Gene profiling of cells expressing different FGF-2 forms

Gene 356 (2005) 49 – 68 www.elsevier.com/locate/gene

Gene profiling of cells expressing different FGF-2 forms Natalina Quarto *, Kenton D. Fong, Michael T. Longaker Department of Surgery, School of Medicine Stanford University, 257 Campus Drive, Stanford, CA 94305-5148, United States Received 3 February 2005; received in revised form 18 April 2005; accepted 4 May 2005 Available online 14 July 2005 Received by R. Di Lauro

Abstract Fibroblast Growth Factor-2 (FGF-2) induces cell proliferation, cell migration, embryonic development, cell differentiation, angiogenesis and malignant transformation. The four forms of FGF-2 (Low Molecular Weight) and (High Molecular Weights) are alternative translation products, and have a different subcellular localization: the high molecular weight (HMWFGF-2) forms are nuclear while the low molecular weight form, (LMWFGF-2) is mainly cytoplasmic. Our previous work demonstrated NIH 3T3 cells expressing different FGF-2 forms, displayed a different phenotype, suggesting that nuclear and cytoplasmic forms of FGF-2 may have different functions. Here we report a cDNA microarray-based study in NIH 3T3 fibroblasts expressing different FGF-2 forms. Several candidate genes that affect cell-cycle, tumor suppression, adhesion and transcription were identified as possible mediators of the HMWFGF-2 phenotype and signaling pattern. These results demonstrated that HMWFGF-2 and LMWFGF-2 target the expression of different genes. Particularly, our data suggest that HMWFGF-2 forms may function as inducers of growth inhibition and tumor suppression activities. D 2005 Elsevier B.V. All rights reserved. Keywords: Gene array; FGF-2; Nuclear form; Growth inhibition; Tumor suppression

1. Introduction The Fibroblast Growth Factors (FGFs) constitute a large family of signaling molecules. Up to date, at least 24 family members have been identified in vertebrates (Yamashita et al., 2000; Ornitz and Itoh, 2001; Draper et al., 2004). Most FGFs are secreted via classical signal peptide dependent secretory pathways, while other family members (FGF-1 and FGF-2) use poorly understood alternate routes. The

Abbreviations: BSA, Bovine serum albumin; DAPI, 4V,6-Diamidino-2phenylindole dimethylsulfoxide; G418, Geneticin; SDS-PAGE, Sodium dodecyl sulfate-polyacrilamide gel electrophoresis; PBS, Phosphate-buffered saline; PCNA, Proliferating cell nuclear antigen; NfX-I, Nuclear factor I; Nupr, Nuclear protein 1; St5, Suppressor tumor 5; Egr-1, Early growth response-1; Mapk-6, Mitogen-activated protein kinase 6; Angptl4, Angiopoietin-like 4; Rps5, Ribosomal protein 5; FGF-2, Fibroblast growth factor 2; HMWFGF-2, High molecular weight FGF-2; LMWFGF-2, Low molecular weight FGF-2; WTFGF-2, Wild type FGF-2. * Corresponding author. Tel.: +1 650 736 1704; fax: +1 650 736 1705. E-mail address: [email protected] (N. Quarto). 0378-1119/$ - see front matter D 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.gene.2005.05.014

biological functions of the extracellular FGFs are mediated by their interaction with tyrosine kinase receptors (FGFR-1 to FGFR-4) (Friesel and Maciag, 1995). FGF-2 is produced naturally in five forms (Renko et al., 1990; Zuniga Mejia Borja et al., 1996). Molecular analysis of FGF-2 has shown that 3 to 4 protein forms are synthesized by alternative translation initiation from a single mRNA (Prats et al., 1988; Florkiewicz and Sommer, 1989), In addition, a fifth form, (isoform) called altFGf-2 is generated by an alternative splicing event in chicken (Zuniga Mejia Borja et al., 1996). The low molecular weight FGF-2 form (LMWFGF-2) initiates at an internal AUG codon, whereas the high molecular weight forms (HMWFGF-2) forms are synthesized by alternative translation from upstream in-frame CUG codons. The mechanism used to generate these multiple forms of FGF-2 from a single mRNA was elucidated by Florkiewicz and Sommer, and Prats and colleagues who showed that initiation of translation at CUG codons located 5V to the AUG codon accounted for the FGF-2 species of 24, 22.5, and 22 kDa (HMWFGF-2),

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whereas the AUG codon was used for the initiation of translation of the 18 kDa form (LMWFGF-2). As result, the three HMW forms of FGF-2 are colinear NH2-terminal extensions of the 18 kDa FGF-2 form at its COOH termini. When exogenously added, both HMWFGF-2 and LMWFGF-2 forms bind to the cell surface FGFRs with equal affinity, eliciting similar biological responses (Moscatelli et al., 1987). As FGF-2 forms do not contain a secretory signal sequence, their secretion from cells uses an alternative pathway which is independent of the ER –Golgi complex (Mignatti et al., 1992). Subcellular distribution of the different forms of FGF-2 analysed in cells expressing endogenous FGF-2, as well as cells transfected with FGF-2 human-cDNA, demonstrated that the 18 kDa form is primarily cytosolic, whereas the three HMWFGF-2 forms are predominantly nuclear (Renko et al., 1990; Bugler et al., 1991; Quarto et al., 1991a). The NH2-terminal extension of HMW forms is responsible for their nuclear distribution. Previous studies showed that the NH2-terminal extension of HMW forms contain a nuclear localization signal (NLS), which mediates their preferential intracrine nuclear accumulation (Bugler et al., 1991; Quarto et al., 1991a). In addition, the NH2-terminal extension of HMW forms contain glyarg-gly-arg-gly repeats in which the arginine residues appear to be mono- and dimethylated (Burgess et al., 1991). Studies by Pintucci and colleagues provided evidence that post-translational modification is essential for the nuclear accumulation of the HMW forms of FGF-2 (Pintucci et al., 1996). Different forms of FGF-2 with different subcellular distributions appear to affect the cell phenotype through autocrine FGF-R-dependent, or intracrine FGF-R-independent pathways (Moscatelli and Quarto, 1989; Bikfalvi et al., 1995; Nindl et al., 2004).The formation of different FGF-2 forms with a distinct subcellular localization suggested that the nuclear forms may be functionally distinct from the 18 kDa cytosolic form. We have previously demonstrated that transfected NIH 3T3 cells with the human cDNA that expresses only the HMWFGF2, the LMWFGF-2 or all forms (WTFGF-2) have different phenotypes. Particularly, the selective expression of HMW forms, at low levels, confers a unique phenotype to NIH 3T3 fibroblasts characterized by an impaired growth compared to the parental cells and tendency to form multinucleate giant cells (Quarto et al., 1991b). In contrast, the expression of LMW form induces NIH 3T3 transformation as measured by enhanced saturation density, growth in soft agar, and growth in low serum (Quarto et al., 1991b, 1989).The co-expression of a dominantnegative FGF receptor reverts the phenotype of cells transformed by the 18 kDa FGF-2, but has no effect on the phenotype of cells expressing the HMW forms (Bikfalvi et al., 1995). Moreover, in cells expressing HMW FGF-2 forms, the down-regulation of FGF-2 high

affinity receptors is similar to that of parental cells, while cells expressing the 18 kDa form have most of their FGF-2 high affinity receptors down-regulated (Moscatelli and Quarto, 1989). Collectively, these data suggested that different forms of FGF-2 with different subcellular distributions appear to affect the cell phenotype through autocrine FGF-R-dependent, or intracrine FGF-R-independent pathways. Indeed, a specific functional role for HMW FGF-2 forms may exist. Predominantly nuclear FGF-2 proteins have been observed in many cultured cells (Renko et al., 1990), but their functions remain largely unknown. During embryogenesis, nuclear translocation appears to be tightly regulated by developmental stage and cell-type specific mechanisms (Shiurba et al., 1991; Woodward et al., 1992; Dono and Zeller, 1994; Riese et al., 1996). Moreover, a predominant translation of HMW FGF-2 forms seems to be associated with more differentiated cell-state (Cowan et al., 2003). One of the ways in which cells adjust to their phenotype and their environment is by altering gene expression patterns. The development of DNA microarray technology has provided a powerful tool to measure gene expression of thousands of genes. In the past few years, the DNA microarray technology has been used to discover gene function, understand biochemical pathways and regulatory mechanisms, classify disease, and discover drug targets. To study possible functional differences of HMWFGF-2 and LMWFGF-2 forms we have investigated the changes in the gene expression profile of NIH 3T3 fibroblasts expressing different FGF-2 forms. Microarray analysis indicated differential expression of genes in cells expressing only HMWFGF-2 forms, only the LMWFGF-2 form, or all forms (WTFGF-2). Taken together, these data shed new light on the mechanisms of action of different FGF-2 forms, and suggests a potential specific role of the nuclear FGF-2 forms.

2. Materials and methods 2.1. Cell culture and transfection NIH 3T3 cells were grown in Dulbecco’s modified Eagle’s medium (DMEM) containing 10% fetal bovine serum (GIBCO, Invitrogen Corporation). Cells (3  105 / 100-mm plate) were transfected with the Zip-neo vector containing the human FGF-2 cDNAs as described previously (Quarto et al., 1989), using the lipofectin reagent according to manufacturer’s instructions. After 42 h in culture, the cells were split 1 : 5 and cultured thereafter in the presence of selective DMEM containing 10% fetal calf serum and 500 ug/ml of G418 (GIBCO, Invitrogen Corporation). Two weeks later, G418 resistant clones were isolated, grown up, and tested for the expression of FGF-2 forms by immunoblot analysis. Cells

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transfected with the cDNA encoding for 22, 22.5 and 24 kDa FGF-2 forms are operationally referred to as HMWFGF-2 clones, cells transfected with the cDNA encoding for 18 kDa FGF-2 form are referred to as LMWFGF-2 clones, whereas cells transfected with the wild-type cDNA encoding for all FGF-2 forms, are referred as to WTFGF-2 clones. Control NIH 3T3 cells were transfected with DNA from plasmid vector minus the FGF-2 cDNA insert and are referred to as Zip clones. Several clones were isolated, expanded and analyed for FGF-2 expression. Four clones for each FGF-2 transfected plasmid were analyzed by microarray. Each clone was used separately for the microarray. Importantly, these clones expressed relatively low levels of FGF-2. Southern blot analysis indicated that 1– 2 copies of cDNA were integrated in the genome of these clones (data not shown). 2.2. Immunoblotting and ELISA assay Cell lysis and immunoblot analysis were performed following protocols that have been described in detail previously (Quarto et al., 1989, 1991b). Briefly, immunoblots on total protein were performed using 50 –150 Ag of protein lysate. Antibodies against FGF-2 (174, sc-79), PCNA (FL-261, sc7907), Egr1 (588, sc-110) MAPK-6 (I15, sc-156) and NFI-X (sc-5567) were purchased from Santa Cruz Biotechnology. To control for protein loading, membranes were re-probed with mouse monoclonal hactin antibodies (AC-15, ab6276-100, abcam.com, Cambridge, MA). Horseradish peroxidase (HPR)-conjugated anti-rabbit and anti-mouse secondary antibodies (Amersham Biosciences) were used, and proteins were visualized by enhanced chemiluminescence (ECL, Amersham Biosciences). ELISA assay was performed using the FGF-2 Chemikine Human bFGF EIA kit (Cat.No. CYT142) purchased from Chemicon International, the assay was carried out according to the manufacturer’s protocol. Each clone was assayed in triplicate. 2.3. Immunofluorescence For the immunofluorescence analysis cells (8  10 4) were seeded on 8-well Lab-Tek II chamber slides (Nalge Nunc International, Naperville, IL) in medium containing 10%FBS. After 24 h, the media was removed and replaced with fresh serum-free medium, cells were cultured for additional 18 h prior performing immunofluorescence. Cells were washed twice with cold PBS and fixed for 5 min with MeOH at 20 -C, then for 3 min with Acetone. After two washes with PBS cells were blocked with PBS/1% triton X-100/2% BSA at 37 -C for 2 h. Cells were incubated with primary antibodies for 1 h at 37 -C, washed twice with phosphate buffered saline solution, then incubated with secondary antibodies for 1 h at 37 -C in the dark and washed twice with phosphate

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buffered saline solution. The primary antibodies to PCNA and NFI-X were for the immunoblot analysis. Fluorescein-conjugated anti-rabbit IgG (Vector FI-1000) were used as secondary antibody (Vector Laboratories, Burlingame, CA). Nuclei were stained with 300 nM DAPI (Molecular Probes, Eugene,OR) for 5 min then rinsed with water. Coverslips were mounted and fluorescence was enhanced with Vectashield (Vector Laboratories, Burlingame, CA). Photographs were taken with a Zeiss AxioPlan Immunofluorescence microscope equipped with Zeiss AxioCam HRc digital camera (Carl Zeiss MicroImaging, Inc., Thornwood, NY). Controls for each antibody consisted of probing with the secondary antibody in the absence of primary antibody. 2.4. RNA isolation In order to eliminate stimuli due to any growth factor present in the serum and enhance FGF-2 activity, exclusively, subconfluent cells were incubated for 18 h in serum free medium prior to RNA isolation. The same culture conditions were used also for immunoblotting and immunofluorescence analysis. Cells were then harvested with cold PBS and RNA extraction (from four clones for each FGF-2 transfected plasmid) was performed using TriZol (Invitrogen Corporation) according to the manufacturer’s instruction. Isolated RNAs were treated with DNase I to minimize any genomic contamination. RNAs were amplified using the MessageAmp aRNA kit (Ambion, Austin, TX) according to the manufacturer’s instructions. 2.5. Microarray analysis 2.5.1. cDNA micro-array chips and postprocessing We utilized mus musculus cDNA large-scale microarrays manufactured by the Stanford Functional Genomics Facility (www.microarray.org). This microarray core facility has printed over 7000 large-scale high quality microarrays containing 42,000 unique cDNA elements created from the RIKEN and NIA mouse clone libraries. Prior to use, slides were rehydrated in a hydration chamber for 20 min at room temperature followed by snap drying at 70 -C. The poly-l-lysine surface was blocked by treatment with 0.05% succinic anhydride prepared in a buffer solution consisting of 350 ml of 1methyl-2-pyrrolidinone and 15 ml of boric acid for 20 min on a shaker. The cDNAs on the slides were then denatured in distilled water for 2 min at 95 -C followed by fixation with 95% ethanol. 2.5.2. Microarray labeled cDNA probe preparation The total RNA extracted from each sample and the Stratagene Mouse Universal Standard (La Jolla, CA) was amplified into aRNA using the Ambion Message AMP Kit (Austin, Tx) following manufacturer protocols. Synthesis

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of the fluorescently labeled cDNA probes used for hybridizing to the microarrays included 5 micrograms of aRNA per sample. The fluorescent-labeled nucleotide analogs Cyanine3-dUTP (Cy3-dUTP) and Cyanine5-dUTP (Cy5-dUTP) were obtained from Amersham (Piscataway, NJ). RNA extracted from FGF-2 expressing cells was labeled with Cy5-dUTP and mouse universal standard RNA (reference sample) was labeled with Cy3-dUTP during the reverse transcription using pdT24 primers. The reverse transcription reaction was performed as follows: a master mix consisting of 200U Superscript II (GibcoBRL), buffer, dNTPs, and fluorescent nucleotides was added to the RNA. The final concentration of dATP, dCTP, and dGTP was 500 AM and the final concentration of dTTP was 200 AM. Cy3-dUTP and Cy5-dUTP was used at a final concentration of 100 AM. The reaction was incubated at 42 -C for two h. One hour into the incubation, an additional 1 Al of Superscript II was added and the incubation continued. Unincorporated fluorescent nucleotides were removed by diluting the reaction mixture with 470 Al of 10 mM tris-HCl (pH 8.0)/1 mM EDTA. Each Cy5-labeled sample (FGF-2 expressing cells) was equally mixed with the Cy3-labeled common reference sample (mouse universal standard). The mixture was concentrated to approximately 20 Al using centricon-30 microconcentrators (Amicon, Bedford, MA). Each labeled cDNA probe derived from a different FGF-2 clone was hybridized on separate microarray slide. Four slides for each FGF-2 group (four clones) were used. Probe hybridization was performed by adding 10 Ag of yeast tRNA, 10 Ag of polydeoxyadenylic acid, and 20 Ag of human CoT1 DNA (Invitrogen) to a solution containing 3X standard saline citrate (SSC), 0.3% sodium dodecyl sulfate (SDS), and labeled probes. This mixture was loaded onto the microarray surface and covered with a glass cover slip. The array was subsequently transferred to a hybridization chamber (GeneMachine) and incubated for 18 h at 65 -C. After incubation, the array was washed for 5 min at room temperature in 2 SSC with 0.03% SDS, then twice for 10 min at room temperature in 2 SSC, followed by 0.1 SSC. Each labeled cDNA probed derived from a different FGF-2 clone was hybridized on separate microarray slide. 2.5.3. Scanning and analyzing arrays Arrays were immediately scanned with a computercontrolled stage and microscope objective using a GenePix Scanner (Axon Instruments, Foster City, CA). Sequential excitation of the two fluorophores was accomplished via a mixed gas, multiline laser. Emitted light is split according to wavelength and detected with two photomultiplier tubes. The final signal was read into a PC utilizing a 12-bit analog-to-digital board. Images were scanned at a resolution of 10 Am per pixel. Normalization of the two channels to overall intensity was performed by

adjusting the photomultiplier and laser power settings such that the signal ratio at these elements was as close to 1.0 as possible. Dividing the intensity of each pixel in a binding box on the array by the total number of pixels yielded the average fluorescence intensity. The array images were analyzed by GenePix Pro 4.0 software (Axon Instruments) and imported to the Stanford Microarray Database (SMD) for further comparison and interpretation. In addition we utilized Significance Analysis of Microarrays (SAM) to perform supervised gene expression analysis. SAM is validated statistical technique to identify differentially expressed genes across high density microarray. In addition to identify significantly differentially expressed genes, this technique gives an estimated false discovery rate that represents genes that have been identified by chances. SAM data was normalized between samples for both gene expression and overall array expression levels. Only data points with correlation coefficients of > 0.4 and mean spot intensity/median background intensity for both the red or green channels < 2.0 were included in our analysis. 2.6. Real-time qRT-PCR Purified and quantified RNA were treated with DNAse I (Ambion, Austin, TX) to clear genomic DNA. Three micrograms of total RNA from each clone was reverse transcribed to cDNA using random primer hexamers (Invitrogen, Carlsbad, CA). The reverse transcription proceeded for 1 h at 42 -C, followed by 5 min incubation at 95 -C to inactivate the reverse transcriptase. For quantitative real-time PCR, primers were designed with Primer Expressi software (Applied Biosystems). Each primer was subjected to PCR to ensure single primary amplicon as evidenced by 2% agarose gel electrophoresis to be < 200 bp for maximum efficiency. SYBR\ Green PCR Master Mix (Applied Biosystems) was used for fluorescence. Samples along with primers and Syber Green Master Mix (Applied Biosystems, Foster City, CA) were loaded in 384 well sealed plates and the reaction was run in an ABI Prism 7900 HT (Applied Biosystems) as follows: samples were cycled 45 times from 95 -C for 15 s (denaturation) to 61 -C for 1 min (annealing, extension). In addition to experimental samples, a set of negative (minus cDNA) controls were run for each primer and probe combination on each 384 well plate. Gapdh (Applied Biosystems) was used for internal control. Standard curve method of quantitation was used to calculate the expression of target genes relative to the house keeping gene Gapdh. Four serial dilutions of cDNA (1 : 4) were made for the calibration curve and trend lines were drawn using Ct values versus log of dilutions for each target gene and Gapdh run in triplicate with correlation coeffient (R 2 > 0.99). Relative expressions were calculated using line equations derived from

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calibration curves and obtaining ratios of target gene to Gapdh. For each gene, experiments were run at least three times using cDNAs obtained from three independent RNA purifications. Similar results were obtained from each experiment. Primers sequence: Angptl-4 forward primer was 5VCAGAGGGACCACTACAGTCCAACTA 3V, Angptl-4 reverse primer was 5VCACCCTGTCTCCAGTCAGTCAA 3V; NfI-X forward primer was 5VACCCAACCATCCGCTACCA3V, NfI-X reverse primer was 5VGAGCACACAAACTGCACAAACTC3V; Nupr-1 forward primer was 5VCCCTTCCCAGCAACCTCTAAA3V, Nupr-1 reverse primer was 5VGGGCCAGGCTGTACTGATCA 3V; Egr-1 forward primer was 5V TCCTTTTCTGACATCGCTCTGA, Egr-1 reverse primer was 5VCGAGTCGTTTGGCTGGGATA 3V; Mapk-6 forward primer was 5VAAAGGTACCACGATTGTCAGTTCTC 3V, Mapk-6 reverse primer was 5VGTCAAGCTGCACCTCATCGATAT 3V; St5 forward primer was 5VGAGAGGCCTTCC GCAAGTC 3V, St5 reverse primer was 5VGAAGCCAGCAAACATCTGAGATT 3V; Rps5 forward primer was 5VCCTCCAATTCCTATGCCATCAA 3V; Rps5 reverse primer was 5VCTGGGAAATCAGCGGTTAGACTT 3V.

3. Results 3.1. Characterization of FGF-2 expressing clones NIH 3T3 cells were transfected with plasmids encoding HMWFGF-2, LMWFGF-2 and WTFGF-2 forms. In our earlier study to analyze the phenotype of cells expressing selective forms of FGF-2, we chose NIH 3T3 fibroblasts because their endogenous level of FGF-2 is extremely low (Quarto et al., 1989), while they express high levels of FGF receptor 1 and receptor 2 (Moscatelli and Quarto, 1989), which confer a prompt and full response to FGF-2 ligand . In addition, several studies of FGF-2 signaling have been performed in these cells. For the microarray analysis, we selected stable transfectants expressing different FGF-2 forms in the same order of magnitude to ensure that differences in gene expression observed between HMWFHF-2 and LMWFGF-2 expressing cells were not due to differences in the amount of FGF-2 level expressed by each clones, but rather due to qualitative differences between the nuclear (HMWFGF-2) and the cytoplasmic (LMWFGF-2) forms of FGF-2. To accomplish this we tested the expression of FGF-2 forms in several clones after stable transfection and isolation. We selected four independent clones for each different FGF-2 form (HMWFGF-2 and LMWFGF-2), as well as for Zip control and WTFGF-2. All selected clones expressed a moderate level of FGF-2. FGF-2 immunofluorescence staining was performed in the Zip control, HMWFGF-2, LMWGF-2 and WTFGF-2 expressing cells (Fig. 1A). Nuclear FGF-2 localization

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was observed in HMWFGF-2 cells, whereas FGF-2 localized primarily to the cytoplasm and perinuclear area in LMWFGF-2 cells. Both nuclear and cytoplasmic staining was present in WTFGF-2 cells, while no FGF2 staining was detected in Zip control cells. Fig. 1B shows the expression levels and qualitative analysis of FGF-2 forms in all selected clones. As expected, 24 and 22.5 kDa bands were detected in HMWFGF-2 clones, while a single 18 kDa band was detected in LMWFGF-2 clones, and all three bands (24, 22.5 and 18 kDa) were present in WTFGF-2 clones. No immunoreactivity bands were detected in Zip control clones. The amount of FGF2 was further quantitated by ELISA immuno assay. (Fig. 1C). As shown in Fig. 1B and C, all clones expressed similar amount of FGF-2, whereas Zip clones expressed extremely low levels of FGF-2 which could be detected only by ELISA assay. These clones were subsequentially used for genomic cDNA microarray analysis. Our previous data demonstrated that fibroblasts expressing exclusively HMWFGF-2 grew much slower than fibroblasts expressing LMWFGF-2, WTFGF-2 or Zip control cells. Those results were obtained by performing a growth curve assay (Quarto et al., 1991b). Here we have tested the proliferating cell nuclear antigen (PCNA) level in different FGF-2 expressing clones. Immunoblotting analysis and immunofluorescence staining showed significantly lower levels of PCNA in HMWFGF-2 expressing cells when compared to the other cells (Fig. 2A and B). Thus, these data are consistent with our earlier observation (Quarto et al., 1991b). Surprisingly, the PCNA gene down-regulation was not detected by microarray analysis, although we could detect difference in PCNA expression by PCR analysis. A possible explanation is that the PCNA spot was missed on the chip arrays. 3.2. Differential gene expression in HMW and LMWFGF-2 expressing cells To evaluate global differences in gene expression associated with selective expression of different FGF-2 forms (nuclear and cytoplasmic), we utilized cDNA microarray analysis to examine mRNA expression patterns from NIH 3T3 fibroblasts expressing HMWFGF-2, LMWFGF-2, WTFGF-2 forms, or Zip vector alone. We analysed a total of four separate clones for each condition using Pearson correlation hierarchical clustering. After data quality filtration and normalization of expression by both mean gene expression and mean array intensity, we filtered differentially expressed clones that had > 2-fold change in at least one array experiment. We further filtered out genes expressed in < 80% of the experiments and arrays having < 70% available data. A dendrogram illustrating the relationship between the expression patterns of each experiment is shown in Fig. 3A. Arrays with the most similar gene expression profiles cluster together with the shortest branches. By utilizing this global overview of

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Fig. 1. Expression of different FGF-2 forms: A. Immunofluorescence showing different FGF-2 subcellular localization in HMWFGF-2, LMWFGF-2 and WTFGF-2 expressing clones. HMWFGF-2 forms localized in the nucleus, while LMWFGF-2 form was perinuclear and cytoplasmic, both nuclear and cytoplasmic FGF-2 localization was observed in WTFGF-2 clones, and no FGF-2 immunofluorescence staining was detected in Zip control cells (40 magnification). B. Qualitative FGF-2 immunoblotting analysis of Zip control, HMWFGF-2, LMWFGF-2 and WTFGF-2 clones. Cell lysates (50 Ag) were run on SDS-13% PAGE, transferred to Immobilon-P membrane (Millipore) and analyzed using anti-FGF-2 antibody as described in Section 2.2. No FGF-2 immunoreactivity was detected in Zip control clones, HMWFGF-2 clones expressed the 24 kDa and 22,5 kDa forms, LMWFGF-2 clones expressed only the 18kDa form, while the 24, 22.5 and 18 kDa forms were present in the WTFGF-2. C. ELISA assay to quantitate the concentration of FGF-2 in different clones showed that the FGF-2 clones selected for the microarray study expressed similar amount of total FGF-2. Very low level of endogenous FGF-2 was detected in Zip control clones. Each clone (20 Ag) was assayed in triplicate. Values are means of triplicate.

differential expression, we found distinct expression patterns in each experimental group (with individual clone replicates clustering closely together by experimental group) (Fig. 3A). Although grossly there appeared to be clear differences in gene expression between HMWFGF-2 and LMWFGF2, we further statistically evaluated the differences in expression with significance analysis of microarrays

(SAM). The SAM procedure was performed on 13,647 genes, and we found 251 genes with greater than 2-fold higher expression and 210 genes with greater than 2-fold lower expression in the HMWFGF-2 samples compared to the LMWFGF-2 samples with a predicted mean number of false significant genes of 3.30615 using a Delta of 0.08443 (Fig. 3B). Comparison of Zip vector to either HMWFGF-2, LMWFGF-2, or WTFGF-2 revealed signifi-

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Fig. 2. Comparison of proliferating cell nuclear antigen (PCNA) in cells expressing different FGF-2 forms: A. Immunoblotting analysis of Zip control, HMWFGF-2, LMWFGF-2 and WTFGF-clones. Cell lysates (100 Ag) were run on SDS-13% PAGE, transferred to Immobilon-P membrane (Millipore) and analyzed using anti-PCNA antibody as described in Section 2.2. Immunoblot analysis showed that HMWFGF-2 clones expressed decreased amount of PCNA. The same membranes were stripped and re-probed with h-actin antibody as loading control. B. Immunofluorescence staining with PCNA antibody showed decreased nuclear staining in HMWFGF-2 clones, thus further demonstrated the low level of PCNA in HMWFGF-2 clones. The same clones were counterstained with Dapi to detect nuclei, as described in Section 2.3.

cant and distinct up-regulation or down-regulation of many genes, including genes that encode proteins playing a role in cell cycle, chromatin remodeling, regulation of transcription and cell adhesion. In addition, many novel genes were also found to be differentially expressed in each experimental group. The list of significantly differentially expressed genes (with more than 3-fold change) between HMWFGF-2 and LMWFGF-2 can be found in Tables 1 and 2. Fig. 3C shows upregulated and downreguraleted genes (at least 2-fold) in HMWFGF-2 cells grouped into different categories. 3.3. Validation of differential expression using real-time qRT-PCR and proteins analysis Because our interest was to elucidate the molecular mechanism(s) underlying the phenotype of HMWFGF-2 cells we focused on validation of genes which could have a potential role in development of this phenotype. We selected seven genes from the list of genes that were differentially regulated for validation by quantitative realtime RT-PCR, as well as protein analysis, when available. The rationale for selecting these seven genes was based on their potential cellular function. Overall, our RT-PCR data agreed well with the microarray data. The first gene we validated by quantitative real-time RTPCR was the Nuclear Factor I-X (NfI-X). NfI-X belongs to the Nuclear Factor I family of site-specific DNA proteins (also known as CTF or CAAT box transcription factors) playing multiple roles in animal physiology, biochemistry, and pathology. Expression of NFI proteins has been

associated with changes in the growth state of cells and with a number of oncogenic processes and disease states (Gronostajski, 2000). To validate the microarray data for NfI-X (Fig. 4A), we performed quantitative real-time RT-PCR, Western blotting and immunofluorescence analysis on different FGF-2 expressing clones. Quantification of NfI-X mRNA revealed upregulation of NfI-X in all HMWFGF-2 clones compared to LMWFGF-2 clones and Zip clones (Fig. 4B). An increased expression of NfI-X was also detected in WTFGF-2 clones, but it was less significant than that observed in HMWFGF-2 clones (Fig. 4A and B). Both Western blotting and immunofluorescence analysis showed data mirroring the mRNA profiles (Fig. 4C and D). Thus, both mRNA and protein results confirmed microarray data. The increased expression of NfI-X in WTFGF-2 clones compared to LMWFGF-2, suggests a ‘‘dominant’’ inducing activity of HMWFGF-2, with respect to LMWFGF-2, on NfI-X gene expression. Nuclear protein 1 (Nupr1) was another gene upregulated in HMWFGF-2 compared to LMWFGF-2 and Zip control clones (Fig. 5A). Nupr1 is a stress-induced DNA-binding protein, biochemically related to the architectural chromatin binding HMG protein family (Mallo et al., 1997). Quantitative real-time RT-PCR showed that Nupr1 was upregulated in HMWFGF-2 clones (Fig. 5B), thus validating the microarray data. Nupr1 was also slightly increased in WTFGF-2 clones (Fig. 5A and B), likely as result of a balanced coexpression of HMWFGF-2 and LMWFGF-2 forms. We also validated the expression of the Suppressor Tumor 5 (St5) gene, which had higher expression in

56 N. Quarto et al. / Gene 356 (2005) 49 – 68 Fig. 3. Expression profiles and clustering of cells expressing different forms of FGF-2: A. Hierarchical clustering overview of differential expression in Zip control, WTFGF-2, HMWFGF-2 and LMWFGF-2 clones. Individual spots for each experiment were filtered for quality (correlation co-efficient of >0.4, signal to background ratio >2.0 for each channel) and then normalized to the mean expression of both individual genes and arrays. Genes with at least 2-fold change in expression in at least one array experiment and having 80% of data present were then hierarchically clustered. Note that individual clone replicate experiments clustered together by condition, and each condition had distinct expression profiles. B. Comparison of expression between HMWFGF-2 and LMWFGF-2 clones using SAM analysis and correlation plot. There were 251 significant genes with >2-fold higher expression in HMW samples compared to LMW samples. There were 210 significant genes with >2-fold lower expression in HMW samples compared to LMW samples. The predicted mean number of false significant genes was 3.30615 with a Delta of 0.08443. Green indicates lower than mean expression and red indicates higher than mean expression. C. Array targets showing upregulated and downreguraleted genes in HMWFGF-2 cells grouped into different categories. The upregulated and downregulated genes were categorized using the S.O.U.R.C.E program (http:// source.standford.edu) based on sequence analysis and are shown in the separate pie graphs. Color coding for the different groupings is shown on the side.

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Table 1 Transcripts up-regulated at least 3-fold in HMWFGF-clones Encoded protein, EST, Riken cDNA Growth inhibition and tumor suppression Nuclear factor I/X* Nuclear protein 1* Suppression of tumorigenicity 5* MAD2 (mitotic arrest deficient, homolog)-like 1 Transformation related protein 53 Cell-adhesion/Extracellular matrix molecules Procollagen, type VI, alpha 3 Cell adhesion molecule-related Neuropilin Development/Differentiation Odd Oz/ten-m homolog 4 (Drosophila) Troponin T2, cardiac four and a half LIM domains 1 Proteolipid protein 2 DNA binding RIKEN cDNA C530025K05 gene AD homolog 4 (Drosophila) RNA binding/processing RIKEN cDNA 0610027F08 gene Biosynthesis Methionine adenosyltransferase, alpha Pyrroline-5-carboxylate reductase 1 Leucyl-tRNA synthetase Procollagen-proline 4-hydroxylase, alpha II polypeptide Hydroxysteroid (17-beta) dehydrogenase 12 Binding protein Secreted acidic cysteine rich glycoprotein Transcribed sequence with strong similarity to protein:sp:P06454 Secreted acidic cysteine rich glycoprotein Kinesin family member 23 Hypotetical H19 fetal liver mRNA Calpain 6 RIKEN cDNA 4930432B04 gene RIKEN cDNA 2610318I01 gene RIKEN cDNA C330027G06 gene Mitosis/meiosis cDNA sequence BC049989 Protein binding Pre-B-cell leukemia transcription factor interacting protein 1 LIM domain containing preferred translocation partner in lipoma Structural protein Smoothelin RIKEN cDNA 3110027H23 gene ceroid lipofuscinosis, Neuronal 3, juvenile Tropomyosin 1, alpha Translation Basic leucine zipper and W2 domains Transport/energy Solute carrier family 6, member 6 Solute carrier family 29 (nucleoside transporters), member 1 Unknown AV025941 AW553597 AV133669 AV111488 Proteolipid protein 2 Expressed sequence C80913

Gene ID

Gene Accession #

Fold change

Nfix Nupr1 St5 Mad2l1 Trp53

BG075770 BG072110 BG075828 AA033406 BG075029

25.96 6.54 4.69 3.89 3.54

Col6a3 Cdon Nrp

AV012366 BG075737 BG073453

3.77 3.76 3.48

Odz4 Tnnt2 Fhl1 Plp2

BG071076 AV006224 BG074978 AI893212

5.00 4.63 3.79 3.88

C530025K05Rik Madh4

AV021105 BG073878

4.01 3.81

0610027F08Rik

BG074255

3.48

Mat2a Pycr1 Lars P4ha2 Hsd17b12

BG073682 BG073280 BG064998 BG063134 BG064974

3.38 4.46 3.32 3.06 3.25

Sparc THY_HUMAN Sparc Kif23

BG072874 AV130619 AV104148 BG068324

6.46 3.51 4.36 4.66

H19 Capn6 4930432B04Rik 2610318I01Rik C330027G06Rik

BG063261 AA050030 BG066884 BG073078 BG074935

14.87 9.92 4.68 4.43 4.41

BC049989

AW557070

3.29

Pbxip1 Lpp

AW556431 BG068912

3.17 3.17

Smtn 3110027H23Rik Cln3 Tpm1

BG066516 BG073013 BG075591 AW548270

4.8l 4.61 3.39 3.88

Bzw1

BG064777

3.28

Slc6a6 Slc29a1

C81465 BG075739

4.35 7.97

BG064776 BG074199

9.87 4.74 4.63 4.43 3.83 3.41

Plp2 C80913

All transcripts result from SAM analysis between HMWFGF-2 and LMWFGF-2 clones. Bold entries highlight genes of interest. Asterisks indicate genes which have been validated by quantitative real-time RT-PCR.

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Table 2 Transcripts down-regulated at least 3-fold in HMWFGF-clones Encoded protein, EST, Riken cDNA

Gene ID

Gene Accession #

Fold change

Cell proliferation/mitogenic signaling(-) Early growth response 1* Early growth response 1 Mitogen-activated protein kinase 6* Kit ligand Nucleolar protein 1

Egr1 Egr1 Mapk6 Kitl Nol1

BG070825 AA119590 BG072253 AA404089 BG064905

7.06 8.02 4.99 4.35 3.87

Angiogenesis Angiopoietin-like 4*

Angptl4

AV084034

6.99

RNA binding/ribosome biogenesis Ribosomal protein S5* Ribosomal protein S24 Ribosomal protein S24 Ribosomal protein S5 Ribosomal protein S27-like Ribosomal protein L6 (Asp-Glu-Ala-Asp) box polypeptide 25

Rps5 Rps24 Rps24 Rps5 Rps27l Rpl6 Ddx25

BG072601 AV103362 BG075284 BG072598 BG063163 BG075206 AV043347

3.99 3.89 2.38 3.98 2.99 3.55 3.19

DNA binding High mobility group AT-hook 2

Hmga2

BG073094

3.93

Biosynthesis S-transferase, alpha 4 Rho, GDP dissociation inhibitor (GDI) beta

Gsta4 Arhgdib

AV084880 AA276890

6.54 3.16

Metabolism Crystallin, lambda 1

Cryl1

AA108958

5.65

Dlk1 S100a13

AA407811 AW542372 AV083696

3.02 3.89 3.35

9130413I22Rik Zfp503

AI841222 BG074838 AU023385 BG072688 BG072823 BG070657 AV149864

3.96 4.56 4.49 4.49 3.32 3.99 3.16

BG064389 AV049179 AV000246 BG074344 BG065216

3.24 3.32 3.27 5.15 3.84

Binding protein Delta-like 1 homolog (Drosophila) S100 calcium binding protein A13 AV083696 Unknown RIKEN cDNA 9130413I22 gene Zinc finger protein 503 AU023385 RIKEN cDNA 2310076K21 gene RIKEN cDNA 9130413I22 gene RIKEN cDNA 9930012K11 gene RIKEN cDNA 1500016L11 gene Cellular component Actin related protein 2/3 complex, subunit 1B DNA AV049179 Actin related protein 2/3 complex, subunit 1B Mesothelin RIKEN cDNA 2700060E02 gene

2310076K21Rik 9130413I22Rik 9930012K11Rik 1500016L11Rik

Arpc1b Arpc1b Msln 2700060E02Rik

All transcripts result from SAM analysis between HMWFGF-2 and LMWFGF-2 clones. Bold entries highlight genes of interest. Asterisks indicate genes which have been validated by quantitative real-time RT-PCR.

HMWFGF-2 clones arrays (Fig. 5C). Quantitative realtime RT-PCR confirmed that St5 expression was upregulated exclusively in HMWFGF-2 clones (Fig. 5D). In contrast to NfI-X and Nupr1 gene expression profiles, St5 expression remained low in WTFGF-2 clones. This result suggests that LMWFGF-2 acts as a ‘‘trans-dominant’’ repressor of HMWFGF-2 inducing activity of St5 gene expression.

Quantitative real-time RT-PCR also validated the genes which were found to be down-regulated in HMWFGF-2 clones, such as early growth response-1 (Egr-1) and mitogen-activated protein kinase 6 (Mapk-6). Egr-1 is a transcription factor induced by stress, injury, mitogens, and differentiation factors (Gashler and Sukhatme, 1995). This gene regulates the expression of genes involved in growth control or survival (Lim et al., 1998).

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Fig. 4. Expression profile of Nuclear FactorI-X gene in cells expressing different FGF-2 forms: A. Microarray profile of NfI-X gene in Zip control, HMWFGF-2, LMWFGF-2 and WTFGF-2 clones showed that the gene is upregulated in all four HMWFGF-2 clones compared to LMWFGF-2, as well as WTFGF-2 and Zip clones. B. Quantitative real-time RT-PCR. Quantified mRNA values were normalized by the amounts of Gapdh mRNA, and results are given as fold induction. All samples were run in triplicate as described in Section 2.6. Quantitative real-time RT-PCR correlated with microarray data. C. Immunoblotting analysis using anti NFI-X antibody demonstrated high NFI-X protein level in HMWFGF-2 clones. Cell lysates (100 Ag) were run on SDS-9% PAGE, transferred to Immobilon-P membrane (Millipore) and analyzed using anti-NFI-X antibody as described in Section 2.2. D. Immunofluorescence, with anti-NFI-X antibody detected intense nuclear staining in HMWFGF-2 cells, and less intense nuclear staining was observed also in WTFGF-2 clones, while very little immunofluorescens signals were detected in LMWFGF-2 and Zip contol clones. Thus, immunofluorescence, confirmed the immunoblotting analysis data. Membranes were stripped and re-probed with h-actin antibody as loading control. Cell culture conditions were the same for each experiment as described in Section 2.1.

We analyzed the expression of Egr-1 in our cells both by quantitative real-time RT-PCR and immunoblotting analysis. As shown in Fig. 6, the expression of Egr-1 was decreased in all HMWFGF-2 clones compared to other clones, both at mRNA (Fig. 6A and B) and protein levels (Fig. 6C). Mitogen-activated protein kinase 6 (Mapk-6) expression was also drastically decreased in cells expressing HMWFGF-2 (Fig. 6D). Mapk-6, known also as the Erk3 gene (Turgeon et al., 2000), belongs to the serine/threonine protein kinase superfamily that plays a major role in transducing extracellular chemical and physical signals into intracellular responses. Both quantitative real-time RT-PCR and immunoblotting analysis validated the initial results obtained from the microarray analysis (Fig. 6E and F).

In LMWFGF-2 clones, the proangiogenic factor angiopoietin-like 4 (Angptl4), was one of the genes specifically upregulated compared to HMWFGF-2, WTFGF-2 and Zip control cells. Our microarray data demonstrated that Angptl4 gene was upregulated 5-fold in cells expressing the LMWFGF-2 form (Fig. 7A). To our knowledge this is the first observation that FGF-2 induces the expression of Angptl4. Quantitative real-time RT-PCR confirmed the upregulation of Angtl4 in LMWFGF-2 clones (Fig. 7B). Interestingly, no increase of the Angtl4 expression level was observed in WTFGF-2 clones. This data suggest that HMWFGF-2 counteracts the inducing activity of LMWFGF-2 for the expression of Angptl4. Ribosomal protein 5 (Rps5) was another gene exclusively upregulated in LMWFGF-2 cells (Fig. 7C).

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Fig. 5. Expression profiles of Nupr and St5 genes in cells expressing different FGF-2 forms: A. Microarray profile of Nupr-1 gene in Zip control, HMWFGF-2, LMWFGF-2 and WTFGF-2 clones showed that the gene was up-regulated in all four HMWFGF-2 clone compared to LMWFGF-2, as well as WTFGF-2 and Zip clones. B. Quantitative real-time RT-PCR. Quantified mRNA values were normalized by the amounts of Gapdh mRNA, and results are given as fold induction. Quantitative real-time RT-PCR correlated with microarray data. C. Microarray profile of St5 gene in Zip control, HMWFGF-2, LMWFGF-2 and WTFGF-2 showed that the gene was up-regulated in all four HMWFGF-2 clone compared to LMWFGF-2, as well as WTFGF-2 and Zip clones. D. Quantitative real-time RT-PCR. Quantified mRNA values were normalized by the amounts of Gapdh mRNA, and results are given as fold induction. Quantitative real-time RT-PCR correlated with microarray data. All samples were run in triplicate described in Section 2.6.

Quantitative real-time RT-PCR analysis confirmed the initial observation obtained from the genomic cDNA microarray study (Fig. 7D). Previous work by Bouche

and colleagues demonstrated that exogenously added LMWFGF-2 enters the nucleolus and stimulates the transcription of ribosomal genes in adult bovine aortic

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Fig. 6. Expression profile of Egr-1 and Mapk-6 genes in cells expressing different FGF-2 forms Expression profile of Egr-1 in cells expressing different FGF-2 forms: A. Microarray profile of Egr-1 in Zip control, HMWFGF-2, LMWFGF-2 and WTFGF-2 clones showed that the gene was down-regulated in all four HMWFGF-2 clones compared to LMWFGF-2, as well as WTFGF-2 and Zip clones. B. Quantitative real-time RT-PCR. Quantified mRNA values were normalized by the amounts of Gapdh mRNA, and results are given as fold induction. Quantitative real-time RT-PCR correlated with microarray data. All samples were run in triplicate described in Section 2.6. C. Immunoblotting analysis using anti-Egr1 antibody demonstrated extremely low Egr-1 protein level in HMWFGF-2 clones. Cell lysates (150 Ag) were run on SDS-9% PAGE, transferred to Immobilon-P membrane (Millipore) and analyzed using anti-Egr-1 antibody as described in Section 2.2. Membranes were stripped and re-probed with h-Actin antibody as loading control. D. Microarray profile of Mapk-6 gene in Zip control, HMWFGF-2, LMWFGF-2 and WTFGF-2 clones showed that the gene was down-regulated in all four HMWFGF-2 clones compared to LMWFGF-2, as well as WTFGF-2 and Zip clones. E. Quantitative real-time RT-PCR. Quantified mRNA values were normalized by the amounts of Gapdh mRNA, and results are given as fold induction. Quantitative real-time RT-PCR correlated with microarray data. All samples were run in triplicate F. Immunoblotting analysis using anti MAPK-6 antibody demonstrated MAPK-6 low protein level in HMWFGF-2. Cell lysates (150 Ag) were run on SDS-9% PAGE, transferred to Immobilon-P membrane (Millipore) and analyzed using anti-MAPK-6-1 antibody as described Section 2.2. Membranes were stripped and re-probed with h-Actin antibody as loading control.

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Fig. 7. Expression profiles of Angptl-4 and Rps5 genes in cells expressing different FGF-2 forms: A. Microarray profile of Angptl-4 gene in Zip control, HMWFGF-2, LMWFGF-2 and WTFGF-2 clones showed that the gene was up-regulated in all four LMWFGF-2 clones compared to HMWFGF-2, as well as WTFGF-2 and Zip clones. B. Quantitative real-time RT-PCR. Quantified mRNA values were normalized by the amounts of Gapdh mRNA, and results are given as fold induction. Quantitative real-time RT-PCR correlated with microarray data. C. Microarray profile of Rps5 gene in Zip control, HMWFGF-2, LMWFGF-2 and WTFGF-2 clones showed that the gene was up-regulated in all four LMWFGF-2 clones compared to HMWFGF-2, as well as WTFGF-2 and Zip clone. D. Quantitative real-time RT-PCR. Quantified mRNA values were normalized by the amounts of Gapdh mRNA, and results are given as fold induction. Quantitative real-time RT-PCR correlated with microarray data. All samples were run in triplicate described in Section 2.6.

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endothelial cells undergoing G0 –G1 transition (Bouche et al., 1987). Here we show that the ribosomal protein 5 gene (Rps5) was upregulated specifically in cell expressing the LMWFGF-2 form. Interestingly, in WTFGF-2 cells both Angptl4 and Rps5 genes were not upregulated. These results suggest a ‘‘dominant-negative’’ effect of HMWFGF-2 on LMWFGF2 inducing activity, at least relatively to the expression of Angptl4 and Rsp5.

4. Discussion The goal of the work presented in this study was to examine the gene expression pattern of cells expressing different FGF-2 forms by using a microarray-based approach. As a strategy to test the potential roles of HMWFGF-2 and LMWFGF-2 forms, we selectively expressed each of these in NIH 3T3 fibroblasts, where the endogenous FGF-2 expression background is very low, but the fibroblasts can fully respond to FGF-2 ligand. We chose to express HMWFGF-2 and LMWFGF-2 selectively in NIH 3T3 cells because we reasoned that co-expression of all forms could mask specific functions of individual forms, indeed, making it difficult to identify potential unique functions of different FGF-2 forms. In our earlier study to address the functional role of different FGF-2 forms, we questioned whether the selective expression of HMWFGF-2 forms (nuclear forms) would alter the phenotype of NIH 3T3 cells. A consequence of expressing the nuclear forms alone was that the cells grew very slowly and their morphology was characterized by multinucleate giant cells (Quarto et al., 1991b). One of the ways in which cells adjust to their phenotype and their environment is by altering gene expression patterns. The development of DNA microarray technology has provided a powerful tool to measure gene expression of thousands of genes. Since comparison of gene expression patterns between cells expressing HMWFGF-2 and LMWFGF-2 may offer clues to the molecular mechanisms underlying the different phenotype, we have taken advantage of this technique to follow-up a study we started several years ago (Quarto et al., 1991b). Here we report a cDNA microarray-based study in HMWFGF-2 and LMWFGF-2 NIH 3T3 fibroblasts suggesting different potential biological function(s) of nuclear and cytoplasmic FGF-2 forms. Gene expression profiling was performed using a cDNA microarray containing 42,000 different mouse gene elements. Our results demonstrated distinct gene expression profiles in HMWFGF-2 and LMWFGF-2 expressing NIH 3T3 cells, supporting the hypothesis of distinct and unique functions of different FGF-2 forms. Among the genes showing differential expression, several were genes involved in transcription, regulatory functions, transport, energy metabolism and cell-cycle. Unidentified genes and those genes involved in signal transduction

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also contributed significantly to the differential gene expression. Interestingly, a larger proportion of the up-regulated genes in HMWFGF-2 were genes playing a role in growth arrest, while a larger portion of the down-regulated genes were ribosomal proteins. We identified genes with different levels of expression in HMWFGF-2 and LMWFGF-2 cells by direct comparison of the two data sets using SAM. We validated changes in gene expression in HMWFGF-2 and LMWFGF-2 cells by quantitative real-time RT-PCR. We also examined, for selected genes, the corresponding protein levels. Both quantitative real-time RT-PCR and protein analysis data correlated with microarray data. We observed consistent and characteristic differences in the gene-expression patterns of HMWFGF-2 and LMWFGF-2 cells. Several genes up-regulated specifically in HMWFGF-2 have been previously implicated in growth arrest, including NfI-X and Nupr1, or in tumor suppression, such as St5. We specifically examined genes that might point to signaling pathways important for the phenotype of cells expressing different FGF-2 forms. The Nuclear Factor I-X gene (NfI-X) was highly expressed in HMWFGF-2 cells. The NFI family is composed of four members in vertebrates (NFI-A, NFI-B, NFI-C, and NFI-X). Expression of NFI proteins has been associated with changes in the growth state of cells and with a number of oncogenic processes and disease states. Work by Luciakova (Luciakova et al., 2003) demonstrated that NFI-X has a direct growth arrest function in human diploid cells. Other studies have also reported changes in the levels of Nfx-X on quiescent and growth-activated NIH 3T3 cells (Goyal et al., 1990). Moreover, it has a been demonstrated that over-expression of NFI proteins (NFI-A, NFI-B, NFI-C and NFI-X) in chicken embryo fibroblasts confers resistance to transformation by nuclear oncogenes jun, fos, junD, myc and qin (Schuur et al., 1995). Nuclear protein-1 (Npr-1) was another nuclear factor upregulated in HMWFGF-2 clones. Studies performed in Nupr1 / mice demonstrated that Nupr1 is involved in cell growth regulation. Interestingly, overexpression of Npr-1 gene causes cell growth arrest, whereas its knock-out promotes growth (Vasseur et al., 2002; Baron et al., 2003). It has also been suggested that Nupr1 is a growth inhibitor that facilitates apoptosis induced in fibroblasts by DNA damage (Baron et al., 2003). Microarray analysis also revealed up-regulation of Suppressor tumorigenicity 5 (St5) in HMWFGF-2 cells. The hST5 gene, was originally identified as the HeLa tumor suppression gene (Lichy et al., 1992). Several studies reported that the expression of the human protein ST5-p70 correlates with reduced tumorigenic phenotype in mammalian cells, reverts their transformed phenotype, and restores their contact-dependent growth (Lichy et al., 1992; Hubbs et al., 1998). Furthermore, expression of ST5-p70 in COS-7 cells suppresses activation of mitogen activated protein kinase MAPK/ERK2 in response to epidermal growth factor

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stimulation (Majidi et al., 1998). Moreover, it has been suggested that ST5 can function as a signaling protein and can provide a link between c-Abl and ERK-2 (Majidi et al., 1998). Transformation related protein 53 (Trp53) also was upregulated in HMWFGF-2 cells (see Table 1). This gene acts as a tumor suppressor in many tumor types inducing growth arrest or apoptosis, depending on the physiological circumstances and cell type. Moreover, deficiency of Trp53 gene has been associated with a high frequency of spontaneous tumors development (Fleischmann et al., 2003). Mitotic arrest deficient 2 (Mad2) was another interesting gene exclusively upregulated in HMWFGF-2 clones (See Table 1). MAD2 is a spindle assembly checkpoint protein, key component of the mitotic checkpoint. It has been reported that overexpression of Mad2 in a mitotic checkpoint-defective carcinoma cell line leads to G2/M cell cycle arrest (Wang et al., 2002). In contrast, genes which promote cell proliferation and tumor progression were specifically down-regulated in HMWFGF-2 cells. The transcription factor Egr-1 was significantly down-regulated in HMWFGF-2 cells. Egr-1 expression is regulated by serum or growth factors throughout the cell cycle. Studies performed in vitro and in vivo by several groups demonstrate that inhibition of Egr1 function blocks cell proliferation and inhibits the transformed phenotype (Baron et al., 2003). Over-expression and/or mis-expression of Egr-1 induces a transformed phenotype (Virolle et al., 2003). Egr-1 is strongly down-regulated in senescent human diploid fibroblasts at the mRNA level as well as at the protein level (Meyyappan et al., 1999). Evidence for the mechanism by which Egr-1 regulates cell growth came from studies showing that a dominant-negative form of Egr-1 blocks progression of the cell cycle from late G1/S phase (Baron et al., 2003). In addition, Egr-1 is a tumor-promoting agent. Mapk-6 (Erk3) expression was also markedly decreased in cells expressing HMWFGF-2. ERK proteins are activated in response to growth factors, including FGF-2, and their activations correlate with tyrosine phophorylation. These enzymes are components of signaling pathways that control a wide variety of biological responses, including embryonic development, cell proliferation, differentiation, cell survival, and adaptation. MAPK-6 phosphorylates microtubule-associated protein-2 (map2) and may promote entry into the cell cycle. The observation that both Egr-1 and Mapk-6 were downregulated in HMWFGF-2 expressing cells is intriguing. Recent work showed that sustained extracellular signalregulated kinases 1 and 2 (ERK1 and ERK2) significantly extends the expression of Egr-1 in fibroblasts (Murphy et al., 2004). These data suggest potential relationships (functional interactions) between the two genes. Therefore, it can be hypothesized that the lack of MAPK-6 signaling accounts for the down-regulation of Egr-1.

Microarray analysis of LMWFGF-2 expressing cells showed a specific up-regulation of the angiopoietin-like 4 gene (Angptl-4). Angptl-4 is a circulating lipoprotein expressed primarily in adipose tissue and liver (Yoon et al., 2000). Moreover, Angptl-4 is a proangiogenic factor and its gene expression is induced by hypoxia in endothelial cells (Le Jan et al., 2003). When tested in the chicken chorioallantoic membrane assay, Angptl-4 induces a strong proangiogenic response, independent of VEGF. It is widely known that FGF-2 and VEGF are potent angiogenesis inducers in vivo and in vitro. A previous work by Seghezzi and colleagues (Seghezzi et al., 1998) showed that in endothelial cells, endogenous LMWFGF-2 production upregulates VEGF expression through extracellular interaction with cell membrane receptors. Our microarray data showed the novel finding that only the LMWFGF-2 form induced the expression of Angptl-4. Since experimental evidence has shown that angiogenesis is regulated by a variety of growth factors, it will be of interest to analyze whether the LMWFGF-2 form also induces the expression of Angptl-4 in endothelial cells, and furthermore, to investigate potential interactions between FGF-2 and angiopoietin-4 in the context of angiogenesis. Ribosomal protein 5 (Rps5) was also specifically upregulated in LMWFGF-2 expressing cells. An elegant work previously performed by Bouche and colleagues demonstrated that exogenously added LMWFGF-2 enters the nucleolus and stimulates the transcription of ribosomal genes in adult bovine aortic endothelial cells undergoing G0 –G1 transition (Bouche et al., 1987). In a more recent report, the same group has shown that exogenous LMWFGF-2 interacts with the free cytosolic ribosomal protein S19 (Rps19) present in NIH 3T3 cells (Soulet et al., 2001). Our microarray data showed that in LMWFGF-2 expressing cells, there was a specific induction of transcription of ribosomal gene Rps5, as well as of other ribosomal genes (see Table 2) thus, supporting data presented by Bouche and colleagues. We speculate that induction of the Rps5 gene occurs in an autocrine manner upon the release from the cell of the LMWFGF-2 form and interaction with its cell-surface receptors. In support of this hypothesis was the observation that several ribosomal genes were specifically downregulated in HMWFGF-2 expressing cells (see Table 2). The biological significance of differential expression of so many ribosomal proteins between HMWFGF-2 and LMWFGF-2 expressing cells is not clear. However, elevated expression of ribosomal protein genes has been found in human tumors and evidence for their contribution to oncogenic transformation has been reported (Loging and Reisman, 1999). Overall, the microarray data showed a differential gene expression profile between HMWFGF-2 and LMWFGF-2 forms. Interestingly, genes up-regulated in HMWFGF-2

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expressing cells were associated with growth arrest and tumor suppression activity, whereas those down-regulated were associated with early mitotic responses and signal transduction activity. In contrast up-regulation of genes associated either with growth arrest or tumor suppression was not identified in LMWFGF-2 expressing cells. These observations support a model in which the different FGF-2 forms are components of a growth/ differentiation regulatory signaling pathway, possibly involving negative and positive interaction of nuclear and cytoplasmic FGF-2 forms. Genes differentially expressed in HMWFGF-2 compared with LMWFGF-2 clones may reflect differential function of nuclear and cytoplasmic forms of FGF-2, as a result of an evolutionary selected and/or conserved function for the different FGF-2 forms. It is known that LMWFGF-2 is secreted and interacts with its cell-surface receptors, eliciting signaling which could lead to transformation (Moscatelli and Quarto, 1989; Quarto et al., 1989). Moreover, conditions such as wound healing and other stress (like, oxidative stress and heat shock) allow the release of FGF-2 ligand, which can act as an autocrine and or paracrine factor. Thresholds of ligand may be responsible for cellular transformation and malignancy. In light of our microarray data it is tempting to speculate that the endogenous HMWFGF-2 forms may counteract, possible through an intracrine-mediated pathway, the potential transforming activity of the LMWFGF2 form by expressing cell-growth arrest and tumor suppressor genes. Therefore, we propose that HMWFGF-2 (nuclear forms) and LMWFGF-2 (cytoplasmic form) may act as the ‘‘Ying-Yang’’ forms of FGF-2, and indeed, the balance between these forms may control the physiological functions of FGF-2 (Fig. 8). However, this is a hypothetical model which deserves further investigation. Our hypothesis is supported by work published by Vagner and colleagues (Vagner et al., 1996), which demonstrated that translation of CUG-initiated FGF-2 (HMWFGF-2) is activated in transformed and stressed cells (in which oxidative and/or heat-shock stress have occurred). Their work suggested that HMWFGF-2 forms are translationally induced by trans-acting factors active in transformed cells and activated as a response to stress in normal cells. Moreover, the balance between FGF-2 forms varies with the cell type and with stress conditions demonstrating the existence of this translational regulation in relation to cell transformation or stress. In addition, our recent work on calvarial osteoblasts suggested that during osteoblast differentiation, the translation of HMWFGF-2 and LMWFGF-2 forms is differentially regulated. We have demonstrated that adult and fully differentiated osteoblasts preferentially translated high levels of HMWFGF-2, while juvenile osteoblasts, which are characterized by a subpopulation of immature and highly proliferating osteoblasts, produced low levels of all three forms (Cowan et

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al., 2003). These data link the preferential expression of HMWFGF-2 to a less proliferative and more differentiated cell-state. The presence within the primary structure of HMWFGF2 proteins of an evolutionary conserved glycine –arginine (GR-motif), which is responsible for their nuclear translocation and accumulation, represents a peculiarity of HMWFGF-2 forms, relative to LMWFGF-2. Our previous study demonstrated that selective expression of HMWFGF-2 in NIH 3T3 fibroblasts confers a unique phenotype characterized by slow growth. Similar work performed later by Dono and colleagues (Dono et al., 1998) also demonstrated that when the NIH 3T3 fibroblasts expressed HMWFGF-2, or a chimeric LMWFGF-2 form in which the GR-motifs have been inserted, cells remain in growth arrest. Moreover, recent work performed in neurons overexpressing either the rat high molecular weight FGF-2 form (23 kDa FGF-2), or the N-aminoterminal portion of the 23 kDa FGF-2 form, showed morphological changes in the cell population characterized by binuclear or even multinuclear neurons (Nindl et al., 2004). Interestingly, treatment of neuronal cultures with neutralizing antibodies against FGF2 did not influence the proportion of binucleated neurons over-expressing FGF-2, indicating that the effect on nuclear morphology was not due to auto-or paracrine signaling after release of the 18 kDa or 23kDa forms into the medium (Nindl et al., 2004). Moreover, karyokinesis induced by HMWFGF-2 overexpression has also been described in non-neuronal proliferating cells like cardiac muscle (Pasumarthi et al., 1996). Furthermore, Hirst and colleagues demonstrated that high levels of HMWFGF-2 cause chromatin compaction and decreased cardiomyocyte mitosis (Hirst et al., 2003). In addition to their presence in the HMWFGF-2 forms, GR-motifs are also present in the CUG initiated FGF-3 form. Studies by Kiefer and Dickson demonstrated that this FGF-3 form localizes exclusively into the nucleus and nucleolus, and that the exclusive production of this nuclear form inhibits DNA synthesis and cell proliferation in mammary epithelial cells (Kiefer and Dickson, 2003). Taken together, these data along with our microarray data, strongly link the selective expression of nuclear FGF-2 forms to cell-growth arrest conditions. The data presented here are consistent with the hypothesis that HMW and LMW FGF-2 forms might play different biological function(s). The key difference that distinguishes HMWFGF-2 from LMWFGF-2 is the specific regulation of genes which are involved in cell growth arrest and tumor suppression. However, the role of these genes and their relationship with different FGF-2 forms remain to be elucidated. Because we can now genetically over-express or down-regulate genes, it will be possible to investigate relationships between the candidate genes identified by this microarray analysis and the cell phenotype elicited by different FGF-2 forms.

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Fig. 8. The Ying and Yang based-model for FGF-2 forms. Schemes representing a hypothetical model for the molecular mechanism of HMWFGF-2 and LMWFGF-2 forms: A. The cartoon depicts a physiological condition in which the cell translates similar amounts of FGF-2 forms. The HMWFGF-2 forms are promptly translocated to the nucleus, due to the presence within their primary structure of nuclear localization signals (NLSs) (see Bugler et al., 1991; Quarto et al., 1991a,b), while the LMWFGF-2 form remains mostly cytoplasmic. In physiological conditions, due to the lack of a canonical signal peptide, little cytoplasmic LMWFGF-2 is released from the cell. B. Conditions such as wound healing and other stress (like oxidative stress and heat shock) allow further release of FGF-2 ligands, which may fully bind to the cell-surface receptors promoting an ‘‘increased’’ autocrine signaling leading to cell transformation. C. In addition, conditions such as cell transformation or stress activate the translation of HMWFGF-2 forms, therefore by increasing their endogenous level (see Vagner et al., 1996) and disrupting the endogenous ratio between HMWFGF-2 and LMWFGF-2 forms. D. The higher level of HMWFGF-2 leads to an increased ‘‘intracrine’’ signaling which triggers the expression a ‘‘repertoire’’ of genes involved in cell-growth arrest and tumor suppression (microarray data). Therefore, the transcriptional activation of these genes might counteract the potential transforming activity of released FGF-2 ligand.

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Acknowledgments We dedicate this work to the memory of Tom Maciag. The authors wish to thank Dr. Amy Colwell for the critical reading of the manuscript. This work was supported by NIH grants R01DE13194 and R01 DE 14526, and The Oak Foundation to M.T.L.

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