Affiliations of authors: C. A. Thrash-Bingham, Fox Chase Cancer Center, Philadelphia, PA; K. D. Tartof, Urologic Oncology Branch, Division of Clinical Sciences, National Cancer Institute, Bethesda, MD.
Correspondence to: Kenneth D. Tartof, Ph.D., National Institutes of Health, Bldg. 10, Rm. 2B47, Bethesda, MD 20892-1501 (e-mail: kdtartof{at} helix.nih.gov).
K. D. Tartof would like to dedicate this work to Dr. Alfred Knudson on the occasion of the 25th anniversary of his two-hit hypothesis and for making this research possible. We thank Dr. Donald Chapman and C. Strobe for their expertise and equipment used in the hypoxia experiments and B. Bingham for comments on statistical analyses.
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ABSTRACT |
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INTRODUCTION |
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pHIF1 not only functions as a hypoxia transcription factor but
also interacts with p53 during the hypoxic response. It has been known
for some time that hypoxic cells accumulate p53 (17).
Recently, it has been shown that, in response to hypoxia,
pHIF1
binds to p53 and protects it from proteolytic degradation,
thereby facilitating its accumulation under conditions of low oxygen
(18). Moreover, it was further demonstrated that, in the
absence of pHIF1
, p53 accumulates in hypoxic cells only when a
proteosome inhibitor is present. This finding suggests that, by
associating with pHIF1
during hypoxia, p53 is protected from
proteolytic degradation via the proteosome pathway.
The rapid response to hypoxia reflects the fact that this condition is a potentially dangerous one that most cells experience only for brief periods of time. However, it has been estimated that perhaps half of all tumors are hypoxic or are composed of hypoxic regions within the malignant growth. As tumors become hypoxic, they may overexpress the genes discussed above in an attempt to influence tumor vascularization (19). From a practical point of view, hypoxic cells pose a particular problem for the management of malignant disease because they are especially resistant to virtually all forms of therapy.
Renal carcinomas are among those malignant diseases whose rate of occurrence is increasing and for which effective nonsurgical treatments are lacking, especially for late stage disease. In general, these tumors may be broadly assigned to one of two histologic patterns of growth, nonpapillary (85% of cases) or papillary (10% of cases). Nonpapillary disease may be further subdivided according to cell type, being either clear cell, granular cell, mixed clear and granular cell, and, more rarely, sarcomatoid. In both nonpapillary and papillary renal cancers, malignant cells arise from the epithelium of the proximal renal tubule.
Nonpapillary renal carcinoma is a distinct morphologic and molecular disease entity that is quite separate from papillary renal carcinoma. Nonpapillary tumors are characterized by dysfunction of the tumor suppressor gene VHL (von Hippel-Lindau), whereas papillary kidney cancers are not (20,21). Moreover, the hereditary form of basophilic papillary renal tumors possesses mutations in the MET proto-oncogene, while such alterations are not present in nonpapillary renal disease (22).
The Von Hipple-Lindau (VHL) locus, located at chromosome 3p25, has been demonstrated to be an early-acting tumor suppressor gene that is either mutated or silenced in most cases of sporadic nonpapillary clear-cell renal carcinomas (23,24). Normally, the VHL protein (pVHL) associates with several other proteins that regulate transcription (25) and control the cell cycle (26). In addition and for reasons unknown, cells in which pVHL is defective also overexpress VEGF, PDGFB, and GLUT1, genes that are controlled by several factors, including hypoxia. It has been shown that the increase in VEGF is due to stabilization of its messenger RNA (mRNA) (11,27), and its increased production by clear-cell renal carcinomas is probably responsible for the highly vascularized stroma characteristic of nonpapillary tumors. In fact, introduction of a functional VHL gene into clear-cell carcinomas corrects the misregulation of these loci under normal aerobic (normoxic) conditions and restores a typical hypoxia-inducible response (11). Therefore, these results suggest that pVHL may affect the post-transcriptional stability of mRNAs involved in the transduction of signals that detect oxygen levels.
In this article, we demonstrate the presence of a natural antisense
transcript, referred to here as aHIF (antisense HIF1), that is
complementary to sequences in the 3' untranslated region (UTR) of
HIF1
mRNA and is strikingly and specifically overexpressed in
nonpapillary clear-cell renal carcinomas. Because HIF1
is of
central importance for the hypoxic response, we have characterized aHIF
in normal and malignant cells.
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MATERIALS AND METHODS |
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Samples of normal and tumor renal tissues were obtained from patients, with no prior exposure to chemotherapy, who underwent surgery for kidney cancer at the Fox Chase Cancer Center. Loss-of-heterozygosity studies on most of these tumors have been described previously (28). Re-examination of the pathologic specimen from patient RCC34 indicates that it should be classified as a papillary renal carcinoma. Conditions for the establishment and maintenance of renal tumor cell lines and blood-derived lymphocytes immortalized with Epstein-Barr virus (EBV) are described elsewhere (29). Total RNA was extracted from 50 mg of tissue or from cultured cells with the use of an RNeasy Isolation Kit (Qiagen, Valencia, CA). After the first extraction, all samples were treated with deoxyribonuclease (DNase) and repurified on a second Qiagen column according to the manufacturer's instructions. This second purification was necessary to eliminate all traces of DNA contamination.
Lymphocytes (106/mL) obtained from patient RCC22 and immortalized with EBV were maintained in glass vessels at 37 °C; they were mixed by a magnetic stirrer and flushed with a water-saturated gas mixture (5% CO2/specified amount of O2/N2) at a rate of 1 L/minute. At the end of the incubation period, cells were immediately chilled on ice, and RNA was extracted as described above.
Oligonucleotides
The oligonucleotides used in reverse transcription-polymerase chain
reaction (RT-PCR) experiments reported here are listed in Table
1. They (RT-PCR) either were obtained from
commercial sources as indicated or were produced on a PE Applied
Biosystems (Foster City, CA) DNA synthesizer.
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Differential display reactions were carried out by use of the RNAimage Kit (GenHunter, Nashville, TN). Briefly, complementary DNAs (cDNAs) were synthesized from 200 ng of total RNA by use of Moloney murine leukemia virus (MMLV) reverse transcriptase and oligo-deoxythymidylate (oligo-dT)-based primers H-T11A, H-T11G, or H-T11C in a 20-µL volume according to conditions recommended by the supplier. Two microliters of the reaction products was amplified by PCR by use of the same oligo-dT primer and one of the following short arbitrary sequence primers: H-AP9, H-AP10, H-AP11, H-AP12, and H-AP13. Samples were subjected to electrophoresis on 5% denaturing Long Ranger acrylamide gels (FMC BioProducts, Rockland, ME). After autoradiography, bands of interest were excised from the gels and DNA was eluted. DNA fragments were re-amplified by use of the same primer pair employed in the original differential display amplification and sequenced by use of the Sequenase PCR Sequencing Kit (Amersham Pharmacia Biotech, Piscataway, NJ).
Semiquantitative RT-PCR
These reactions were performed according to published procedures
(30-32) as follows: Total RNA (200 ng) was reverse
transcribed in 20-µL reactions with 1 mM each of the four
standard deoxyribonucleotide triphosphates and 2.5 µM each
of gene-specific reverse primers for aHIF, HIF1, and ACTB
(ß-actin; Table 2)
by use of MMLV reverse
transcriptase (GenHunter) for 1 hour at 37 °C. PCR
amplification of each gene product was carried out in parallel
20-µL reactions to avoid depletion of substrates. Each tube
contained 4 µL of the RT reaction, 1 µM of forward and
reverse gene-specific primers, 30 µCi
[
-32P]deoxyadenosine triphosphate (dATP), and 2.5 U
Taq polymerase (Perkin-Elmer Applied Biosystems, Foster City,
CA). Amplifications were carried out for 16 cycles for ACTB, 22 cycles
for HIF1
, and 26 cycles for aHIF. These cycle numbers were chosen
because they are within the exponentially increasing range with a
reaction efficiency of 1.9 to 2.1 (31) and allow comparable
amounts of [
-32P]dATP incorporation into each product.
Equal volumes from each PCR amplification were loaded onto denaturing
acrylamide gels and subjected to electrophoresis for 2 hours. After
visualization by autoradiography, the DNA bands were excised from the
gel, and radioactive incorporation was determined by use of a
scintillation spectrometer. For the comparison of the amount of
amplified aHIF and HIF1
fragments produced from different RNA
samples, the amplified ACTB product of each sample was used as an
internal standard. The radioactivity incorporated into aHIF and
HIF1
fragments of each RNA sample was divided by the radioactivity
incorporated into the ACTB fragment produced by that same sample, and a
ratio was then calculated for each tumor/normal pair.
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RNA from an EBV-transformed lymphocyte cell line obtained from patient RCC39 served as template for use with a 5' and 3' RACE (rapid amplification of cDNA ends) Systems Kit (Life Technologies, Inc. [GIBCO BRL], Gaithersburg, MD). Two types of 5' RACE experiments were performed with reverse transcriptase and gene-specific primer KT482, followed by either one or two nested PCR reactions. The first nested PCR used the Anchor primer and KT481. The second reaction utilized the Universal primer and KT477. For 3' RACE, an oligo-dT Adapter primer (Life Technologies. Inc.) was employed to synthesize cDNAs, and primers KT476 and KT480 were used in nested PCR amplification reactions with the oligo-dT primer. Amplifications were carried out as recommended by the supplier by use of aHIF gene-specific primer KT482 for cDNA synthesis and primers KT481 and KT477 for nested reactions in conjunction with the Anchor and Universal amplification primers. PCR products were cloned into the vector pCR2.1 (Invitrogen Corp., Carlsbad, CA). Cloned DNAs were cycle sequenced by use of an Applied Biosystems sequencer.
Ribonuclease Protection Assays
A 1049-base-pair (bp) aHIF fragment was PCR amplified from 5' RACE clone 3-7 by use of primers KT485 and KT489, designed to eliminate the oligo-C tail created by the RACE protocol. This fragment was cloned into pCR2.1 in both orientations. BamHI linearized plasmid served as substrate for riboprobe synthesis in a reaction containing 300-400 ng of DNA; 500 mM ATP, guanosine triphosphate, and cytosine triphosphate; 12 µM uridine triphosphate (UTP); 50 µCi [32P]UTP; 10 mM dithiothreitol; 40 U RNAsin inhibitor (Promega Corp., Madison, WI); 20 U T7 RNA polymerase; and 1x transcription buffer (Promega Corp.). Transcription products were separated on 5% denaturing Long Ranger gels. Gel slices containing full-length transcripts were excised and incubated overnight at 4 °C in gel elution buffer (Ambion, Austin, TX). Ribonuclease (RNase) protection assays were performed by use of a HybSpeed RPA Kit (Ambion). Typically, 40 µg of total RNA isolated from the RCC22 tumor cell line was mixed with 7 x 104 cpm of riboprobe labeled to a specific activity of about 109 cpm/µg. Conditions of probe hybridization and RNase T1 digestion were as recommended by the supplier (Ambion). The resulting protected fragments were separated on a 5% acrylamide gel and visualized by autoradiography. RNA size markers were synthesized by in vitro transcription of Century markers (Ambion) in the presence of [32P]UTP as recommended by the supplier.
Statistics
For the comparison of differences in expression of aHIF and
HIF1 transcripts between tumor types, radioactive counts from
excised bands were normalized against counts for ACTB (ß-actin)
amplified in the same experiment. RT-PCR reactions for all samples
obtained from the patients were performed in duplicate. Data were
expressed as ratios of normalized tumor cpm divided by adjusted normal
cpm from the same patient. The data were analyzed by two-sided nested
analysis of variance (Nested ANOVA) (33,34).
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RESULTS |
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To search for genes whose expression might be specifically
associated with nonpapillary clear-cell renal carcinoma, we examined
RNAs from five normal and nonpapillary clear-cell tumor pairs as well
as from one renal oncocytoma normal/tumor pair in duplicate by
differential display (35). As a result of these experiments,
we discovered a distinct transcript that is strikingly overexpressed
only in clear-cell tumors relative to normal kidney tissue and not
expressed in either normal or tumor tissue from the renal oncocytoma
(Fig. 1). No other transcript detected in this
experiment was similarly overexpressed.
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First, primer-specific RT-PCR amplification demonstrates
overexpression of an aHIF transcript in clear-cell tumors. Forward and
reverse oligonucleotide primers capable of amplifying a 92 nt fragment
contained within the 187 nt UTR were synthesized (Table 1).
DNase-treated RNA from clear-cell renal tumor RCC33 and corresponding
normal kidney tissue were reverse transcribed in separate reactions by
use of either the reverse or the forward primer. The resulting cDNAs
were then PCR amplified in the presence of both primers and
[32P]dATP, and the products were separated by acrylamide
gel electrophoresis. As illustrated in Fig. 2,
RT
reactions with the reverse primer yielded similar amounts of
HIF1
-derived product in both normal and tumor tissues. In
contrast, RT with the forward primers produced a conspicuous
aHIF-derived fragment only with tumor RNA. These HIF1
and aHIF
products were not due to spurious DNA contamination because PCR
amplification of RT reactions lacking reverse transcriptase
(RT- lanes) failed to produce HIF1
or aHIF fragments.
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Finally, an RNase protection experiment demonstrates that aHIF is an
antisense transcript derived from the opposite strand of the 3' UTR
of HIF1. As illustrated in Fig. 4
, a cDNA obtained from the RACE
experiments containing the last 1049 bp of the aHIF transcript
(see "Materials and Methods" section) was cloned, in
both orientations, adjacent to a T7 promoter. The full-length in
vitro synthesized riboprobe is 1163 nt, owing to an additional 114
nt derived from plasmid sequences flanking the insert. HIF1
and
aHIF transcripts are expected to overlap with 737 nt and 1049 nt of
their complementary riboprobes, respectively. Full-length
32P-labeled riboprobes complementary to HIF1
and aHIF
were synthesized in vitro, purified on an acrylamide gel, and
then hybridized to total RNA isolated from RCC22, a cell line
established from a clear-cell renal carcinoma (28). Following
RNase T1 digestion, the RNA duplexes were denatured and subjected to
electrophoresis on an acrylamide gel.
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aHIF Overexpression in Clear-Cell Renal Tumors
Semiquantitative RT-PCR was used to assess the relative abundance
of aHIF and HIF1 transcripts with respect to ACTB in various renal
tumors. The efficiency of RT-PCR amplification of aHIF, HIF1
, and
ACTB transcripts was found to be 2.09 ± 0.02 (mean ± standard
deviation), 2.06 ± 0.03, and 1.97 ± 0.05 per cycle, respectively,
over the cycle numbers used here. The PCR cycle numbers chosen for the
assay of these products in the experiments described below are within
the exponentially increasing range of the reactions and permit
comparable amounts of [
-32P]dATP incorporation without
exhausting the substrate.
RT-PCR experiments using primers within the unique 5' domain of
aHIF demonstrated that this antisense transcript was specifically
overexpressed in all nonpapillary clear-cell renal cell carcinomas
examined. RNA from each of 10 different primary clear-cell tumors and
the corresponding normal tissues was reverse transcribed in a reaction
containing the reverse primers for aHIF, HIF1, and ACTB
transcripts. The resulting cDNAs were PCR amplified in three separate
reactions using primers specific for each transcript, and the
synthesized fragments were examined by acrylamide gel electrophoresis.
As illustrated by the four representative examples presented in Fig.
5,
A, aHIF was strikingly overexpressed in all
nonpapillary carcinomas in comparison with normal kidney tissue,
whereas the abundance of HIF1
and ACTB transcripts in these same
specimens was relatively constant. In contrast, papillary renal
carcinomas (Fig. 5
, B) displayed no consistent differential elevation
of either aHIF, HIF1
, or ACTB products.
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Overexpression and Hypoxic Induction of aHIF in Renal Carcinoma Cells and Lymphocytes
Because aHIF is associated with the HIF1 locus, we wanted to
determine if the aHIF transcript could be induced by hypoxia.
Therefore, RCC22, one of our established nonpapillarly renal carcinoma
cell lines, was exposed to varying oxygen conditions. Cells were
equilibrated in a normal oxygen atmosphere (21%) for 1 hour and
then in 21%, 0.3%, 0.1%, and 0% oxygen for 2 hours.
After RNA purification, the samples were assayed by RT-PCR for the
presence of aHIF and ACTB transcripts. As shown in Fig. 6
, A, the
expression of aHIF remained elevated and at about the same levels,
whether in a normoxic or in a hypoxic atmosphere. The amount of ACTB
remained relatively constant under these conditions as well.
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Because the amount of aHIF was measured relative to ACTB in the cell being assayed, it was not possible by this method to compare the absolute levels of aHIF in RCC22 and lymphocytes. The difference in aHIF/ACTB ratios in RCC22 and lymphocytes might be due to differences in the amounts of aHIF, ACTB, or both. Nevertheless, it is clear that RCC22 cells express aHIF at high levels in normoxic and hypoxic conditions, whereas lymphocytes express high levels of aHIF only in a hypoxic environment. It is possible that aHIF is not further induced by hypoxia in RCC22 because it is already maximally expressed in normoxic conditions.
Two other cell lines were also assayed for possible inducibility of aHIF by hypoxia. aHIF remains very low and not induced by hypoxia in Hep3B (human hepatocarcinoma) cells and HeLa (human ovarian carcinoma) cells (data not shown). The reason for this is not known. It may be an intrinsic property of these cell types, or it may the reflect changes that such cell lines have undergone over the very long time they have been maintained in culture that have impaired aHIF expression. The pattern of expression of aHIF in a variety of mammalian cells and tissues is being investigated.
For the determination of the effect of hypoxia on aHIF, HIF1, and
ACTB over time, lymphocytes were equilibrated in an atmosphere of
21% for 1 hour and then shifted to 0.1% oxygen. At appropriate
intervals, aliquots of cells were removed, RNA was extracted, and the
presence of aHIF, HIF1
, and ACTB RNA was assayed by RT-PCR. As
shown in Fig. 6
, C, induction of aHIF began at about 2 hours and
increased over the next 6.5 hours. During that period, the amount of
ACTB remained virtually constant. Quantitation of the radioactivity
incorporated in each band is graphically displayed in the lower portion
of Fig. 6
, C. The results indicate that, by 6.5 hours of hypoxia, aHIF
increased about eightfold while HIF1
appeared to be reduced about
twofold. This inverse relationship suggests that aHIF expression may
negatively affect the abundance of HIF1
mRNA.
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DISCUSSION |
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It is known that, in at least 80% of all sporadic nonpapillary clear-cell renal carcinomas, there is loss of VHL function as a result of mutation or gene silencing (21,24). This situation indicates that loss of VHL function is a very early event in the development of nonpapillary renal cancer. Moreover, considerable molecular data demonstrate that pVHL mediates a post-transcriptionally controlled increase in the expression of a number of loci associated with oxygen stress, including VEGF, GLUT1, and PDGFB (11,27). The overexpression of aHIF is consistent with this pattern. Hence, it will be of interest to determine if VHL acts through a transcriptional or post-transcriptional mechanism to cause the increased expression of aHIF RNA.
In the nonpapillary renal carcinoma cell line RCC22, hypoxia fails to
increase further the already elevated expression of aHIF. This result
suggests that the aHIF transcript may be maximally expressed in these
cells. However, in lymphocytes, aHIF is present at low levels under
normoxic conditions but is readily inducible by hypoxia. This
inducibility implies that aHIF may play a regulatory role in the
hypoxic response. In this regard, it is of interest to note that the
increase in aHIF and the decline in HIF1 occur at a time (about 4
hours) when pHIF1
begins to decrease (2). Thus, aHIF may
facilitate the reduced expression of pHIF1
during periods of
prolonged hypoxia. It is also worth noting that the reciprocal
regulation of the sense transcript by expression of its counter
transcript has been reported for several genes (36-40). Assay
of pHIF1
in conjunction with the expression of aHIF and HIF1
transcripts will help to decide this issue.
The potential for aHIF to reduce HIF1 abundance could have
important implications for the regulation of p53. Increased aHIF
expression could lead to a decrease in the amount of pHIF1
and,
therefore, a lower abundance of p53. Because p53 acts as a tumor
suppressor, loss of p53 function results in deregulated cell
proliferation (41-43). An interesting example of how the
level of p53 can be altered to achieve oncogenic transformation
involves the MDM2 locus. MDM2 binds p53 and accelerates its degradation
through the ubiquitin pathway (44,45). Of particular interest
is the fact that MDM2 is amplified in more than one third of 47 human
sarcomas examined (46). It has been proposed that, in this
subgroup of cancers, loss of p53 function occurs not by mutation of p53
but rather by enhancing its degradation. Experiments to determine the
effect of aHIF expression on p53 and pHIF1
are under way.
Much remains to be discovered about the aHIF transcript itself. Because
the aHIF sequence reported here is incomplete, it is not known if it is
linked to a protein-coding region or possesses other regulatory
information. Also, we do not know the sequences regulating aHIF
expression, although a hypoxia response element would appear to be
implicated. If aHIF forms a duplex structure with the 3' UTR of
HIF1 mRNA in vivo, this might affect the abundance of the
HIF1
transcript or the protein it encodes. In addition, such a
putative duplex might interact with proteins such as
double-stranded RNA adenosine deaminase-editing enzymes
(47-49), double-stranded RNA-activated protein kinase
(50), interferon-induced RNase L (51), or
inosine-specific
RNase (52). Experiments are under way to address these points.
The overlap between aHIF and HIF1 transcripts is substantial,
corresponding to 882 bp of the 1169 bp in the 3' UTR of HIF1
mRNA. It is of interest to note that the HIF1
mouse and human
3' UTRs are about 89% identical over their entire length
(53). This high degree of noncoding sequence conservation is
relatively common. Recent comparisons of vertebrate genes included in
the Genbank database reveal that about 5%-30% have highly
conserved 3' UTRs (54,55). Such extensive homology
throughout a noncoding sequence implies that there is strong selective
pressure acting on the entire region that has an important or essential
role in the cell (54).
An intriguing hypothesis to explain the strong conservation in 3' UTRs such as this proposes that long, nearly perfect RNA duplexes are formed between sense and antisense transcripts and function as a signal for the stabilization or destruction of the target mRNA (56). Although there may be some small nt sequences that bind regulatory proteins within or near these duplexed regions, it is the long length of the duplex per se that is the signaling element. Sequence conservation over the entire length of 3' UTR is maintained because the presence of mutations would, in the heterozygote, disrupt the duplex.
The overexpression of a natural antisense transcript in clear-cell renal carcinoma suggests a novel regulatory mechanism that may be of importance for understanding the control of gene expression in normal cells as well as in several diseases, including cancer. The system reported here provides a means to explore this phenomenon further.
Supported by grants from the Lucille P. Markey Charitable Trust; Public Health Service grant CA06927 from the National Cancer Institute, National Institutes of Health, Department of Health and Human Services; a gift from the Tuttleman Family Foundation in tribute to Dr. Michael Kriegler; and an appropriation from the Commonwealth of Pennsylvania.
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Manuscript received June 18, 1998; revised October 21, 1998; accepted November 10, 1998.
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