ARTICLE

Loss of the Tumor Suppressor PML in Human Cancers of Multiple Histologic Origins

Carmela Gurrieri, Paola Capodieci, Rosa Bernardi, Pier Paolo Scaglioni, Khedoudja Nafa, Laura J. Rush, David A. Verbel, Carlos Cordon-Cardo, Pier Paolo Pandolfi

Affiliations of authors: Molecular Biology Program and Department of Pathology (CG, RB, PPP), Department of Pathology (PC, CCC), Molecular Biology Program and Departments of Pathology and Medicine (PPS), Department of Medicine (KN), Department of Epidemiology and Biostatistics (DAV), Memorial Sloan-Kettering Cancer Center, Sloan-Kettering Division, Graduate School of Medical Sciences, Cornell University, New York, NY; Department of Veterinary Biosciences, The Ohio State University, Columbus, OH (LJR).

Correspondence to: Pier Paolo Pandolfi, MD, PhD, Molecular Biology Program and Department of Pathology, Sloan-Kettering Institute, Memorial Sloan-Kettering Cancer Center, Box 110, 1275 York Ave., New York, NY 10021 (e-mail: p-pandolfi{at}ski.mskcc.org)


    ABSTRACT
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
Background: The PML gene is fused to the RAR{alpha} gene in the vast majority of acute promyelocytic leukemias (APL) and has been implicated in the control of key tumor-suppressive pathways. However, its role in the pathogenesis of human cancers other than APL is still unclear. We therefore assessed the status and expression of the PML gene in solid tumors of multiple histologic origins. Methods: We created tumor tissue microarrays (TTMs) with samples from patients with colon adenocarcinoma (n = 109), lung carcinoma (n = 19), prostate adenocarcinoma (n = 36), breast carcinoma (n = 38), central nervous system (CNS) tumors (n = 51), germ cell tumors (n = 60), thyroid carcinoma (n = 32), adrenal cortical carcinoma (n = 12), and non-Hodgkin's lymphoma (n = 251) and from normal tissue corresponding to each histotype and analyzed PML protein and mRNA expression by immunohistochemistry and in situ hybridization, respectively. Tumor cell lines (n = 64) of various histologic origins were analyzed for PML protein and mRNA expression by immunofluorescence and northern blotting, respectively. DNA from microdissected tumor samples and cell lines was analyzed for PML mutations and loss of heterozygosity (LOH). For some tumor types, the association between PML expression and tumor stage and grade was analyzed. Statistical tests were two-sided. Results: All normal tissues expressed PML protein. PML protein expression was reduced or abolished in prostate adenocarcinomas (63% [95% confidence interval {CI} = 48% to 78%] and 28% [95% CI = 13% to 43%], respectively), colon adenocarcinomas (31% [95% CI = 22% to 40%] and 17% [95% CI = 10% to 24%]), breast carcinomas (21% [95% CI = 8% to 34%] and 31% [95% CI = 16% to 46%]), lung carcinomas (36% [95% CI = 15% to 57%] and 21% [95% = 3% to 39%]), lymphomas (14% [95% CI = 10% to 18%] and 69% [95% CI = 63% to 75%]), CNS tumors (24% [95% CI = 13% to 35%] and 49% [95% CI = 36% to 62%]), and germ cell tumors (36% [95% CI = 24% to 48%] and 48% [95% CI = 36% to 60%]) but not in thyroid or adrenal carcinomas. Loss of PML protein expression was associated with tumor progression in prostate cancer (the progression from prostatic intraepithelial neoplasia to invasive carcinoma was associated with complete PML loss; P<.001), breast cancer (complete PML loss was associated with lymph node metastasis; P = .01), and CNS tumors (complete PML loss was associated with high-grade tumors ; P = .003). PML mRNA was expressed in all tumor and cell line samples. The PML gene was rarely mutated and was not subject to LOH. Conclusions: PML protein expression is frequently lost in human cancers of various histologic origins, and its loss associates with tumor grade and progression in some tumor histotypes.



    INTRODUCTION
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
PML (formerly known as Myl) has become the object of intense research due to its involvement in the pathogenesis of acute promyelocytic leukemia (APL) (1). In the vast majority of APL patients, the PML gene (on chromosome 15) is fused to the retinoic acid receptor alpha (RAR{alpha}) gene (on chromosome 17) as a consequence of reciprocal and balanced chromosomal translocation resulting in the production of a PML–RAR{alpha} fusion protein. PML encodes a founding member of a growing family of proteins that all contain a distinctive C3HC4 zinc-binding domain termed RING finger. Some members of this family (e.g., BRCA1) have also been implicated in tumor suppression and control of genomic stability (2).

PML is typically found in multiprotein speckled subnuclear structures termed PML nuclear bodies (2,3). To date, more than 50 proteins have been reported to co-localize with PML in the nuclear body, either transiently or constitutively, including p53, pRb, Daxx, and CBP (2,3). In APL blast cells, PML–RAR{alpha} causes the de-localization of PML into microspeckled nuclear structures and the consequent disruption of the PML nuclear bodies. In Pml–/– primary cells, nuclear body components acquire an aberrant nuclear localization pattern that can be restored to normal when PML is added back (4,5). Thus, PML is essential for the formation and stability of the nuclear body. This conclusion implies, in turn, that PML may regulate the nuclear body–associated functions of multiple nuclear body components and that these functions may be impaired in APL blasts or in cells lacking PML function.

In vivo analyses have underscored the importance of the functional disruption of PML and the nuclear body in tumorigenesis. For instance, when expressed in the promyelocytic/myeloid compartment of transgenic mice, PML–RAR{alpha} caused leukemia with APL-like features (6,7). Moreover, the progressive reduction of the dose of PML obtained by crossing PML–RAR{alpha} transgenic mice with Pml–/– mice resulted in a dramatic increase in the incidence of leukemia and in an acceleration of leukemia onset (7,8). Finally, Pml–/– mice are highly susceptible to developing tumors in several in vivo models of physically or chemically induced carcinogenesis (9,10).

Until recently, little was known about the biologic and biochemical roles of PML in tumorigenesis and tumor progression. It is now becoming apparent that PML and the PML nuclear body are essential for critical tumor-suppressive pathways (9,1115). PML acts as a p53 transcriptional coactivator and is required for p53-dependent induction of apoptosis and cellular senescence upon exposure to ionizing radiations and oncogenic transformation (1215). Moreover, Pml–/– mice and cells are protected from multiple caspase-dependent apoptotic stimuli, such as Fas, tumor necrosis factor, ceramide, and interferon (9). PML is also required for transcriptional repression mediated by the tumor suppressors Mad and Rb (16,17).

Thus, at the cellular level, PML controls, at least in part as a result of its localization in the nuclear body, functions—such as induction of apoptosis, growth suppression, and cellular senescence in response to oncogenic transformation and DNA damage—that are essential for tumor suppression (11). However, it remains to be determined whether PML loss is a critical event in human tumorigenesis. We conducted a comprehensive analysis of the status of the PML gene, including its mRNA and protein expression levels, in human cancer of diverse histologic origins.


    PATIENTS AND METHODS
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
Tumors and Cell Lines

The cohort analyzed consisted of 608 patients and included patients with colon adenocarcinoma (n = 109: stage I [n = 12], stage II [n = 14], stage III [n = 24], stage IV [n = 39], liver metastasis [n = 20]), lung carcinoma (n = 19: T1 [n = 8], T2 [n = 6], T3 [n = 3], T4 [n = 2]), and prostate adenocarcinoma (n = 36: pT2 [n = 17], pT3 [n = 19]; 33 of 36 prostate cancers were multifocal, i.e., they displayed prostatic intraepithelial neoplasia [PIN] and invasive cancer lesions in different areas); breast carcinoma (n = 38: stage I [n = 4], stage II [n = 14], stage III [n = 14], stage IV [n = 6]; matching lymph node metastases were available in 25 of 38 cases), central nervous system (CNS) tumors (n = 51: grade I and II [n = 8], grade III and IV [n = 43]), germ cell tumors (n = 60: pT1 [n = 19], pT2 [n = 6], pT3 [n = 3], pT4 [n = 6]; for 26 samples, staging information was not available), thyroid carcinoma (n = 32; staging information was not available), adrenocortical carcinoma (n = 12; staging information was not available), and non-Hodgkin's lymphoma (n = 251; staging information was not available). The patients were treated and followed at Memorial Sloan-Kettering Cancer Center, and tumor samples were collected at the time of surgical resection with written informed consent. Survival data were available for patients with colon adenocarcinoma and non-Hodgkin's lymphoma. Normal tissues corresponding to all histotypes were obtained as anonymized samples of surgical specimens from patients with benign conditions who had undergone routine surgical procedures (colon, n = 6; liver, n = 8; lung, n = 3; prostate, n = 6; breast, n = 4; CNS, n = 6; thyroid, n = 3; adrenal gland, n = 4; lymph node, n = 3; skin, n = 4; tonsil, n = 4; spleen, n = 4). All specimens were obtained under Institutional Review Board–approved protocols.

We also analyzed tumor cell lines (n = 64), including 10 colon carcinoma lines (WiDr, COLO320DM, COLO320HSR, COLO205, HCT-15, SW620, LoVo, SW403, SW48, and SW948; from American Type Culture Collection, Manassas, VA), five prostate carcinoma lines (PC-3, LNCAP, DU145, TSU, BU145; from American Type Culture Collection), two germ cell tumors (NCCIT, NTERA-2; from ATCC), 10 neuroblastoma lines (LAN1, SK-N-JO, NMB7, CHIP100, LAN5, SK-N-ER, SK-N-AS, SK-N-SH, SH-SEP, 5Y5Y; all provided by Dr. A. N. Houghton, Memorial Sloan-Kettering Cancer Center and Dr. G. Melino, Department of Experimental Medicine, Biochemistry Laboratory, Istituto Dermopatico dell'Immacolata-Istituto di Ricovero e Cura a Carattere Scientifico [IDI-IRCCS], University of Rome Tor Vergata, Rome, Italy), 17 melanoma lines (SK-Mel29, SK-Mel28, SK-Mel22A, SK-Mel90, SK-Mel94, SK-Mel146, SK-Mel186, SK-Mel37, SK-Mel19, SK-Mel64, SK-Mel31, SK-MelH, SK-MelR, SK-MelD, SK-MelV, SK-MelC, SK-MelJ; all provided by Dr. A. N. Houghton), 12 multiple myeloma lines (XG1, XG2, XG4, XG5, XG6, XG7, RPMI8266, FR4, JJN3, U266, SKMM1, EJM, all provided by Dr. R. Dalla Favera, Institute for Cancer Genetics, Columbia University, New York, NY), and 10 lymphoma lines (LY1, LY7, LY8, LY17, BL29, BL36, BL54, BL74, BL113, BL136, all provided by Dr. R. Dalla Favera). All cell lines were maintained in the appropriate medium (RPMI-1640 or Dulbecco's modified Eagle medium) supplemented with 10% (vol/vol) fetal calf serum and 1% (vol/vol) penicillin–streptomycin in a humidified incubator at 37 °C and 5% CO2.

Tumor Tissue Microarray Analysis of Tissue Samples

Tissue samples were fixed in formalin and embedded in paraffin, and 5-µm sections were stained with hematoxylin–eosin to identify viable, morphologically representative areas of the specimen. Tumor tissue microarrays (TTMs) were then created as previously described (21). Briefly, triplicate tissue cores with a diameter of 0.6 mm from each specimen and normal controls were punched and arrayed on a recipient paraffin block. Sections (5 µm thick) of these tissue array blocks were cut and placed on charged polylysine-coated slides.

For immunohistochemical analysis of TTMs, we treated deparaffinized sections with 1% H2O2 to block endogenous peroxidase activity, immersed them in boiling 0.01% citric acid (pH 6.0) in a microwave oven for 15 minutes to enhance antigen retrieval, and allowed them to cool. Avidin–biotin blocking was achieved following the manufacturer's instructions for the ABC Vector Kit (Vector Laboratories, Burlingame, CA), and the TTMs were then incubated with 10% normal horse serum for 30 minutes at room temperature. A well-characterized mouse monoclonal antibody to PML (PG-M3; Santa Cruz Biotechnology, Santa Cruz, CA; 1 : 50 dilution) was then added, and the TTMs were incubated overnight at 4 °C. The TTMs were then incubated with biotinylated horse anti-mouse immunoglobulin G antibody (1 : 25 final dilution; Vector Laboratories) for 30 minutes at room temperature and then with avidin–biotin immunoperoxidase complexes (1 : 25 dilution; Vector Laboratories) for 30 minutes at room temperature. Diaminobenzidine was used as the final chromogen, and hematoxylin was used as the nuclear counterstain. Double staining was also performed on normal testis and lymph node tissue using mouse monoclonal antibodies against inhibin{alpha} (1 : 600 dilution; Serotec, Oxford, U.K.) and Bcl-6 (1 : 20 dilution; Novocastra Laboratories, Newcastle-upon-Tyne, U.K.). Diaminobenzidine and NBT/XP (Boehringer Mannheim, Mannheim, Germany), respectively, were used as chromogens.

Contiguous TTMs were also analyzed for PML mRNA expression. One microgram of recombinant plasmid pCMV-Tag2 (Stratagene, La Jolla, CA), which contains DNA coding for the RBB motif of PML [i.e., the PML RING domain followed by two distinct cysteine-rich motifs, termed B-boxes (B1 and B2) (13)], was linearized by digestion with BamHI and EcoRI. Sense (negative control) and antisense riboprobes were generated by treating linearized plasmid with T7 and T3 polymerases for 2 hours at 37 °C in 1x transcription buffer (Boehringer Mannheim, Indianapolis, IN); 20 U of RNase inhibitor; ATP, GTP, and CTP at 1 mmol/L each; UTP at 6.5 mmol/L; and digoxigenin–UTP at 3.5 mmol/L. Deparaffinized TTM sections were rinsed in water and phosphate-buffered saline for 10 minutes. The slides were digested in pre-warmed citrate buffer (10 mM citric acid, 10 mM sodium citrate) for 5 minutes in a microwave at full power. Slides were pre-hybridized for 30 minutes at 45 °C in 50% deionized formamide and 2x standard saline citrate (SSC). Hybridization was performed overnight at 45 °C in 50 µL of hybridization buffer (50% deionized formamide [vol/vol], 10% dextran sulphate, 2x SSC, 1% sodium dodecyl sulfate, and 0.25 mg/mL of herring sperm DNA) containing 10 pmol/L of digoxigenin-labeled riboprobe per section. After the hybridization solution was added, the section was covered with a coverslip to prevent evaporation. Slides were washed in pre-warmed 2x SSC for 20 minutes at 42 °C twice and then in pre-warmed 1x SSC at 42 °C for 20 minutes. The slides were then incubated in normal sheep serum diluted in buffer (2 M Tris–HCl [pH 7.5], 5 M NaCl) and successively in the same buffer with alkaline phosphatase–labeled anti-digoxigenin antibody (Boehringer Mannheim, Indianapolis, IN) at a dilution of 1 : 500 for 1 hour at room temperature. Digoxigenin was visualized by addition of the substrate nitro-blue tetrazolium 5-bromo-4-chloro-3-indolylphosphate. The slides were counterstained with methyl green and mounted (23). Two replicates of each slide were analyzed. The slides were scored for expression levels based on the presence or absence of the signal and, when the signal was present, based on the number of PML nuclear bodies.

Analysis of Cell Lines

Tumor cell lines were analyzed for protein expression using immunofluorescence and western blotting and for RNA expression using northern blotting. Indirect immunofluorescence and confocal analysis of tumor cell lines were carried out as described (4) with antibodies against PML (PGM13) and the nuclear body components SUMO-1 (FL101) and Daxx (MG112) from Santa Cruz Biotechnology. Slides were viewed on an Olympus fluorescence microscope or an Axiovert 100M confocal microscope. Western blot analysis on tumor cell lines was carried out as described (13) with anti-PML (PGM13), anti-SUMO-1 (FL101), anti-Daxx (MG112), anti-p27 (N-20), anti-{beta}-actin, and anti-heat shock protein 90 antibodies (Santa Cruz Biotechnology) and anti-cyclin D1 antibodies (DCS-6; BD Pharmingen, San Jose, CA). Immunofluorescence and western blot analyses were repeated in at least three independent experiments.

Northern blotting on tumor cell lines was carried out as described (10). In brief, total cellular RNA was extracted from cell lines using Trizol (Life Technologies, Frederick, MD), separated on a 1% agarose–formaldehyde gel, transferred to a nylon membrane (Amersham Pharmacia Biotech, Piscataway, NJ), and hybridized with a PML probe containing a BamHI/EcoRI cDNA fragment covering the PML–RBB motif. A probe for the GAPDH gene was used to control for mRNA loading. Northern blot analysis was repeated twice.

Mutation and Loss of Heterozygosity Analysis of PML

For mutation analysis, DNA was purified from microdissected tumor samples and from cell lines using the PUREGENE DNA Isolation Kit (Gentra Systems, Minneapolis, MN) according to the manufacturer's instructions. Specific areas of tumor tissues were marked on the hematoxylin–eosin-stained slides. Unstained slides were aligned by morphology to the stained slide using operating loupes (2.5x magnification), and corresponding areas were manually microdissected. DNA from 44 anonymous healthy volunteers collected at the Memorial Sloan-Kettering Cancer Center served as controls. Fourteen primer sets were used to amplify the entire PML coding sequence and intron–exon boundaries. The primers selected to produce polymerase chain reaction (PCR) products of various sizes, ranging from approximately 300 to 500 base pairs (bp), are listed in Table 1. PCR was performed in a volume of 25 µL containing 100 ng of genomic DNA and 10 pmol of each primer, using HotStarTaq Master Mix Kit (Qiagen, Valencia, CA). Initial denaturation at 95 °C for 15 minutes was followed by 35 cycles on an automated thermal cycler (GeneAmp PCR System 9700; PerkinElmer, Wellesley, MA). Each cycle included denaturation at 94 °C for 30 seconds, annealing at 60–66 °C for 30 seconds, and extension at 72 °C for 30 seconds. A final step at 72 °C for 10 minutes was added. Because optimal sensitivity for single-strand conformation polymorphism (SSCP) analysis is achieved using 100- to 400-bp DNA fragments, we digested the greater-than-400-bp PCR-amplified fragments with MboII (second fragment of exon 2 and second fragment of exon 7b) or BamHI (first fragment of exon 7b). All PCR products were then subjected to SSCP analysis using a GenePhor Electrophoresis Unit and GeneGel Excel 12.5/24 (Amersham Pharmacia Biotech) according to the manufacturer's instructions. Gels were silver-stained using the PlusOne DNA Silver Staining Kit (Amersham Pharmacia Biotech) according to the manufacturer's instructions. PCR products displaying an altered electrophoretic mobility by SSCP were re-amplified and sequenced on a semiautomated sequencer (Applied Biosystems, Foster City, CA) using the ABI PRISM Dye Terminator Cycle Sequencing-Ready Detection Kit (Perkin Elmer, Shelton, CT). Each amplicon was sequenced bidirectionally, and each sample was sequenced at least twice from independent PCR reactions.


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Table 1. Sequence of the primers used to PCR-amplify the different PML exons

 
For loss of heterozygosity (LOH) analysis, DNA was purified as described above from microdissected tumor samples and matched normal tissue. The PML gene is localized within the interval between D15S124 and D15S114 located on 15q. Four microsatellite markers (D15S124, D15S197, D15S160, and D15S114, primer sequences, and PCR conditions are provided at http://www.ncbi.nlm.nih.gov/genemap99) were PCR amplified. The PCR products were electrophoresed on a GenePhor Electrophoresis Unit using GeneGel Clean 15/24 gels (Amersham) according to the manufacturer's instructions. Gels were silver-stained as described above.

Proteasome Inhibitor Analysis

Tumor cell lines COLO320HSR, COLO320DM, SW620, NCCIT, and NTERA-2 were grown in the appropriate medium (RPMI-1640 or Dulbecco's modified Eagle medium) supplemented with 10% (vol/vol) fetal calf serum and 1% (vol/vol) penicillin–streptomycin in a humidified incubator at 37 °C and 5% CO2 in six-well plates and then incubated for 3 and 6 hours in the presence of 10 µM carbobenzoxy-L-leucyl-L-leucyl-L-leucinal (MG132; Calbiochem, San Diego, CA) diluted in dimethyl sulfoxide. The tumor cells were then harvested and analyzed for PML expression by both immunofluorescence and western blot analysis, as described above.

Statistical Analysis

PML protein expression patterns were divided into three mutually exclusive categories: no loss, partial loss, and complete loss. Complete loss was defined as undetectable levels of PML, and partial loss was defined by two or fewer PML nuclear bodies per cell. This cutoff was based on the observation that normal tissues showed, on average, 10–20 PML nuclear bodies per nucleus. The baseline variables examined for their association with PML expression patterns were tumor stage (T1–T4) in colon adenocarcinoma, tumor grade (grades I and II versus grades III and IV) in CNS tumors, PIN versus invasive tumor in prostate adenocarcinomas, and primary tumors versus lymph node metastases in breast adenocarcinomas. Fisher's exact test was used to assess the associations between PML expression and these variables. The associations between PML expression patterns and overall survival of colon cancer and non-Hodgkin's lymphoma patients were investigated by Kaplan–Meier analysis (24). The association between PML expression levels and tumor progression in prostate cancer and breast cancer was assessed using the Stuart–Maxwell test for marginal homogeneity. This test, an extension of McNemar's test (25), is useful when the number of categories in each of the variables being compared is greater than two, but equal (e.g., 3 x 3, 4 x 4). In this case, the possible values for PML expression are complete loss, partial loss, and no loss. The Stuart–Maxwell test analyzes the hypothesis of marginal homogeneity for all categories simultaneously, i.e. whether the distribution of PML expression between PIN versus primary tumors and invasive tumors versus lymph node metastases is the same. It is also appropriate because the data analyzed are considered to be paired data, being measurements taken from the same patient. Statistical correlation between CNS tumor grading and PML expression levels was assessed by the Pearson correlation coefficient (26). All P values are two-sided. The statistical analysis was performed using SAS release 8.2 (SAS Institute, Cary, NC).


    RESULTS
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
To assess the status of the PML gene, including its transcript and protein levels, we analyzed sections from TTMs (Fig. 1, A). We first compared the sensitivity of the PG-M3 antibody for immunohistochemical staining of frozen tissues (data not shown) and of the formalin-fixed, paraffin-embedded tissues on TTMs. Because we obtained similar results for the two techniques in initial analyses (data not shown), we used the fixed tissues in TTMs for subsequent analyses. We next studied expression of PML in normal tissues, including breast, colon, lung, prostate, brain, testis, lymph node, thyroid, and adrenal gland, from control individuals. All of these tissues displayed intense nuclear immunoreactivity for PML expression (Fig. 2 and data not shown). PML displayed a nuclear speckled staining pattern compatible with its normal localization in the PML nuclear body. On average, 10–20 PML nuclear bodies were detected per nucleus. In some instances, we also found a concomitant diffuse nucleoplasmic immunostaining. By contrast, in tumor specimens, PML staining was frequently partially (defined as two or fewer PML nuclear bodies per cell on average) or completely lost (Fig. 2). Endothelial cells and tumor-infiltrating lymphocytes were strongly positive for PML expression in both normal and tumor tissues, thus serving as internal positive controls.



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Fig. 1. High-throughput analysis of the PML status in human cancers. A) Normal and tumor tissues were embedded in paraffin, and 5-µm sections were stained with hematoxylin–eosin to identify viable, morphologically representative areas of the specimen. From these areas of each specimen (left, arbitrary specimen number is shown on each block), triplicate tissue cores with a diameter of 0.6 mm were punched and arrayed on a recipient paraffin block (second from left). Five-µm sections of these tissue array blocks were cut and placed on charged polylysine-coated slides (third from left) and used for morphological study and analysis of the PML mRNA and protein (right). B) The PML locus was studied by loss of heterozygosity (LOH) and single-strand conformation polymorphism (SSCP), followed by sequencing analysis of DNA from microdissected tumors negative for PML expression. C) PML is located on chromosome 15q22 and includes nine exons. Fourteen sets of primers (arrowheads) were generated and used to amplify the entire PML coding sequence and intron–exon boundaries.

 


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Fig. 2. Representative immunostaining of PML expression in tissue tumor microarrays of normal human tissues and corresponding tumor types. A) Normal colon epithelium. B) Colon adenocarcinomas. Strong nuclear immunoreactivity is displayed by connective tissue elements, including endothelial cells lining blood vessels (arrow). C) Normal testis. Double staining of PML (blue speckles) with inhibin{alpha} (brown staining reveals that PML is expressed in spermatogonia [arrow]). D) Germ cell tumors. Left panel: teratocarcinoma; right panel: seminoma. Intense nuclear immunostaining of inflammatory elements and endothelial cells is evident within the seminoma (arrow). E) Normal prostate. F) Prostatic carcinoma. Staining is evident in endothelial and other connective tissue cells (arrow). G) Normal brain. Staining is evident in neuroglial cells (arrow) and neurons (asterisk). H) Glioblastoma (arrow indicates infiltrating cells positive for PML staining). I) Normal lymph node. PML is expressed throughout germinal center differentiation as confirmed by double staining with BCL-6 (blue speckles, PML; brown staining, BCL-6), which is normally expressed in germinal center B cells. J) Lymphoma (arrow points to PML-positive endothelial cells). Original magnifications: A, H, and I, x100; C, E, G, and J, x200; B, D, and F, x400. Insets are, in all cases, higher magnifications of the images and show representative cells.

 
The overall results from the TTM analysis are summarized in Table 2. PML protein expression was partially or completely lost in prostate adenocarcinomas (63% [95% CI = 48% to 78%] and 28% [95% CI = 13% to 43%], respectively), colon adenocarcinomas (31% [95% CI = 22% to 40%] and 17% [95% CI = 10% to 24%]), breast carcinomas (21% [95% CI = 8% to 34%] and 31% [95% CI = 16% to 46%]), lung carcinomas (36% [95% CI = 15% to 57%] and 21% [95% = 3% to 39%]), lymphomas (14% [95% CI = 10% to 18%] and 69% [95% CI = 63% to 75%]), CNS tumors (24% [95% CI = 13% to 35%] and 49% [95% CI = 36% to 62%]), and germ cell tumors including teratocarcinomas (36% [95% CI = 24% to 48%] and 48% [95% CI = 36% to 60%]). As already mentioned, in each case, normal control tissue samples revealed intense PML staining. PML loss was not associated with cellular differentiation stages, as indicated by the analysis in GC tumors, in which PML protein expression was completely lost in both seminomas as well as in teratocarcinomas, which are terminally differentiated along multiple lineages (Table 2; Fig. 2, D). In some tumor types (thyroid and adrenocortical carcinomas; Table 2) no loss of PML protein expression was detected. In addition, two colon cancer, two neuroblastoma, and one germ cell tumor cell lines among those analyzed displayed partial or complete loss of PML expression (Table 2).


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Table 2. PML expression in primary tumors and cell lines*

 
We next determined whether PML status was associated with tumor progression in prostate, breast, and CNS tumors. In 33 of 36 prostate cancer samples, expression of PML was assessed in both PIN and invasive carcinoma (these 33 prostate cancers were multifocal, i.e., the same patient was carrying both PIN and invasive cancer lesions; thus, each dataset was considered to be paired). As shown in Fig. 3, A, a statistically significant association was found between complete loss of PML expression and progression from PIN to invasive carcinoma (P<.001). Similarly, analysis of matched primary tumor samples and lymph node metastases from breast cancer patients (available for 25 of 38 patients) showed a statistically significant association between complete PML loss and progression to lymph node metastases (Fig. 3, B; P = .01). In CNS tumors, PML loss was also observed in a large percentage of tumors and was more frequently associated with high-grade tumors than with low-grade tumors (stages III and IV versus stages I and II; Fig. 3, C; r2 = 0.20; P = .003). In colon cancer, 65% (95% CI = 45% to 85%) of the metastatic lesions to the liver (n = 20) but only 44% (95% CI = 34% to 54%) of primary tumors (n = 89) had lost PML expression, although this difference did not reach statistical significance. No statistically significant association was found between loss of PML expression and overall survival in non-Hodgkin's lymphoma or colon cancer patients (data not shown). Thus, PML protein expression is frequently lost in human cancers, and its loss associates with tumor progression or tumor grade in some neoplasms.



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Fig. 3. Association of loss of PML expression with tumor progression. A) PML protein expression in both prostatic intraepithelial neoplasia (PIN) and invasive prostate cancer tissues from 36 patients was determined by immunohistochemical analysis on tumor tissue microarrays. PML was found to be partially lost in 80% of PINs and in 63% of invasive carcinomas. Complete PML loss was detected in 27.5% of invasive carcinomas. A statistically significant association was found between the progression from PIN to invasive carcinoma and complete PML loss (P<.001). B) PML protein expression in breast cancer tissues and lymph node metastases from 38 patients. PML was partially or completely lost in 21% and 31% of breast cancer, respectively, and in 31% and 65% of lymph node metastases, respectively. For matched samples (i.e., the 25 patients with both breast cancer and lymph node metastasis samples), a statistically significant association was found between PML loss and lymph node metastasis (P = .01). C) PML protein expression in central nervous system (CNS) tumors from 51 patients. PML expression was partially lost in 37.5% of grade I and II and in 23% of grade III and IV CNS tumors. Complete PML loss was detected in 58% of grade III and IV CNS tumors. Statistical analysis showed that PML is frequently lost in high-grade CNS tumors (grade III and IV versus grade I and II; P = .003). Shaded bars = PML partial loss (two or fewer nuclear bodies per cell); solid bars = PML complete loss (no detectable PML expression).

 
To analyze the molecular mechanisms through which PML is lost in human cancers, we subjected contiguous TTM sections from the blocks used for immunohistochemical analyses of PML protein expression to in situ hybridization assays using sense and antisense PML probes. As expected, the PML sense probe did not reveal any signal in any of the sections analyzed (Fig. 4). By contrast, using the PML antisense probe, we detected an intense signal in all tumors analyzed, irrespective of the status of the PML protein (Fig. 4 and data not shown). Levels of PML mRNA expression varied but not in association with protein levels (e.g., higher levels of PML mRNA were often detected in tumors negative for PML protein expression; data not shown). Tumor cell lines subjected to northern blot analysis showed comparable levels of PML mRNA expression.



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Fig. 4. PML mRNA expression in human cancers. Representative PML immunohistochemistry (A and C) and in situ hybridization (B and D) of human colon carcinoma (A and B), and non-Hodgkin's lymphoma (C and D). Control in situ hybridization performed with the sense probe is shown in the upper insets. Original magnification, x200 for main panels; x400 for lower and upper insets.

 
Because PML mRNA expression was retained in samples negative for PML protein expression, we next analyzed the PML locus for mutations and LOH (Fig. 1, B and C). Mutation screening of the entire coding region and intron–exon boundaries of the PML gene was performed on DNA from both primary microdissected tumors and cell lines, including samples positive and negative for PML protein expression (Table 2). Twenty-eight lymphomas (18 primary tumors and 10 cell lines), 15 lung carcinomas (all primary tumors), 22 germ cell tumors (20 primary tumors and two cell lines), 20 prostate adenocarcinomas (all primary tumors), 17 melanomas (all cell lines), five colon cancers (all cell lines), and five neuroblastomas (all cell lines) were examined, as was DNA from 44 control individuals. Our analysis revealed several PML variants in the tumor samples. For example, among the melanoma cases, we detected a missense mutation: exon 2 (371A->G [D124G]). In addition, variants were observed in the 5' untranslated region (–14C->G) from one lung cancer patient, in intron 4 (IVS4+24G->C) from one lymphoma patient, in intron 7 (IVS7–22C->T) from one lung cancer patient, and in exon 9 (1933C->T [F645L], 1956C->T [A652A], and 2025C->T [A675A]) from several patients of all tumor histotypes analyzed. The variants identified in exon 9 probably represent polymorphisms because they were also frequently found in control individuals. PML protein expression was not affected in the melanoma, lung cancer, and lymphoma samples in which PML mutations were detected.

For the LOH analysis, matched tumor and normal DNA samples were examined. Using four different microsatellite markers (D15S124, D15S197, D15S160, and D15S114), we did not observe LOH at the PML locus in any of the tumors analyzed (data not shown), including tumors that harbored mutations in the PML gene. In addition, the tumor samples were all heterozygous for each of the microsatellite markers.

Because these results indicated that mechanisms other than inactivating mutations or LOH led to PML protein loss, we therefore assessed whether post-translational mechanisms would affect PML expression. Specifically, we investigated whether PML might be degraded through proteasome-dependent mechanisms, as has been shown for other tumor suppressors, such as p53 and p27 (27,28). For these experiments, we used cell lines representative of tumors that were found to be negative for PML protein expression. Although most cell lines expressed PML protein, some of the colon, neuroblastoma, and germ cell tumor–derived cell lines were partially or completely negative for PML protein expression (colon adenocarcinoma lines COLO320HSR and COLO320DM (Fig. 5, A), germ cell tumor line NTERA-2, and neuroblastoma lines LAN1 and SK-N-JO; Table 2). In the colon cancer and neuroblastoma cell lines, the PML protein was completely lost (i.e., it was undetectable by either western blot analysis or immunofluorescence; Table 2; Fig. 5, A and B). In the germ cell tumor line, PML protein was partially lost (i.e., two or fewer PML nuclear bodies per nucleus were detected by immunofluorescence, and low levels of protein expression were revealed by western blot; Table 2; Fig. 5, B). The cell lines defective for PML expression expressed the PML nuclear body components SUMO-1 and Daxx at levels similar to those in cell lines that were positive for PML expression (data not shown; Fig. 5, B) but with aberrant nuclear localization patterns (Fig. 5, A) consistent with the essential role of PML in the proper formation and stability of the PML nuclear body. SUMO-1 displayed a patchy nuclear distribution, and Daxx was diffused in the nucleus. As in primary tumors, in these cell lines the PML mRNA was expressed at normal levels (Fig. 5, C) and the PML gene was not mutated (data not shown).



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Fig. 5. Loss of PML protein through post-translational, proteasome-dependent mechanisms. A) Representative immunofluorescence staining of a colon cancer cell line (COLO320DM) negative for PML protein expression with anti-PML (upper and lower left panels), anti-SUMO-1 (upper central panel) and anti-Daxx antibodies (lower central panel). DAPI staining, to reveal the nucleus, is also shown. B) Western blot analysis of PML and Daxx in two colon cancer lines (COLO320DM [Co]) and SW620 [SW]) and two germ cell tumor lines (NCCIT [NC] and NTERA-2 [NT]). PML protein was not detected in COLO320DM cells and was barely detected in NTERA-2 cell lines, while Daxx, despite its abnormal nuclear localization in COLO320DM cells, was expressed at a similar level among the different cell lines. Control for protein loading and transfer using a heat shock protein 90 antibody is shown. C) Northern blot analysis of PML in the same four tumor cell lines. A GAPDH probe was used as control for mRNA loading. Asterisks indicate PML isoforms. D) Confocal immunofluorescence analysis of the expression and subnuclear localization of PML, SUMO-1, and Daxx in COLO320DM cells after a 6-hour treatment with the proteasome inhibitor MG132. E)Western blot analysis of the levels of PML in Co and SW cells after treatment with the proteasome inhibitor MG132 for 3 and 6 hours. To control for protein loading, the same blots were also stained with a {beta}-actin antibody.

 
To investigate the possible involvement of proteasome-dependent mechanisms on PML protein expression in these cells, we treated three of these cell lines (COLO320HSR, COLO320DM, and NTERA-2) and two control cell lines that express normal levels of PML (the colon carcinoma line SW620 and the germ cell tumor line NCCIT) with the proteasome inhibitor MG132. As Fig. 5, D, shows, COLO320DM cells treated with MG132 for short periods (3 or 6 hours) expressed PML in the normal speckled nuclear body localization pattern (Fig. 5, D). Western blot analysis confirmed PML accumulation in COLO320DM cells treated with MG132 (Fig. 5, E). In addition, nuclear body components such as SUMO-1 and Daxx were relocalized to the newly formed PML nuclear body. Similar results were obtained for COLO320HSR and NTERA-2 cells (data not shown). By contrast, treatment of SW620 and NCCIT cell lines with MG132 led to relocalization of PML into nucleoli-like structures (data not shown), as previously reported (29).

We carried out additional analyses to investigate the possibility that the effect of MG132 on PML was an indirect result of an effect on cell growth. Our results suggest that such indirect effects are unlikely. The cell cycle distribution in both PML-positive and PML-negative tumor cell lines was not affected by treatment with MG132 within the experimental time frame (3 and 6 hours; data not shown), suggesting that the PML accumulation in MG132-treated cells is not an indirect result of the proteasome inhibitor's effect on the cell cycle. In addition, treatment of both PML-negative and PML-positive tumor cell lines with MG132 for 6 hours increased the expression of the p27 cyclin-dependent kinase inhibitor (data not shown), as expected and as previously reported (28). By contrast, no modulation of cyclin D1 was found. These data suggest that proteasome-dependent mechanisms regulate the stability of PML.


    DISCUSSION
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
The genetics underlying the pathogenesis of leukemia and lymphoma have historically been regarded as distinct from those underlying the pathogenesis of solid tumors because hemopoietic malignancies are often associated with characteristic chromosomal translocations that are leukemia- or lymphoma-specific. However, because many of the genes involved in these translocations are also expressed in non-hemopoietic cells and control key survival and proliferation pathways, it is tempting to hypothesize that their aberrant function or loss of function may participate in the pathogenesis of non-hemopoietic tumors as well. In this study, we used a high-throughput approach to demonstrate that the protein product of the PML tumor suppressor gene of APL is frequently lost in human cancers.

Several lines of evidence have pointed to an important role for PML in tumor suppression (11). First, inactivation of PML obtained by homologous recombination provides cells of various histologic origins with a marked survival and proliferative advantage (9,13). Loss of PML function also impairs cellular senescence in response to oncogenic stimuli (14,15). In addition, PML inactivation impairs cellular differentiation upon treatment with differentiating agents such as retinoic acid and vitamin D (8,9,30). PML modulates key tumor-suppressive pathways, such as the ones controlled by p53 and Rb, which are impaired in cells lacking its function (1217). As a consequence Pml–/– mice are tumor prone (9,10). Together, these observations predicted that PML loss may represent an important step in human tumorigenesis and tumor progression. Our comprehensive analysis of the status of the PML gene and its encoded protein in human cancer provides compelling evidence that PML may participate in the pathogenesis of malignancies other than APL. Data from our study reveal that loss of PML protein is a frequent event in human cancers of various histologic origins and suggest that PML may be lost through proteasome-dependent mechanisms. PML protein expression was lost in certain tumor types and not in others (e.g., PML was never lost in thyroid carcinomas; Table 2). Furthermore, PML loss was associated with tumor progression in prostate, breast, and CNS tumors. Our data are in agreement with previous reports that documented a decrease in PML protein levels in invasive epithelial tumors (18) and loss of PML expression in a few tumors of diverse histological origins (19,20). However, none of these reports provided firm conclusions on whether PML loss was associated with a specific tumor type or grade because of the limited number of specimens analyzed.

We found that, although PML was more frequently completely lost in advanced cancers, it was also lost in early stages of tumorigenesis (e.g., in PIN; Fig. 3). This observation raises the question of whether PML loss is an important event in tumor initiation and/or progression. The ability of PML to control proliferation following oncogenic stimulation and apoptosis in cells experiencing DNA damage or other pro-apoptotic stimuli provides a straightforward explanation for how loss of PML protein would favor tumor initiation. Such an explanation is particularly likely for lymphoma and other hemopoietic malignancies, in which the survival advantage conferred by PML loss could be a critical determinant in oncogenesis. However, PML loss could also favor tumor initiation in non-hemopoietic malignancies. In this context, it is interesting to note that PML and the PML nuclear body have recently been implicated in the suppression of another important oncogenic signaling pathway that is critical for colon cancer pathogenesis: the pathway triggered by Wnt/{beta}-catenin. PIASy, a nuclear matrix–associated SUMO E3 ligase, has been found to repress the activity of the Wnt-responsive transcription factor LEF1 by sequestering it into the PML nuclear body (31). Early inactivation of PML in colon cancer could therefore lead to de-repression of this oncogenic pathway.

In addition, PML inactivation could contribute to tumor progression by favoring the accumulation of additional genetic lesions. Intriguingly, several proteins involved in the maintenance of genomic stability accumulate in the PML nuclear body, such as Bloom's syndrome protein (BLM), nibrin/p95, MRE11, and topoisomerase III{alpha} (also a BLM-interacting protein). PML could affect the function of these proteins by regulating their localization into the PML nuclear body. Support for this hypothesis comes from the finding that BLM is delocalized in Pml–/– cells as well as in APL blasts (32). Moreover, in Pml–/– mouse embryonic fibroblasts, the frequency of sister chromatid exchange, a distinctive molecular feature of the genomic instability found in cells from Bloom syndrome patients, is greatly augmented, suggesting that the localization of BLM in the PML nuclear body is required for its normal function (32).

Mutations at the PML locus have also been identified in this study. However, PML mutations were not accompanied by LOH. These mutations could lead to the generation of either dominant negative or functionally impaired PML mutants. Pml+/– primary cells grow faster than Pml+/+ cells, suggesting that PML may be haploinsufficient for some of its tumor-suppressive functions (9,10). Based on our analysis, however, mutations at the PML locus occur infrequently. Moreover, they are not detected in tumors that have lost the PML protein, suggesting that mutation is not the main mechanism of PML inactivation in the tumor types we analyzed. Our findings demonstrate that PML mRNA is expressed in virtually all the tumors analyzed and that PML protein expression can be lost through proteasome-dependent mechanisms. Post-translational loss of tumor-suppressive proteins has previously been documented. For example, the p27 cyclin-dependent kinase inhibitor has also been found lost during tumor evolution through proteasome-dependent mechanisms triggered by increased expression of the Skp2 ubiquitin ligase (33,34). As for p27, it will be important in the future to identify the ubiquitin ligase(s) that are responsible for mediating PML degradation by the proteasome because such ligase(s) could participate in tumor progression. It will also be critical to assess whether the expression of this PML ubiquitin ligase is increased in tumor cells lacking PML or whether PML degradation is triggered by the activation of major oncogenic signaling pathways.

It is also important to reconcile the infrequent complete loss of PML protein in established cancer cell lines compared with that in solid tumors of the same histotype. PML protein loss, in the context of the whole tumor, could be exacerbated by reinforcing stimuli resulting from the molecular cross-talk among tumor cells and/or the surrounding stromal component.

Although the molecular mechanisms underlying the aberrant degradation of PML in tumor cells remain to be elucidated, our findings provide a further rationale for the possible utilization of proteasome inhibitors as anticancer drugs (35). Whether loss of PML dictates response to therapy in human cancers also remains to be established. This question is of particular relevance, given that PML inactivation in mouse models renders cells resistant to the pro-apoptotic effects of ionizing radiation and chemotherapeutic agents (9,13).


    NOTES
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
C. Gurrieri and P. Capodieci contributed equally to this work.

Supported in part by Public Health Service grants CA71692 and CA74031 (to P. Paolo Pandolfi), CA47179 and CA87497 (to C. Cordon-Cardo), T32 grants CA080618 and CA009207 (to R. Bernardi and P. Paolo Scaglioni), and Spe-cialized Programs of Research Excellence (SPORE) grant 92629 (to C. Cordon-Cardo and P. Paolo Pandolfi) from the National Cancer Institute, National Institutes of Health, Department of Health and Human Services. P. Paolo Scaglioni is a recipient of the American Society for Clinical Oncology (ASCO) Young Investigator Award and of the Cancer and Leukemia Group B (CALGB) Oncology Fellow Award.

We thank Francesca Bernassola, Ilhem Guernah, Taha Merghoub, Francesco Piazza, Davide Ruggero, and Paolo Salomoni for technical help and discussions and Nai-Kong Cheung, Riccardo Dalla Favera, William Gerald, Cristina Ferrone, Cyrus Hedvat, Axel Hoos, Alan Houghton, Robert Hromas, Susan McKeinzie, Gerry Melino, Stephen Nimer, and Christoph Plass for critical material and experimental help.


    REFERENCES
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 

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Manuscript received June 9, 2003; revised December 17, 2003; accepted December 23, 2003.


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