were especially high. Levels of cyclin E and Cdk2 were not changed between control and FB1-treated livers with the exception of a single sample (250 mg/kg of diet) in which a decrease in both proteins was detected.

Cyclin D1 protein stability as a post-translational event
Northern blot analysis was performed in order to test whether the levels of the mRNA were increased in the FB1-treated samples in which the overexpression of cyclin D1 protein was detected. Both RNA and protein samples were taken from the same liver lobe. An ~4 kb cyclin D1 transcript was detected in all samples with no consistent change in cyclin D1 mRNA expression between control and experimental samples (Figure 3Go). To test whether the overexpression of cyclin D1 protein is due to post-translational events, the same protein lysates in which its stabilization was detected (Figure 2Go) were analyzed by IEF and 2D electrophoresis followed by probing of membranes with a monoclonal antibody for cyclin D1 (DCS-6). Although this antibody is directed against human cyclin D1, the epitope of the DCS-6 antibody is at the C-terminal portion of the cyclin box, an area with 100% conservation between rat and human cyclin D1 (41). 2D gels from control samples showed two distinct closely-running spots in the middle of the membranes at ~36 kDa, corresponding to the expected size for the cyclin D1 protein (Figure 4Go, left panel). These represent the two forms of the cyclin D1 [a] and [b] recently reported to be detected by the DCS-6 antibody (Neomarkers) (38). This monoclonal antibody does not possess cross-reactivity to cyclin D2. On membranes from gels for all six protein samples from FB1-treated livers (representative blot, Figure 4Go, right panel), we detected one form of cyclin D1 in the same position as the lower spot in the control membranes. In the FB1-treated samples, the upper spot (detected in controls) was not seen; instead, an additional spot lying toward the right portion of the membrane was detected. Clearly, in the experimental samples, the upper form [a] of cyclin D1 became less negatively charged and was shifted to the right. In contrast, two spots were also found for cyclin E, but these did not vary in position in control compared with FB1-treated livers (data not shown). Also, 2D electrophoresis of the same samples did not reveal any differences in the migration pattern of the Cdk4 protein (data not shown), thus confirming the specific effects of FB1 treatment on post-translational modification of cyclin D1.



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Fig. 3. Northern blot analysis for cyclin D1 mRNA from FB1-treated and control livers. The blot was probed with a 1.3 kb mouse cyclin D1 cDNA probe, then stripped and probed with a rat cDNA probe for 18S ribosomal RNA as an internal control for RNA loading. Data are representative of two identical assays.

 


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Fig. 4. Representative 2D gel electrophoresis of cyclin D1. One-hundred micrograms of the same protein lysates used for determination of cyclin D1 expression levels (Figure 2Go) were separated via IEF on gels run according to the manufacturer's instructions (Novex). Gel slices containing the separated proteins were positioned for running in the second dimension, with the top (containing the more negatively charged, rapidly migrating proteins in the first separation) positioned next to the protein marker (left of each panel). After the second electrophoresis, the proteins were electroblotted to Immobilon-P membranes and probed with a monoclonal antibody to cyclin D1. Data are representative of two identical assays.

 
Inhibition of GSK-3ß by activated Akt in FB1-treated livers
Recently, GSK-3ß has been shown to be a specific kinase that phosphorylates cyclin D1 on Thr-286, thus triggering its rapid degradation (25). Upon phosphorylation by activated Akt on Ser9, GSK-3ß kinase activity is inhibited (26). In order to test if the detected post-translational modification of cyclin D1 documented by the presence of a less negatively charged form of cyclin D1 (possibly a form with fewer phosphorous groups) in FB1-treated samples is linked to inhibition of GSK-3ß, the same liver specimens were analyzed for both the expression level and phosphorylation status of GSK-3ß. We detected ~50 kDa GSK-3ß protein expressed in both control and FB1-treated samples to an equal extent (Figure 5AGo). However, in FB1-treated samples, the protein migrated more slowly compared with control samples. In addition, more phosphorylated GSK-3ß protein was detected (~2-fold increase in rats fed 50 p.p.m. of FB1; ~1.5-fold increase in rats fed 100 p.p.m.), indicative of its possible inactivation (Figure 5BGo). However, we did not detect a dose–response-related phosphorylation of GSK-3ß (Figure 5CGo), possibly due to the limitations of the analysis (it is difficult to evaluate the linearity of kinase activation by using this approach and further complication arises when using protein from tissue samples). Because Akt has been shown to inhibit GSK-3ß by phosphorylation, we predicted that FB1 would activate Akt, possibly due to its toxic action on both sphingolipids and phospholipids. This could lead to alterations of the signaling molecules involved in the control of Akt activation (29). Because GSK-3ß is a target of activated Akt, we analyzed the same lysates for both expression and phosphorylation status of Akt. Akt protein was detected at ~60 kDa and total levels of Akt were similar in both FB1-treated livers and controls (Figure 6AGo); however, in FB1-treated livers, a slower migrating Akt was detected. Furthermore, more phosphorylated Akt (on Thr 308) was detected in FB1-treated samples (4-fold increase in rats fed 50 p.p.m. and 7-fold increase in rats fed 100 p.p.m.) (Figure 6BGo), indicative of dose-dependent activation of Akt kinase.




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Fig. 5. Western immunoblot analysis for expression levels and activity of total GSK-3ß in livers from control rats and rats fed FB1 (50 p.p.m. and 100 p.p.m.) for 21 days. The band at ~50 kDa represents total GSK-3ß detected using an antibody recognizing GSK-3ß protein independent of its phosphorylation status (A). More highly phosphorylated (inactivated by phosphorylation on Ser9) GSK-3ß protein was detected in FB1-treated samples (B). In rats fed 50 p.p.m. of FB1 the increase in phosphorylated GSK-3ß was ~2-fold, whereas in rats fed 100 p.p.m. of FB1, the increase was only 1.5-fold (C). Data are representative of two identical assays. R.S.U., relative scan units.

 


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Fig. 6. Western immunoblot analysis for expression and phosphorylation status of Akt in livers from controls and FB1-treated rats. (A) Akt protein was detected at ~60 kDa in controls,whereas a slower migrating protein form was detected in FB1-treated livers. (B) More highly phosphorylated Akt (Thr 308; one of the Akt activation sites in vivo) was detected in livers from rats treated with 50 p.p.m. (4-fold) and 100 p.p.m. (7-fold) FB1 as compared with controls. Data are representative of two identical assays.

 
Increased cyclin D–Cdk4 complex formation and Cdk4 activity in FB1-treated livers
Cyclin D-containing complexes were immunoprecipitated and immunoblots were then probed with an antibody against Cdk4. Cyclin D–Cdk4 complexes were more prominent in the FB1-treated samples than in control samples (Figure 7AGo). This increase was due to overexpression of cyclin D1, because the total levels of the Cdk4 were not changed in the FB1-treated livers (Figure 2Go). To investigate whether the increased levels of cyclin D1 upregulated Cdk4 activity, we performed an in vitro kinase assay for Cdk4 activity using recombinant GST-pRb fusion protein as the kinase substrate. Cdk4 from FB1-treated livers had much higher kinase activity compared with Cdk4 in control livers where no visible GST-pRb phosphorylation could be detected (Figure 7BGo), in spite of the presence of small amounts of cyclin D/Cdk4 complexes and equal levels of Cdk4 protein in both control and experimental samples (Figures 2 and 7CGoGo). The reason why Cdk4 activity does not parallel the increase in cyclin D1 dose-dependent overexpression is not clear. It is possible that linearity of the assay was exceeded. In other words, the assay is in the area of the curve where it is impossible to see a dose–response. The absence of a detectable level of kinase activity in control samples could be predicted, since it is likely that the activation of cyclin D1/Cdk4 complexes is very critically restricted in normal, quiescent adult rat hepatocytes.



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Fig. 7. Complex formation, activity and amounts of Cdk4. (A) Cyclin D1 was immunoprecipitated from the protein lysate, separated by gel electrophoresis, blotted to membrane and the membrane was probed with an antibody to Cdk4. (B) Cdk4 kinase activity in the liver lysates was measured by immunoprecipitating Cdk4 from the lysates, incubating with fusion protein (GST-pRb) in the presence of 32P, separating the products in denaturing gels, and detecting phosphorylated Rb by autoradiography. (C) The Cdk4 immunoprecipitates used in the kinase activity assay were additionally analyzed by western blot and equal levels of Cdk4 expression in two control and high-dose FB1 samples were observed. All data represent two identical assays.

 
Overexpression of cyclin D1 protein in FB1-induced tumors
Immunohistochemical staining was performed in order to analyze the level of cyclin D1 expression in tumors from a long-term FB1-feeding study and to determine both the liver cell type overexpressing cyclin D1 as well as its subcellular localization. Staining in normal livers was limited to the nuclei of a few hepatocytes (Figure 8AGo), while an increase in the number of cells with weak nuclear staining (grade 1) was seen in preneoplastic foci which previously stained positive for {gamma}-glutamyltranspeptidase (Figure 8BGo) (1). All benign tumors showed evidence of grade 2 and all HCCs showed strong grade 3 nuclear staining for cyclin D (Figure 8C and DGo). The intensity of the staining was correlated with the grade of the lesion. These findings suggest that cyclin D1 deregulation is involved in development of early lesions as well as in malignant conversion. Highly cirrhotic and necrotic areas within the HCCs and focal aggregates of lymphocytes, common within the connective tissue septa, were both found to be negative for cyclin D1. Cyclin D1 overexpression was detected in the epithelial cells of the bile ducts in two tumors with lesions typical of cholangiofibrosis (data not shown).



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Fig. 8. Immunohistochemical staining for cyclin D1 in control and FB1-exposed rat livers. (A) In control livers, only occasional nuclei (arrow) show light staining for cyclin D1. (B) A preneoplastic focus in the center of the picture shows increased numbers of nuclei with moderate cyclin D1 staining (arrow). (C) A liver adenoma with most nuclei stained for cyclin D1. (D) In HCCs, the majority of the nuclei stain with greater intensity (dark brown) than liver adenomas.

 
Absence of cyclin D1 overexpression in other tissues
Alteration in cyclin D1 expression was a liver-specific finding as all other tissues examined lacked cyclin D1 overexpression with the possible exception of the kidney (data not shown). Kidney showed some positive nuclear staining in the proximal tubules in both untreated and treated rats. This finding must be interpreted with caution due to the tendency of the proximal tubules to stain non-specifically in IHC. Because chronic interstitial nephritis was present in the kidneys and FB1 has been shown to have toxic effects in rat kidneys (42), this finding may also reflect an actual role of cyclin D1 in this pathology.

Specificity of the FB1 effect on cyclin D1 in hepatocytes
To test whether the overexpression of cyclin D1 was a common property of rat HCCs induced by other carcinogens, liver sections from paraffin-embedded rat HCC caused by nitroglycerin or DEN/PB (32,33) were compared with those induced by FB1. Tissues from all three studies were processed identically. The cyclin D1 overexpression, which was characteristic of FB1-induced HCC (Figure 9AGo), was not seen in HCC caused by either DEN/PB (Figure 9BGo) or nitroglycerin (Figure 9CGo).



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Fig. 9. Comparison of cyclin D1 staining in tumors from FB1-treated livers versus tumors from other treatments (includes analysis of levels of PCNA in these livers). (A) Strong nuclear staining is apparent in a FB1-caused rat liver tumor, whereas staining was absent from tumors caused by DEN/PB (B) and only a few nuclei (arrow) stained in carcinomas induced by nitroglycerin (C). The intensity of staining for PCNA in the HCCs induced by the three carcinogens: FB1 (D), DEN/PB (E) and nitroglycerin (F) is very similar.

 
To test whether increased cyclin D1 expression was merely a reflection of the rate of cell proliferation, the tissues were stained for PCNA and cyclin D1 in consecutive sections for all experimental and control livers. In control livers, staining for both antigens was detected in only a few cells (graded as 0). In preneoplastic foci in the FB1-treated livers, the PCNA-positive cells exceeded the number of those positively staining for cyclin D1 by ~2-fold; in benign and malignant FB1-induced tumors, the same areas stained positively for both antigens, with cyclin D1 slightly exceeding the PCNA-positive staining cells (data not shown). HCC induced by nitroglycerin or DEN/PB showed PCNA staining rates similar to those in the FB1 tumors (Figure 9D–FGo). Thus, although a high rate of cell proliferation is a common property of the HCCs induced in each of the three different induction models, only FB1-induced HCCs exhibited overexpression of cyclin D1.

Absence of K-ras and H-ras activating mutations
To test whether mutations in ras oncogenes could be driving the cyclin D1 protein overexpression, we analyzed all experimental and control samples for K-ras and H-ras mutations. Only two samples with third position A->T transversions were found in codon 61 of exon 2 of H-ras (data not shown), thus ruling out activating mutations in ras as a major contributor to the observed cyclin D1 overexpression.


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
To date, fumonisin B1 hepatocarcinogenicity in rats occurs by an unknown mechanism. The present study reveals that FB1 treatment results in cyclin D1 overexpression via protein stabilization due to post-translational modification(s) without effect on cyclin D1 mRNA levels. In addition to cyclin D1 stabilization, we detected Akt activation resulting in phosphorylation and inhibition of GSK-3ß. These findings indicate that the activation of a major survival molecule (Akt) and its downstream effectors leads to alteration of cell cycle progression in hepatocytes; therefore, this could prove to be a mechanism or part of a mechanism responsible for FB1 carcinogenesis.

Our 2D gels electrophoresis patterns indicated that a less negatively charged form of cyclin D1 predominated in FB1-treated livers as compared with control livers from a short-term FB1 feeding study where both forms detected were of similar charge. It is likely that the two spots detected were [a] and [b] forms of cyclin D1, as reported by Sawa et al. (38). Cyclin D1 form [a] associates in a ternary complex with PCNA, WAF-1 and Cdk4. It has been known for some time that cyclin D1 contains a C-terminal PEST sequence (24). Removal of the PEST sequence from yeast G1 cyclins prolongs their half-life (43). Cyclin D1 form [a], not form [b], contains a PEST-rich region which is probably responsible for its rapid turnover (38). One interpretation of our findings is that the cyclin D1 spot, which has an altered position by 2D gels electrophoresis after FB1 treatment, is the form [a] which recently has been implicated in controlling cell cycle entry (38). It appears that FB1 treatment causes post-translational modification of cyclin D1 form [a] in such a way as to make it less negatively charged (possibly less phosphorylated), thus leading to an increase in its stability. Efforts to identify the exact nature of this post-translational change are currently in progress. Also, additional confirmation of the involvement of form [a] is derived by detection of overexpressed cyclin D1 in FB1-treated samples by western blotting (Figure 2Go) using an anti-cyclin D1 polyclonal Ab (UBI) reported to recognize only form [a] of cyclin D1 (38). We used the same antibody (UBI) in the analysis of binding between cyclin D1 and Cdk4; therefore, it appears that the [a] form of cyclin D1 is responsible for cyclin D1/Cdk4 binding in our studies and is responsible for the pRb phosphorylation we detected.

A lack of correlation between cyclin D1 protein and mRNA expression has been reported in some human cell lines and tumors (20,44). There is frequently a correlation between cyclin D1 gene amplification and cyclin D1 protein overexpression in human hepatic, esophageal and head and neck cancer, whereas in breast cancer, amplification is detected in only 13% of the tumors despite evidence that >50% exhibit cyclin D1 protein overexpression. Similar findings have been reported in sarcomas, colon cancers and melanomas (20), further implicating post-translational modification of cyclin D1 as a mechanism for overexpression.

Recently, cyclin D1 has been shown to be regulated post-transcriptionally by GSK-3ß (25,45), calpain (46) and retinoic acid (47). GSK-3ß has been shown to phosphorylate cyclin D1 specifically on Thr-286, thus triggering rapid turnover of the protein. Defective GSK-3ß kinase activity could be responsible for the high stability of cyclin D1 (which remains in the nucleus throughout the cell cycle) (25). Because the activity of GSK-3ß can be inhibited by signalling through a pathway that involves Ras/PI3K/Akt (25) (a major survival pathway in cells), the involvement of some of these molecules in the regulation of GSK-3ß activity and cyclin D1 stability was analyzed. We detected more phosphorylated and activated Akt in FB1-treated livers as compared with control livers, and have reasoned that this alteration is responsible for the inhibition of GSK-3ß as well as the less negatively charged state of the [a] form of cyclin D1 (possibly less phosphorylated).

Also, because cyclin D1 protein overexpression has been linked to both the ras/MAPK (21,22) and PI3K pathways (25), and K-ras is frequently mutated in rat liver tumors (48), we tested our samples for mutations in ras oncogene. Only two mutations in codon 61 of exon 2 of the H-ras oncogene were found; therefore, a potential role for ras-activating mutations in the overexpression of cyclin D1 in our FB1 experimental samples is very limited. This is in agreement with previous reports involving human esophageal cell lines and primary esophageal cancers where cyclin D1 overexpression is detected frequently (49,50), whereas activating mutations in ras oncogenes are rare to non-existent (51).

Recent evidence suggests that cyclin D1 expression from a heterologous promoter can lead to apoptosis in serum-starved rat fibroblasts; potentially, this outcome may involve phosphorylation of pRb (52). FB1 has been reported to be capable of inducing apoptosis in rat liver cells (11). It appears that the absolute level of cyclin D1 expression can determine whether cells will undergo transformation (moderate overexpression) or apoptosis (high overexpression).

In the majority of cell types, activated Akt has been shown to inhibit apoptosis through inhibition of pro-apoptotic molecules: Bad, caspase 9, caspase 3 and Fas/CD95 (53,54). Activated Akt detected in FB1-treated liver could potentially be responsible for the inhibition of apoptosis in those hepatocytes that are resistant to FB1 toxicity (those that probably give rise to tumor development), whereas the majority of hepatocytes are sensitive to FB1-induced apoptosis due to its toxicity. Although it is not completely clear which molecules are participating in its activation, Akt has been reported to be negatively regulated by the tumor suppressor gene PTEN (55), and the lipid signalling molecule ceramide (29). FB1 has been shown to inhibit ceramide synthase both in vitro and in vivo, therefore leading to a decrease in cellular ceramide levels and an increase in intracellular sphinganine (5). These events could potentially contribute to the detected activation of Akt in our study. Additionally, due to the effects of FB1 on total phospholipids and fatty acids in hepatocytes (6,7), it is possible that FB1 is altering molecules involved in the activation/deactivation of Akt (PI3K, PDK1, PDK2 or tumor suppressor gene PTEN) (53,55).

The ability of FB1 to cause a relatively early dose-dependent increase in cyclin D1 protein expression was shown by both IHC and immunoblot analysis of livers from rats exposed to FB1 for only 3 weeks. Although levels of Cdk4 protein were not increased by FB1 treatment, the amount of cyclin D1–Cdk4 complex formed as well as Cdk4 kinase activity were greatly elevated as shown by increased phosphorylation of a GST-pRb substrate. These findings are in agreement with previous reports in which the expression level of cyclin D1, regardless of Cdk4 level, is rate limiting for complex formation and is responsible for Cdk4 kinase activity (49). In addition, more hyperphosphorylated forms of pRb were detected in vivo in some samples with high cyclin D1 levels. The key role of cyclin D1 was further underscored by lack of alteration in the level of proteins involved in control of the late G1 phase of the cell cycle (cyclin E and Cdk2). Although cyclin E is sometimes overexpressed in rodent and human tumors (56,57), the overexpression of cyclin D1 and consequent hyperphosphorylation of pRb occur more frequently (20).

The effects of FB1 on cyclin D1 during short term exposure were further substantiated by results from FB1 chronic treatment: clear association of its nuclear overexpression with liver neoplasia (especially in progression to carcinoma). This was evidenced by immunohistochemical analysis of pre-neoplastic foci, adenomas and carcinomas in the livers of rats chronically fed FB1 (50 mg/kg diet) over a period of 20–26 months (1). The overexpression of cyclin D1 was specific to the liver as a targeted organ, with few to no cells stained in the control livers nor in the 12 other tissues analyzed from the same rats (with the possible exception of kidney which is also a target for the toxic effects of FB1). In the majority of the experimental rats, proximal tubules of the kidney stained intensively for cyclin D1. However, it is difficult to interpret this finding, because in some of the control rats, we detected a similar staining, although less intense.

Notably, rat HCCs induced by two other carcinogens failed to show overexpression of the protein when stained for cyclin D1 in parallel with the FB1 tumors despite similar cell proliferation as indicated by PCNA staining. Thus, the effect on cyclin D1 was unique to FB1. There are a few chemical carcinogens reported to have an effect on cyclin D1 overexpression in rodent tumors similar to that of FB1: in mouse skin tumors, dimethylbenz[a]anthracene was used as an initiator followed by 12-O-tetradecanoylphorbol-13-acetate as the promoter (56,58) and N-methyl-N-nitrosourea in rat mammary carcinomas (59). However, to our knowledge, there are no other reports of a chemical liver carcinogen affecting cyclin D1 post-translationally.

We hypothesize that there are several possible mechanisms by which FB1 is causing cyclin D1 post-translational modification(s). (i) Although there are no activating mutations in the ras oncogenes, it is possible that FB1, by affecting the metabolism of sphingolipids (5), phospholipids and fatty acids (7), important constituents of cellular membranes, could increase p21 ras membrane association or signaling through the PI3K pathway (ultimately impacting on Akt and GSK-3ß, signaling molecules involved in cyclin D1 degradation). In addition to these effects, it is possible that FB1 is affecting the Raf/MEK/MAPK pathway and possibly influencing signals involved in cyclin D1 synthesis and assembly. (ii) Independent of ras, FB1 might alter the function of some proteins and signalling molecules in the control of the activation or inactivation of Akt. (iii) In addition to its effects on cyclin D1 protein stability/degradation or post-translational modifications due to the inhibition of GSK3-ß, it is possible that FB1 can affect some additional proteins responsible for cyclin D1 degradation and translation. Currently, we are investigating all of these possibilities.

In conclusion, dietary FB1 specifically causes activation of Akt, inhibition of GSK-3ß and overexpression of cyclin D1 related to post-translational modification(s). This causes a consequent increase in Cdk4 kinase activity, resulting in hyperphosphorylation of pRb and alteration in cell-cycle progression (G1/S) of rat hepatocytes. Our data contribute significantly to the understanding of the carcinogenic mechanism of FB1, a common environmental contaminant, recently shown to be carcinogenic in both rats and mice by the National Center for Toxicological Research of the US Food and Drug Administration (60).


    Notes
 
7 To whom correspondence should be addressed Email: ramljak{at}mail.ncifcrf.gov Back


    Acknowledgments
 
We wish to thank Drs K.Vousden, A.O.Perantoni, G.S.Buzard and M.Hunter Jamerson and Mrs A.B.Jones for valuable reviews; Ms K.Breeze for excellent secretarial assistance; Dr I.Karavanov for scientific input; and Mr J.F.Hochadel for technical help. We are grateful to Dr M.E.Ewan for the pGEX-Rb plasmid, Dr C.J.Sherr for the mouse cyclin D1 cDNA plasmid, and Dr W.H.Lee for the pRb antibody. This project has been funded in part by the National Cancer Institute under contract NO1-CO-56000. The content of this publication does not necessarily reflect the views nor policies of the Department of Health and Human Services, nor does the mention of trade names, commercial products or organizations imply endorsement by the US Government.


    References
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 Results
 Discussion
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Received ; accepted December 10, 1999.