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1 Dipartimento di Biologia e Patologia Cellulare e Molecolare e/o Istituto di Endocrinologia ed Oncologia Sperimentale del CNR, Facoltà di Medicina e Chirurgia di Napoli, Università degli Studi di Napoli "Federico II," Naples, Italy; 2 Kimmel Cancer Center, Jefferson Medical College, Philadelphia, Pennsylvania; 3 Department of Clinical Biochemistry, Aarhus University Hospital, Aarhus, Denmark; and 4 Istituto Dei Tumori Di Napoli "Fondazione Pascale," Naples, Italy.
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ABSTRACT |
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INTRODUCTION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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HMGA1 proteins seem to play their major physiological role during embryonic development (5) . In fact, HMGA1 expression is very high during embryogenesis, whereas it is negligible in normal adult tissues. HMGA1 proteins has been found abundant in malignant neoplasias (6) , where their expression appears critical for the acquisition of the neoplastic phenotype (7 , 8) .
To identify the differentiation pathways in which HMGA1 is involved and to assess the role of the HMGA1 proteins in development, we generated embryonic stem (ES) cells in which one or both hmga1 alleles are disrupted. We reported recently that hmga1–/– ES cells generate less T-cell precursors than do wild-type ES cells after in vitro-specific differentiation. Indeed, they preferentially differentiate to B cells, probably consequent to decreased IL-2 expression and increased IL-6 expression, both of which are regulated directly by the HMGA1 proteins (9) . Moreover, a lack of HMGA1 expression results in altered hemopoietic differentiation (i.e., there is a reduction in the monocyte/macrophage population and an increase in megakaryocyte precursors, erythropoiesis, and globin gene expression). Re-expression of the hmga1 gene in hmga1–/– ES cells restores the wild-type phenotype (9) . These results indicate that drastic changes occur in the transcriptional activity of the hmga1–/– cells, and presumably they depend on the modification of the expression of HMGA1-regulated genes.
Using the powerful oligonucleotide microarray hybridization technique, we analyzed the expression profile of ES cells carrying two, one, and no hmga1 functional alleles to identify the genes that are regulated, positively or negatively, by HMGA1. We screened an array in which 13,059 transcripts were represented, and we identified 87 transcripts that increased and 163 transcripts that decreased with a 
4-fold change in hmga1–/– ES cells with respect to the wild-type ES cells. Semiquantitative and quantitative reverse transcription (RT)-PCR confirmed the differential expression between wild-type and hmga1-knockout ES cells. We obtained different results when we measured the expression of these genes in murine embryonic fibroblasts (MEF) and various adult tissues from hmga1 knockout mice. The differential expression of some genes matched that found in ES cells, whereas the expression of other genes was either unchanged or opposite to that found in ES cells. Finally, electrophoretic mobility shift assay and chromatin immunoprecipitation experiments demonstrated that HMGA1 proteins bind to the promoters of some representative HMGA1-regulated genes, indicating a direct role of HMGA1 in the regulation of their transcription.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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RNA Extraction from Tissues and Cells.
Tissues were snap-frozen in liquid nitrogen and stored at –80°C until use. Total RNAs were extracted from tissues and cell culture using TRI REAGENT (Molecular Research Center, Inc.) solution, according to the manufacturer’s instructions. The integrity of the RNA was assessed by denaturing agarose gel electrophoresis (virtual presence of sharp 28S and 18S bands) and spectrophotometry.
Microarray Analysis.
Microarray analysis was performed as described previously in detail.5
Briefly, cRNA was prepared from 8 µg of total RNA, hybridized to MG-U74 Affymetrix oligonucleotide arrays (containing 13,059 murine transcripts), scanned, and analyzed according to Affymetrix (Santa Clara, CA) protocols. Scanned image files were visually inspected for artifacts and normalized by using GENECHIP 3.3 software (Affymetrix). Comparisons were made for each mutated sample versus wild-type sample, taking the wild-type sample as baseline by using GENECHIP 3.3. The fold-change values, indicating the relative change in the expression levels between mutated samples and the wild-type sample, were used to identify genes differentially expressed between these conditions.
Semiquantitative and Quantitative RT-PCR.
RNAs were treated with DNaseI (Invitrogen) and reverse-transcribed using random exonucleotides as primers and MuLV-reverse transcriptase (Perkin-Elmer). To ensure that RNA samples were not contaminated with DNA, negative controls were obtained by performing PCR on samples that were not reversed-transcribed but otherwise identically processed.
The PCRs were performed with the same RNAs used for array analysis, and the primers sequences are available upon request. For semiquantitative PCR, reactions were optimized for the number of cycles to ensure product intensity within the linear phase of amplification. The PCR products were separated on a 2% agarose gel, stained with ethidium bromide, and scanned using a Typhoon 9200 scanner. Digitized data were analyzed using Imagequant (Molecular Dynamics).
Quantitative PCR was performed with SYBR Green PCR Master Mix (Applied Biosystems) as follows: 95°C 10 minutes and 40 cycles (95°C 15 seconds and 60°C 1 minute).
Protein Extraction, Western Blotting, and Antibodies.
Tissues and cell culture were lysed in buffer 1% NP40, 1 mmol/L EDTA, 50 mmol/L Tris-HCl (pH 7.5), and 150 mmol/L NaCl, supplemented with complete protease inhibitors mixture (Roche Diagnostic Corp.). Total proteins were separated by SDS-PAGE and transferred to nitrocellulose membranes (Bio-Rad). Membranes were blocked with 5% nonfat milk and incubated with antibodies against Id3 and tubulin. All of them were purchased from Santa Cruz Biotechnology. Bound antibody was detected by the appropriate secondary antibody and revealed with an enhanced chemiluminescence system (Amersham-Pharmacia Biotech).
Electrophoretic Mobility-Shift Assay.
DNA-binding assays with the purified proteins were performed as described previously (10)
. Five to 20 ng of recombinant protein were incubated in the presence of radiolabeled oligonucleotide. A 200-fold excess of specific unlabeled competitor oligonucleotide was added. The double-strand oligonucleotides used were Id3 spanning from base –632 to –615 of the murine Id3 promoter region, (5'-tgattttttttttttttcaaatctg-3'; ref. 11
) and p96 spanning from base –901 to –872 of the 5' untranslated region of the murine p96 gene (5'-aagaaatatttgatattttttcttttatcc-3'; Ref. 12
).
The same oligonucleotides were also used in binding assays with total extract from wild-type and hmga1-knockout murine spleen tissues. Eight micrograms of extracts were incubated in 20 mmol/L HEPES (pH 7.6), 40 mmol/L KCl, 0.1 mmol/L EDTA, 0.5 mmol/L MgCl2, 0.5 mmol/L DTT, and 0.1 mmol/L phenylmethylsulfonyl fluoride in a volume of 20 µl containing 1 µg of poly(dC-dG), 2 µg of BSA, and 10% glycerol, for 10 minutes at room temperature. Binding reactions were incubated for 10 minutes after the addition of 2.5 fmol of a 32P-end-labeled oligonucleotide (specific activity, 8,000–20,000 cpm/fmol). For the antibody supershift analysis, the reactions were performed by preincubating extracts with 0.5 µg of antibody anti-HMGA1 (Santa Cruz Biotechnology) on ice for a minimum of 30 minutes.
The DNA-protein complexes were resolved on 6% non-denaturing acrylamide gels and visualized by exposure to autoradiographic films.
Chromatin Immunoprecipitation.
Approximately 3 x 107 wild-type and hmga1-knockout MEF were grown on 75-cm2 dishes and cross-linked by the addition of formaldehyde (to 1% final concentration) to attached cells. Cross-linking was allowed to proceed at room temperature for 5 minutes and was terminated with glycine (final concentration, 0.125 mol/L). Cells were collected and lysed in buffer containing 5 mmol/L PIPES (pH 8.0), 85 mmol/L KCl, 0.5% NP40, and protease inhibitors (1 mmol/L phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, 10 µg/ml leupeptin), on ice for 10 minutes. Nuclei were pelleted by centrifugation at 5,000 rpm for 5 minutes at 4°C and resuspended in buffer containing 50 mmol/L Tris-Cl (pH 8.1), 10 mmol/L EDTA, 1% SDS, the same protease inhibitors, and incubated on ice for 10 minutes. Chromatin was sonicated on ice to an average length of about 400 bp with a Branson sonicator model 250. Samples were centrifuged at 14,000 rpm for 10 minutes at 4°C. Chromatin was pre-cleared with protein G Sepharose (blocked previously with 1 mg/ml BSA) at 4°C for 2 hours. Pre-cleared chromatin of each sample was incubated with 2 µg of antibody anti-HMGA1 at 4°C overnight. An aliquot of wild-type sample was incubated also with anti-IgG antibody. Next, 60 µl of a 50% slurry of blocked protein G Sepharose was added, and immune complexes were recovered. The supernatants were saved as "input." Immunoprecipitates were washed twice with 2 mmol/L EDTA, 50 mmol/L Tris-Cl (pH 8.0) buffer and 4 times with 100 mmol/L Tris-Cl (pH 8.0), 500 mmol/L LiCl, 1% NP40, and 1% deoxycholic acid buffer. The antibody-bound chromatin was eluted from the beads with 200 µl of elution buffer (50 mmol/L NaHCO3, 1% SDS). Samples were incubated at 67°C for 5 hours in the presence of 10 µg RNase and NaCl to a final concentration of 0.3 mol/L to reverse formaldehyde cross-links. Samples were then precipitated with ethanol at –20°C overnight. Pellets were resuspended in 10 mmol/L Tris (pH 8)-1 mM EDTA and treated with proteinase K to a final concentration of 0.5 mg/ml at 45°C for 1 hour. DNA was extracted with phenol/chloroform/isoamyl alcohol, ethanol-precipitated, and resuspended in water. Input DNA and immunoprecipitated DNAs were analyzed by PCR for the presence of Id3 and p96 promoter sequences. PCR reactions were performed with AmpliTaq gold DNA polymerase (Perkin-Elmer). The primers used to amplify the sequence of the Id3 promoter were 5'-agggtttatgcagcaagcac-3' (forward) and 5'-atttgctgctcgtctgacct-3' (reverse). The primers used to amplify the sequence of the p96 promoter were 5'-aactccagctgtgtcaagtt-3' (forward) and 5'-gaaagaaagagaggggaaag-3' (reverse). PCR products were resolved on a 2% agarose gel, stained with ethidium bromide, and scanned using a Typhoon 9200 scanner.
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RESULTS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. Of the 13,059 transcripts represented on the array, 1,300 had a 2- to 3-, 313 had a 3- to 4-, 227 had a 4- to 10-, and 23 had a >10-fold change. We examined the 250 transcripts (1.9%) that had a 4-fold-change in the homozygous mutant versus the wild-type sample. Among these 250 transcripts, 87 were increased and 163 were decreased, including 103 known genes (37 increased and 66 decreased), 118 expression sequence tags (ESTs) (40 increased and 78 decreased), and 29 unknown genes (10 increased and 19 decreased). As a control of microarray analysis, we verified that the HMGA1 was not expressed in hmga1–/– ES cells. The genes with 4-fold change in hmga1–/– ES cells were grouped according to their function: (a) signal-transduction pathways, (b) transcription factors, (c) cell proliferation, (d) extracellular-matrix and cellular-structure proteins, (e) metabolic pathways, transport and secretion, (f) growth factors and related proteins, (g) genes with immune functions, and (h) other genes. The relative fold changes in these genes, grouped as described above, are shown in Table 1
. It is noteworthy that among the HMGA1-regulated genes, we found Id3 (13)
, lefty (14)
, and Wnt-6 (15)
that are important in embryonic development and some oncogenes such as c-myc, junB, pim-2, and c-fos.
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. The expression of some genes (i.e., TFEB, LKLF, and Id3) was hmga1 dose-dependent. In fact, the changes in hmga1+/– ES cells were intermediate between those found in wild-type and hmga1–/– ES cells. Conversely, the expression of other genes (i.e., cubilin, p96, D-9K, legumain, and collagen) was not modified in hmga1+/– ES cells in comparison to the wild-type ES cells. We also analyzed 12 genes with quantitative RT-PCR. The results coincide with those of the microarray analysis (Fig. 2B)
. The primers used for semiquantitative and quantitative PCR are available upon request.
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. We next evaluated the expression of the HMGA1-regulated genes in adult heart, liver, and spleen tissue from hmga1+/– and hmga1–/– mice.
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. Some other genes only changed in one type of tissue. For example, gly96 and LKLF expression was different only in spleen, whereas TFEB and Laminin 
1 were different only in heart. Interestingly, the regulation of some genes, such as TFEB and Laminin 1, in adult tissues was opposite to that found in ES cells. In fact, they were decreased in ES cells but increased in heart (Fig. 3A)
. These results suggest that HMGA1 function depends on the cellular context.
To verify that changes in RNA levels were associated with changes at protein levels, we analyzed by Western blot the expression of Id3 in MEF and tissues from wild-type and hmga1-knockout mice. As shown in the Fig. 3B
for the Id3 protein, protein levels paralleled RNA levels and were characteristic in each tissue.
Analysis of the HMGA1-Dependent Genes in a Transformed Cell System.
We demonstrated previously that HMGA1 overexpression is a necessary event in cell transformation. In fact, when HMGA1 expression was blocked by transfecting rat thyroid cells (FRTL-5) with an antisense hmga1 cDNA construct and infected with the Kirsten murine sarcoma virus (KiMSV) carrying the v-ras-Ki oncogene, they (FRTL-5-HMGA1as-KiMSV) did not acquire the typical markers of neoplastic transformation (ability to grow in soft agar and induce tumors after injection into athymic mice), although the differentiation markers (i.e., thyrotropin-dependency, ability to trap iodide, thyroglobulin synthesis, and secretion) were lost. Conversely, the neoplastic markers were shown by the untransfected rat thyroid cells infected with the same murine retrovirus (FRTL-5-KiMSV; ref. 8
). Therefore, we analyzed, by semiquantitative and quantitative RT-PCR, the expression of some hmga1-dependent genes in FRTL-5, FRTL-5-KiMSV, and FRTL-5-HMGA1as-KiMSV cells. The experiments revealed two sets of genes. Some genes showed the same regulation observed in the ES cells, i.e., carboxypeptidase E (Cpe) that decreased in hmga1-knockout ES cells and increased in the neoplastic cells or cathepsin H that increased in hmga1-knockout ES cells and decreased in the neoplastic cells compared with the wild-type controls. Other genes were regulated in an opposite direction compared with ES cells (i.e., p96 and mac25), which demonstrated an increased and a decreased level, respectively, in both hmga1-knockout ES and the FRTL5 KiMSV cells compared with the respective controls, although the HMGA1 proteins were expressed only in the latter cells. Some representative results are shown in Fig. 4, A and B
. In Fig. 4C
we show the expression of the proteins v-ras-Ki and HMGA1 in the normal and neoplastic thyroid cells.
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To investigate whether the HMGA1 proteins were able to bind the AT-rich promoter regions of both Id3 and p96, we performed an electrophoretic mobility shift assay using oligonucleotides spanning nucleotides –632 to –615 of the murine Id3 promoter region and –901 to –872 of the 5' untranslated region of the murine p96 gene. As shown in Fig. 5A
, a recombinant HMGA1 protein was able to bind directly to these regions. Binding specificity was demonstrated by competition experiments showing loss of binding with the addition of 200-fold molar excess of a specific unlabeled oligonucleotide. Subsequently, we performed binding assays with total extract from wild-type and hmga1-knockout murine spleens. Two specific complexes with mobility corresponding to the HMGA1 proteins (isoforms A1a and A1b) were present in extracts from wild-type and heterozygous (data not shown) spleens, whereas they were absent in extracts from homozygous hmga1-knockout mice (Fig. 5B)
. These complexes were specifically displaced by the incubation with an antibody directed against the HMGA1 proteins, demonstrating that these complexes do consist of the HMGA1 proteins (Fig. 5B)
. A control gel shift for Sp1 was performed to normalize the spleen extracts used (Panel C).
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demonstrate that HMGA1 proteins bind to these promoters. In fact, the Id3 and p96 promoter regions were amplified from the DNA recovered with anti-HMGA1 antibody in wild-type and hmga1+/– but not in hmga1–/– MEFs. Moreover, no amplification was observed in samples immunoprecipitated with normal rabbit IgG. |
DISCUSSION |
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4-fold change in hmga1–/– ES cells. The validity of these assays was confirmed by the absence of HMGA1 expression in the ES knockout cells. Semiquantitative and quantitative RT-PCR confirmed that all of these genes were differentially expressed in wild-type and hmga1-knockout ES cells. Several genes displayed hmga1 dose-dependency, the phenotype of heterozygous cells was intermediate between those of wild-type and homozygous knockout cells. Thus for some genes, the level of hmga1 expression may be critical for appropriate gene expression. In this case both alleles seem to be necessary to regulate the expression of these genes. For some other genes, the dependency on the hmga1 expression levels was even more pronounced because the gene expression level in heterozygous ES cells was very close to that observed in homozygous cells. This type of regulation by hmga1 expression levels may explain the appearance of pathologies, such as cardiac hypertrophy and B cell lymphomas, in mice heterozygous for hmga1 gene disruption.6
Several other genes showed the same expression level in wild-type and heterozygous ES cells. In this case, one hmga1 allele is sufficient to regulate gene expression. The genes regulated by HMGA1 in ES cells were also analyzed in MEF and in liver, spleen, and heart from wild-type, hmga1+/– and hmga1–/– mice. Different results were obtained in comparison to those observed in ES cells. In fact, the expression of some genes was either not modified by hmga1 gene expression, or their regulation occurred in an opposite direction. It is noteworthy that the HMGA1 regulation of several genes was cell- and tissue-specific. It is known that by interacting with partner proteins, the HMGA1 proteins are able to enhance or suppress the effect of more "traditional" transcriptional activators and repressors. The fact that partner proteins are critical for HMGA1 activity may account for the cell- and tissue-specific regulation exerted by the HMGA1 proteins.
The same occurred when the HMGA1-regulated genes in ES were investigated in a cell system constituted by normal rat thyroid cells (FRTL-5) that do not express the HMGA1 proteins, the same cells malignantly transformed by the KiMSV (FRTL-5-KiMSV) that express high-HMGA1 levels and FRTL-5-KiMSV cells in which the synthesis of the HMGA1 protein was blocked by an antisense construct (FRTL-5-HMGA1as-KiMSV). These experiments revealed two sets of genes, those showing the same kind of regulation observed in ES and those genes showing regulation that occurred in an opposite direction.
The differential gene expression depending on the HMGA1 presence could depend on an indirect effect of the HMGA1 proteins in the sense that HMGA1 might induce some proteins that may interfere with the expression of some genes. To exclude this possibility and demonstrate a direct effect of HMGA1 on the regulation of some genes expressed differentially in hmga1-knockout cells, we performed electrophoretic mobility shift assay and chromatin immunoprecipitation experiments. We demonstrated the binding of the HMGA1 proteins to the promoters of Id3 and p96. We note of particular interest the finding that p96 and Id3 are regulated by the HMGA1 proteins because they are believed to have a critical role in the process of carcinogenesis. In fact, although no putative alterations on Id genes have been identified in primary human tumors to date to certify Ids as true cellular proto-oncogenes, Id proteins that are basic helix-loop-helix transcription factors have been implicated in regulating a variety of cellular processes (i.e., cellular growth, senescence, differentiation, apoptosis, and angiogenesis) that regulate tumorigenesis (16) . In particular, Id3 has been frequently found increased in human neoplasias (16) . Equally, p96, a mitogen-responsive phosphoprotein cloned from a mouse macrophage cell expression library, is consistently down-regulated in mouse mammary carcinogenesis and in human ovarian carcinomas as compared with normal surface epithelium (17 , 18) . It is likely that Id3 up-regulation and p96 down-regulation in human neoplasias depend also on the HMGA1 overexpression, a feature of most of the human-malignant neoplasias (19) .
When we analyzed the expression of some genes in MEF and adult tissues taken from HMGA2 knockout mice, we found that several genes do not appear to be regulated by HMGA2 (data not shown). This result could depend on the different action of these two members of the same HMGA protein family and confirms that although HMGA1 and HMGA2 have a similar structure and expression profile (high during embryogenesis and neoplastic tissue), they exert different functions. This is consistent with a body of evidence indicating that the two proteins exert different function: (a) the BRCA1 promoter is regulated negatively by HMGA1 but not by HMGA2 (20) ; (b) HMGA2 is critical for adipocytic cell growth (21 , 22) , whereas HMGA1 has negative effect on the growth of the preadipocytic cells 3T3 L1 (23) ; and (c) the phenotype of the hmga1- and hmga2-knockout mice is divergent: i.e., a reduction in size and fat tissue of hmga2-null mice and in cardiac hypertrophy and B-cell lymphomas of hmga1-null mice.7
In conclusion, this study indicates that HMGA1 proteins are involved in the regulation of several genes. For some genes, such as Id3 and p96, we demonstrate that the regulation is direct. The positive or negative regulation appears to be tissue-specific because it likely depends on the multiprotein complex in which HMGA1 proteins are inserted.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Requests for reprints: Alfredo Fusco, Dipartimento di Biologia e Patologia Cellulare e Molecolare e/o Instituto di Endocrinologia ed Oncologia Sperimentale del CNR, Facoltà di Medicina e Chirurgia di Napoli, Università degli Studi di Napoli "Federico II," via Pansini 5, 80131 Napoles, Italy. Phone: 39-081-746-3749 or 3056; fax: 39-081-746-3037; E-mail: afusco{at}napoli.com
5 http://www.cancergenetics.med.ohio-state.edu/microarray. 
6 M. Fedele, V. Fidanza, S. Battista, A. Fusco, manuscript in preparation.
7 M. Fedele, V. Fidanza, S. Battista, A. Fusco, manuscript submitted for publication.
Received 4/21/04. Revised 6/17/04. Accepted 6/23/04.
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-tubulin-specific antibodies. The sources of proteins are indicated. Cpe, carboxypeptidase E.
