Affiliations of authors: L. A. Koopman, G. J. Fleuren (Department of Pathology), A. R. van der Slik, M. J. Giphart (Department of Immunohematology and Bloodbank), Leiden University Medical Center, The Netherlands.
Correspondence to: Louise A. Koopman, M.Sc., Department of Pathology, L1-Q/P1-40, Leiden University Medical Center, P.O. Box 9600, 2300 RC Leiden, The Netherlands (e-mail: L.A.Koopman{at}pathology.medfac.leidenuniv.nl).
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ABSTRACT |
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INTRODUCTION |
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From studies on various cell lines (e.g., from colon, skin, and cervical tumors and from
Burkitt's lymphoma), it appears that distinct molecular mechanisms are associated with
different HLA class I phenotypes (11-13). HLA class I expression may be
abolished because of impaired ß2-microglobulin synthesis (14,15) or may be markedly reduced by a defective transporter associated with antigen
processing-dependent peptide translocation (16). Loss of an
HLA-A,-B,-C haplotype may result from loss of a chromosome 6 copy or can be caused by
deletion of a chromosomal unit embracing the HLA class I genes at 6p21.3 (4,13,17,18). Regulatory defects may cause a transcriptional modulation leading to
decreased or complete loss of expression of HLA-A or HLA-B locus gene products (3,13). The mechanisms responsible for HLA class I allelic losses, however, were
molecularly defined in only a few cases that pertained to partial genomic deletions in which the
relevant allele was involved. Browning et al. (19) identified a
chromosomal breakpoint in the HLA-A11 gene, resulting in the selective loss of the allele in
LS411, a colon carcinoma cell line. In addition, the chromosomal breaks and somatic
recombination underlying the selective loss of HLA-A2 expression were recently described in
SK-MEL-29.1.22 cells, derived from an HLA-A2-positive melanoma cell line by
-irradiation-induced mutagenesis and selection with HLA-A2 monoclonal antibody (20). Point mutations underlying allelic loss have so far been described
only in mutant cells generated by mutagenesis and immunoselection in vitro (21).
In some cell lines directly propagated from tumor lesions, the loss of expression of single alleles was found together with the presence of relevant genes, as determined by molecular typing techniques for HLA class I (22,23). Although mutations may be present in these genes, these mutations have not been demonstrated. In a previous study (13), CC11- and CSCC-7, two cervical cancer cell lines, displayed a loss of allelic expression that could be caused by mutations. In CC11- and CSCC-7 cells, which have a loss of HLA-A24 and -B15 antigen expression, respectively, the relevant genes were detected in genomic DNA (13). In the same study, we showed that the lack of HLA-A24 and HLA-B15 antigen expression persisted after interferon gamma treatment and that reduced amounts of allele-specific messenger RNA were present.
In this study, we supply the unequivocal proof that HLA class I mutations account for the loss of allelic expression in these cell lines and in (subpopulations of) their original primary tumors. Distinct nucleotide changes, each leading to a stop codon, were found in exons encoding extracellular domains of the HLA-A24 and HLA-B15 genes, thus blocking the expression of functional proteins. Such genetic alterations may represent an important escape mechanism by which tumors avoid being killed by CTLs that are restricted by the lost alleles.
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MATERIALS AND METHODS |
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Tumor cells. Informed consent was obtained from all patients whose tumors were
used. Cell line CSCC-7 was derived from a cervical squamous cell carcinoma that had lost the
expression of HLA-Bw6. Cell lines CC11- and CC11+ were
derived
from an adeno/squamous collision tumor that had lost expression of the HLA-A24 antigen in part
of its squamous component. CC11- (HLA-A24 antigen-negative) and CC11+ (HLA-A24 antigen-positive) cells were separated from the primary cell culture by
HLA-A24 antigen-specific flow cytometry sorting and were cultured as previously described (13). The allele-specific defects relevant for this study are shown in Table
1. Lymphoblastoid cell line LCL-7, an Epstein-Barr virus-transformed
peripheral blood lymphocyte line derived from the patient who was the donor for CSCC-7 cells,
served as the autologous control. CC11+ and LCL-7 cells served as the wild-type
HLA-A24 controls for CC11- cells (Table 1
).
Primary invasive carcinoma,
premalignant cervical intraepithelial neoplasia (CIN) tissue, metastatic tissue, and tumor-free
normal tissue were obtained separately by microdissection of 10-µm
hematoxylin-eosin-stained, formalin-fixed, paraffin-embedded tissue sections. A cryopreserved
sample of the tumor from which CC11+ and CC11- cells were
derived
was used for separate microdissection of HLA-A24-negative and -positive squamous carcinoma
tissue after staining with anti-HLA-A24 monoclonal antibody A11.1M (24), as previously described (13).
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DNA Isolation
DNA from primary tissues, cell lines, lymphocytes, or B-lymphoblastoid cell lines was isolated by standard protocols with proteinase K (50 µg/mL) digestion at 56 °C, phenol/chloroform extraction, or sodium chloride salting out, followed by ethanol precipitation.
Amplification Reactions
Oligonucleotide primers and biotinylated probes were locally synthesized on an ExpediteTM DNA synthesizer. Biotin (product DMT-Biotin-C6-PA; Genosys, Pampisford, U.K.) was coupled directly to the 5' end of the probe during synthesis.
HLA-B15-specific polymerase chain reaction (PCR). Primers B15F
(5'-ATGAGGTATTTCTACACCGCCA-3'; melting temperature [Tm] = 64 °C), B#31R
[5'-GCTCTGGTTGTAGTAGCC-3' (26);Tm = 56 °C] amplified an HLA-B15-specific 251-base-pair (bp) product
(Fig. 1). Primers were tailed or untailed at the 5' end with the
-21
M13 sequence (TGTAAAACGACGGCCAGT) for forward and reverse M13-based sequencing.
PCR was performed on genomic DNA, as described (26), with minor
modifications. In brief, each 100-µL reaction mixture contained 10 µL of 10x
PCR buffer (150 mM [NH4]2SO4, 500
mM Tris-HCl [pH 8.8], 0.5 mM EDTA, 15 mM MgCl2, 0.1% gelatin, and 10 mM 2-mercaptoethanol), 800 ng of DNA, all
four
deoxynucleoside triphosphates (each at 20 pmol as a mixture from Amersham Pharmacia Biotech,
Uppsala, Sweden), 300 pmol of each primer, 0.5 U Ampli-Taq polymerase (The Perkin-Elmer
Corp., Norwalk, CT), and distilled sterile H2O. After a 1-minute denaturation, the
following amplifications were done: five cycles of 25 seconds at 96 °C, 45 seconds at 70
°C, and 45 seconds at 72 °C; then 21 cycles of 25 seconds at 96 °C, 50 seconds at
65 °C, and 45 seconds at 72 °C; and finally 10 cycles of 25 seconds at 96 °C and
60 seconds at 55 °C. Final elongation incubations were for 2 minutes at 72 °C and for
10 minutes at 72 °C.
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Amplification was performed in a total volume of 100 µL containing PCR buffer (50 mM KCl, 10 mM TrisHCl [pH 8.4], bovine serum albumin [0.06 mg/mL], and 1.5 mM MgCl2), 500 ng of DNA, all four deoxynucleoside triphosphates (each at 20 pmol as a mixture from Amersham Pharmacia Biotech), 50 pmol of each primer, 0.5 U Ampli-Taq polymerase (The Perkin-Elmer Corp.), and distilled sterile H2O. The PCR profile consisted of a 5-minute denaturation at 96 °C; five cycles of 30 seconds at 96 °C, 30 seconds at 65 °C (-1 °C per cycle), and 2 minutes at 72 °C; 30 cycles of 30 seconds at 96 °C, 30 seconds at 60 °C, and 2 minutes at 72 °C; and a final 6-minute extension step at 72 °C. For the last two HLA-A24-specific reactions, elongation and final extension at 72 °C were shortened to 30 seconds and 3 minutes, respectively. Ten microliters of the PCR products was visualized on 1%-2% agarose gels, and the remaining quantity was kept for further analyses.
Sequencing Analyses
Before sequencing, PCR products were purified on Microspin S-400 HR columns (Amersham
Pharmacia Biotech). Sequencing analyses were performed with the Thermo Sequenase core
sequencing kit as described by the manufacturer (Amersham Pharmacia Biotech). For sequencing
-21 M13-amplified DNA templates, 1 pmol of Texas Red -21 M13 forward
sequencing primer was used per reaction. For each of the "nested" sequencing
reactions of the HLA-A24 gene, 1 pmol of Texas Red-labeled forward (F) or reverse (R)
oligonucleotide primer (Table 2) was used. Samples were subjected to 25
PCR cycles (each of 95 °C for 30 seconds, 60 °C for 20 seconds, and 72 °C for 20
seconds) on a PTC-200 Peltier Thermal Cycler (MJ Research, Watertown, MA). After PCR, 2
µL of loading dye was added, and samples were desiccated to reduce their volumes to
approximately 3 µL. Samples were directly loaded in a sharks-tooth comb on a 6%
Rapid-XL sequencing gel (U.S. Biochemical, Cleveland, OH), subjected to electrophoresis on a
Vistra DNA sequencer 725 (Amersham Pharmacia Biotech) at 35 W for 12 hours, and
subsequently analyzed with the Assign Version 5.0 software package (Amersham Pharmacia
Biotech). Sequences were assembled with Auto AssemblerTM DNA sequence assembly
software (The Perkin-Elmer Corp.). A consensus was built from the assembled sequences by
pairwise comparisons, with requirements set at a 10-bp minimum overlap and an error allowance
of 0%. All sequence ambiguities were validated by careful manual inspection of the gel.
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The sequence-specific biotinylated oligonucleotide probes used are shown in Fig. 3. For each dot blot hybridization, 510 µL of PCR product was
diluted with 100 µL of denaturation buffer (0.4 M NaOH and 25 mM
EDTA). Membranes (Hybond-N+; Amersham Pharmacia Biotech) were washed with distilled
sterile water and, after transfer of 100 µL of the denatured amplification products to the
membrane, hybridized as previously described (29) with minor
modifications. In brief, membranes were boiled in stripping mixture (0.5% sodium dodecyl
sulfate [SDS] in distilled water) for background reduction, prehybridized for 30
minutes at 58 °C in 5 mL of TMAC buffer (3 M tetramethylammonium chloride
[Sigma-Aldrich, Bornem, Belgium], 50 mM Tris-HCl [pH 7.5],
5
mM EDTA, and 1% SDS), and hybridized with the specific oligonucleotide probe
(1 pmol/mL of TMAC) for 1 hour at 58 °C. After being washed in standard saline
phosphate/EDTA (0.15 M NaCl, 10 mM sodium phosphate [pH
7.4], and 1 mM EDTA) containing 0.1% SDS for 10 minutes, the
membranes were treated with streptavidin-coupled horseradish peroxidase (Pierce Chemical Co.,
Rockford, IL) at 0.2 µg/mL, washed, soaked in buffer (8 M urea, 0.1 M
NaCl, 5% Triton X-100, and 1% dextran sulfate), and washed again as above. To
visualize the reaction product, the enhanced chemiluminescence kit (Amersham Pharmacia
Biotech) was used according to the manufacturer's instructions. Finally, membranes were
applied to Kodak X-Omat AR film, which was developed after the appropriate exposure time,
usually 1 minute.
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Microsatellite analysis for LOH was performed as previously described (30,31) with fluorescein-labeled microsatellite markers F13A1, D6S265, TNF-a, D6S291 for the short arm of chromosome 6 (6p), and D6S421 for the long arm of chromosome 6 (6q). Markers D6S294 (6p) and D6S1010 (6q) were used in separate 32P-based LOH analyses performed as reported (32). Genomic DNA was used for PCR amplification, followed by gel electrophoresis on an automated laser fluorescence-DNA sequencer (ALF) (Amersham Pharmacia Biotech). External-size markers in the size range appropriate for each microsatellite locus were subjected to electrophoresis with the samples. Data acquisition and quantitative analysis were done with ALF manager software and fragment manager software (Amersham Pharmacia Biotech).
Cytotoxicity Assays
A standard 51Cr-release assay was performed to measure the cytolytic activity of TILs. In brief, TIL effector cells and 100 µL of 51Cr-labeled tumor target cells (1000 cells/well) were incubated at effector-to-target cell ratios of 5 : 1 and 20 : 1 for 4 hours at 37 °C. The percentage-specific lysis was calculated as 100 x (cpm experimentally released cpm spontaneously released)/(cpm released by 10% Triton X-100 - cpm spontaneously released). All assays were carried out in triplicate. The standard deviation of triplicate samples was always less than 5% of the specific 51Cr release.
Statistical Analysis
Cytolytic activities of TILs on tumor cell targets were compared with use of the nonparametric Wilcoxon signed rank test. A two-sided P value of less than .05 was considered to be significant. All P values are two-sided.
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RESULTS |
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The failure of one of a series of HLA-B15-specific PCRs fortuitously
defined the probable site of mutation in CSCC-7 cells (Fig. 1). We
designed an HLA-B15-specific sequence reaction around codons 29-35 in
exon 2 (Fig. 1
) and amplified genomic DNA from CSCC-7 tumor cells,
lymphoblastoid control cells (LCL-7), and the original tumor tissue.
M13-based sequencing results presented in Fig. 3,
A, show the insertion
of TGGG at the last position of codon 32 in CSCC-7 cells. In the
primary tumor, the seemingly heterozygous sequence results from the
mixed presence of wild-type and inserted sequences. Presence of the
insertion in the primary tumor material was confirmed by dot blot
hybridizations that used wild-type and insert-specific oligonucleotide
probes (Fig. 3,
B). The mutated sequence was not detected in the CIN
III lesion with this method. The wild-type sequence was found in all
samples except the tumor cell line, which contained the inserted
sequence only. The four-nucleotide insertion at codon 32 in exon 2
shifts the open-reading frame throughout exon 2 and generates a TGA
stop at codon 151 in exon 3, which truncates the HLA class I protein at
the
2-domain.
Identification of the HLA-A24 Gene Defect in CC11- Cells
CC11- (HLA-A24 antigen-negative) and CC11+
(HLA-A24 antigen-positive) cells were obtained from the same
adeno/squamous collision tumor, which displayed heterogeneous HLA-A24
antigen expression in the squamous component in vivo (13). The
HLA-A74-containing haplotype, present in peripheral blood lymphocytes
from the donor patient, was absent in both CC11- and
CC11+ cells (Table 1). Microsatellite analysis of
chromosome 6 in CC11- and CC11+ cells showed
complete LOH from 6p21.3-21.2 (D6S291) to the telomere (F13A1),
whereas heterozygosity was retained at 6p12-6p11 (D6S294) (Fig. 4,
A).
This pattern suggests the presence of a chromosomal break between
D6S291 and D6S294. The same pattern is present in the primary tumor,
where imbalance values of 2.8-3.8 represent LOH (Fig.
4,
A). Thus, with the apparently normal chromosome
6 karyotype of the CC11 cell lines (Fig. 4,
B), these data imply the
presence of two duplicated and recombined copies of the short arm of
chromosome 6 (6p) carrying the HLA-A24 haplotype. Because the HLA-A24
gene was normally detected with several sets of HLA-A24-specific PCR
primers (13), the site of the possible mutation in
CC11- cells was unknown. Most of the HLA-A24 gene was,
therefore, sequenced (Fig. 2
).
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Each substitution in exons 2 and 5 had apparently occurred in only
one of the duplicated chromosome copies, accounting for the observed
heterozygosity. Because the HLA-A24 antigen was not expressed in
CC11- cells, each chromosome should carry either the exon
2 substitution or the exon 5 substitution, which would lead to proteins
truncated at the 1-domain or the transmembrane domain,
respectively (Fig. 2
). To test this hypothesis, we amplified a 1.6-kb
product that contained the sequence from the heterozygous position at
exon 2 to the heterozygous position in exon 5 by use of primers
specific for the wild-type and substituted sequence (Fig. 6,
A). Only
those reactions amplifying a sequence with a substituted nucleotide in
exon 2 and a wild-type nucleotide in exon 5 and vice versa were
positive in CC11- cells. Conversely, the reaction
amplifying wild-type nucleotides at both ends was positive only in
CC11+ cells, and the reaction that used primers specific for
substituted nucleotides at both exons was negative in both cell lines.
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To assess the presence of substitutions in exon 2 and/or exon 5 in
clinical samples available from the CC11- tumor, we first
sequenced smaller M13 PCR-amplified templates surrounding codons 89 and
279 in exons 2 and 5, respectively. With this method, the presence of
substitutions in exons 2 and 5 from CC11- cells was
confirmed, but only wild-type sequences were detected in the original
CIN and tumor lesions (data not shown). Titration experiments have
shown that initial mixtures of wild-type and mutant cells should
contain at least 5%-10% of mutant cells to detect mutant sequences
after DNA extraction and PCR (data not shown). Although the exact
proportion of tumor cells in these microdissected samples could not be
quantified, the presence of relatively large numbers of wild-type cells
in the clinical samples could have masked detection of possibly small
(<5%-10%) numbers of mutant cells by the preceding sequencing
technique. Therefore, we carried out a second independent PCR-based
approach by use of primers specific for the substituted sequences in
exons 2 and 5 (Fig. 6, B). In this experiment, substitutions in both
exons 2 and 5 were detected in the primary squamous tumor section,
which was heterogeneous for HLA-A24 antigen expression. Neither
substitution was detected in the adjacent HLA-A24-positive squamous or
adenocarcinoma tissue. The mutant-specific reactions were also negative
when DNA from the CIN lesion was tested (data not shown).
Lysis of CC11+ and CC11- by Autologous Bulk TILs
To compare the lytic activity of autologous TILs on HLA-A24 antigen-expressing CC11+ cells with the activity on HLA-A24-negative CC11- cells, we performed initial 51Cr-release cytotoxicity assays by use of autologous bulk TILs. Flow cytometric analysis with the use of two-color immunofluorescence showed that, for the CD3+ T-lymphocyte population (90% of the bulk), 50% of the cells also expressed CD8+, 30% of the cells also expressed CD4+, and 10% of the cells also expressed CD56+. The CD3+ T-lymphocyte population did not express CD16 (data not shown). Data of three cytotoxicity assays at two effector-to-target cell ratios indicate that CC11+ cells are lysed more efficiently than CC11- cells by autologous bulk TILs. At an effector-to-target cell ratio of 5 : 1, the cytolytic activity, measured as specific 51Cr release, was 9.2%, 15.8%, and 7.5% on CC11+ cells versus 0.1%, 0.4%, and 0.1%, respectively, on CC11- cells. At an effector-to-target cell of 20 : 1, the lytic activity was 16.7%, 27.0%, and 11.8% on CC11+ cells versus 1.5%, 5.1%, and 3.1%, respectively, on CC11- cells. Under these six experimental conditions, the lysis of CC11+ cells was statistically significantly higher than the lysis of CC11- cells (Wilcoxon signed rank test, two-sided P = .028).
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DISCUSSION |
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In this study, we show the occurrence of small insertions or point mutations in HLA class I
genes responsible for the loss of allelic expression in two cervical carcinomas. Initial selection of
the derivative cell lines for mutation analysis was based on previous phenotypic and genotypic
characterizations (Table 1). Insertion of the sequence TGGG in the
HLA-B15 gene of CSCC-7
cells probably resulted from the inversion of the preceding CCCA sequence (Fig. 3
). This error
and G
T and C
T substitutions in exons 2 and 5 of CC11- cells
(Fig. 5
)
generated stop codons. These changes result in truncation of the HLA-B15 polypeptide at the
2-domain and of the HLA-A24 molecule at the
1-domain and at the transmembrane
segment, respectively (Fig. 2
). This truncation is in accordance with the
diminished presence of
HLA-B15 and HLA-A24 transcripts previously demonstrated in CSCC-7 and CC11- cells, respectively (13).
The sequence variations found in the cell lines were confirmed in tissue from the original
tumors. Because the loss of HLA-A24 antigen expression in vivo was restricted to a
small portion of the squamous component in the original tumor (13), the
two substitutions in clinical samples of CC11- were detected only after
microscopic
dissection of HLA-A24 antigen-negative cells (Fig. 6, B). The
adenocarcinoma component of this tumor was homogeneously positive for HLA-A24 antigen
expression, which is consistent with the absence of mutations in this tissue or in the lymph node
metastasis from the adenocarcinoma of this patient. The selective loss of HLA-A allelic expression
despite the presence of the relevant gene has been reported in three colon carcinoma cell lines (23) and in two Burkitt's lymphoma cell lines (22). Whether HLA gene mutations have caused a loss of expression of these specific
alleles remains to be proven. One HLA class I mutation was previously identified in
patient-derived (renal) tumor material; expression of a mutant HLA-A2, resulting from a single
base change in exon 3, induced recognition by autologous CTLs (33). In a
few tumor cell lines, chromosomal break and/or somatic recombination at 6p was identified as the
molecular mechanism responsible for the loss of single HLA class I alleles (19,20). This mechanism also predominates in lymphoblastoid cells that spontaneously lose
HLA-A2, at a rate of approximately 5 x 10-6 per cell per generation,
when
cultured in vitro (34). We demonstrated similar chromosomal
breaks and recombination
at 6p that were associated with the loss of an HLA-A,-B,-C haplotype in the CC11 cells and in
primary tumor samples (Fig. 4
; Table 1
). Of
interest, the inactivation of the resulting duplicated
HLA-A24 gene copies by separate mutations resembles the biallelic inactivation described for
tumor suppressor genes. Certain alleles may be more susceptible to mutation than others. For
example, HLA-B15 and -A24 may be prone to mutations because germline mutations in these
genes have been observed in normal individuals (35,36). At present,
however, it is not known whether such mutations also occur in other alleles.
The frequency of allele-specific HLA class I loss observed in primary cervical tumor tissues
may amount to approximately 70% when adequate allele-specific monoclonal antibodies
are used (37,38). This figure takes into account that the phenotype
characterized by the loss of three alleles may result from a combination of haplotype deletion and
single allelic loss, as was found in CC11- cells (Table 1). Of the five tumors from
which we obtained cell lines, three tumors showed some type of HLA class I loss (13). In two of these tumors presented in this study, mutations were identified as the
cause for the observed loss of allelic expression. To establish whether mutations leading to allelic
loss represent a frequent phenomenon in cervical cancer or other cancers in vivo,
however, a large number of primary tumors with known HLA class I phenotypes should be
studied. Our present findings provide a direction for future research, the screening for HLA gene
mutations in cancer [e.g., by methodologies based on double-strand conformation analyses,
recently described for HLA typing (39)].
Genetic changes that affect HLA class I expression may provide clonal populations with a growth advantage. Loss of expression of HLA class I alleles allows tumor cells to escape from immunosurveillance and is regarded as evidence for immunoselection (4,5,40). The occurrence of HLA gene mutations in tumor cells may result from a combination of genetic instability and immunoselective pressure. The notion of immunoselection appears to be supported by the observation that the HLA-A24 allele was inactivated by two distinct mutations in CC11- cells. Moreover, the data obtained in preliminary cytotoxicity assays with these cells and autologous bulk TIL effector cells support the hypothesis that (loss of) HLA-A24 antigen expression by these particular tumor cells is indeed important for (escape from) recognition by autologous TILs. Although several studies (9,10,41) suggest that HLA class I loss is associated with disease progression, proof of the immunologic relevance of our data awaits the detailed study of CTL-mediated cell recognition before and after genetic reconstitution of tumor cells with the original HLA alleles. Additional experiments involving such reconstitution of HLA antigen expression by transfection, the use of clonal CTL populations, and antibody-mediated inhibition are currently in progress to further substantiate functional evidence.
Bontkes et al. (41) showed that in early-stage human papillomavirus
16-positive CIN lesions, HLA-B44, when lost, was associated with disease progression. Whether
expression of the HLA-A24 and HLA-B15 antigens was lost in the CIN lesions of both tumors in
this study could not be determined. The HLA-A24 and HLA-B15 gene mutations were, however,
not detectable in the CIN III lesions of either tumor. We cannot ascertain whether the number of
cells possibly carrying either mutation in the isolated CIN lesions was large enough to be detected
by the applied methodology. It is conceivable that distinct mechanisms of HLA loss occur at
different stages of cervical carcinogenesis. LOH at 6p has been shown to represent an early and
frequently observed event in cervical neoplasia (32,42). Indeed, the allelic
imbalance at 6p observed in the CC11 tumor and cell lines was also detected in the CIN III lesion
(Fig. 4). Three of the four HLA-B44 losses in the study by Bontkes et al. (41) might have been caused by haplotype loss secondary to LOH, but this
remains to be
studied.
In conclusion, we have defined, to our knowledge for the first time, the nature of nucleotide insertions and single-base substitutions responsible for the complete absence of HLA class I molecules in cervical cancer cells in vitro and ex vivo. Structural abnormalities like these may underlie single allelic loss observed in various tumors and may thus constitute an important structural barrier to the eradication of tumors by T-cell-based immunotherapy.
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NOTES |
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Manuscript received March 19, 1999; revised July 26, 1999; accepted August 3, 1999.
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