Human Leukocyte Antigen Class I Gene Mutations in Cervical Cancer

Louise A. Koopman, Arno R. van der Slik, Marius J. Giphart, Gert Jan Fleuren

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).


    ABSTRACT
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
BACKGROUND: Various mechanisms contribute to the loss of human leukocyte antigen (HLA) class I expression that is frequently observed in cancers. Although some single allele losses have been ascribed to mutations in HLA class I genes, direct evidence for this phenomenon in vivo is still lacking. Thus, we investigated whether HLA class I gene mutations could account for the loss of allele-specific expression in cervical carcinomas. METHODS: We used polymerase chain reaction-based techniques, including sequencing, oligonucleotide hybridization, and microsatellite analysis, to identify HLA class I gene defects in two tumor-derived cell lines and to confirm the presence of these defects in the original tumors. RESULTS: In one tumor, in exon 2 of the HLA-B15 gene, a four-nucleotide insertion resulted in a stop codon in exon 3. In the other tumor, in two duplicated copies of the HLA-A24 gene, single-point mutations resulted in stop codons in exons 2 and 5. CONCLUSIONS: To our knowledge, this is the first report of HLA class I gene mutations identified in primary tumors that lead to loss of allelic expression in tumor cells. Such tumor-specific mutations may permit the cell to escape HLA class I-restricted cytotoxic T-cell responses.



    INTRODUCTION
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
To recognize and destroy cancer cells, the cellular arm of the immune system requires that tumor epitopes are adequately presented. Polymorphic human leukocyte antigen (HLA) class I molecules function as ligands for the T-cell receptor on cytotoxic T lymphocytes (CTLs) or for inhibitory receptors on CTLs and/or natural killer cells (1). Many tumors, however, show selective loss of HLA class I expression (2,3). This may enable tumor clones with altered HLA phenotypes to selectively escape recognition by both CTLs and natural killer cells, as was demonstrated in a melanoma patient (4,5). Immunoselection is also substantiated by the increased loss of HLA class I expression on metastases from different tumor types, including cervical cancer (6-8). In addition, loss of HLA-A2 and HLA-B7 on cervical tumor cells is associated with worse prognosis and shorter survival (9,10).

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 {gamma}-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.


    MATERIALS AND METHODS
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
Cell Lines and Tissue Samples

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.Go 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 1Go). 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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Table 1. Phenotype/genotype alterations in cervical carcinoma cell lines CSCC-7, CC11-, and CC11+

 
Tumor-infiltrating lymphocytes (TILs). Bulk TILs from the primary tumor for CC11 cells were obtained, as previously described (25), with a modified interleukin 2 concentration of 30 IU/mL. RPMI-1640 medium (Life Technologies, Inc. [GIBCO BRL], Gaithersburg, MD) supplemented with glutamine (4 mM ), penicillin (50 IU/mL), streptomycin (50 µg/mL), indomethacin (1 µg/mL), leucoagglutinin (1 µg/mL), 10% human type AB serum, and interleukin 2 (30 IU), was used to expand TILs in 96-well culture plates. At each harvest of TILs for cytotoxicity experiments, 2 x 106 cells were simultaneously phenotyped with monoclonal antibodies directed against the human leukocyte differentiation antigens CD3, CD4, CD8, CD16, and CD56 by flow cytometry, as described elsewhere (25).

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. 1Go). 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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Fig. 1. Sequencing approach for human leukocyte antigen (HLA)-B15*21. Primer reactions 1-6 were as previously described (13). Primers chosen for amplification of a 251-base-pair (bp) sequencing template around codons 29-35 in exon 2 incorporate the recognition sites for the -21 M13 sequencing reaction in the forward and reverse directions. PCR = polymerase chain reaction.

 
HLA-A24-specific PCR. Two amplification reactions were carried out to provide a sequencing template covering the entire HLA-A24 gene (Fig. 2Go). The first reaction amplified a 2.3-kilobase (kb) fragment spanning the 5' flanking region to exon 4 by use of primers -501F [5'-AAGCTTACTCTCTGGCACCAA-3' (27); melting temperature [Tm] = 62 °C] and 4-53R (5'-CAGGGCCCAGCATCTCAGA-3'; Tm = 62 °C). The second reaction used primers 4-53'F (5'-ACCATGAGGCCACTCTGAGA-3'; Tm = 62 °C), located in exon 4, and A31R [5'-AACTCTTGCCTCTCAGTCCC-3' (28); Tm = 62 °C], located downstream of exon 8, yielding a 1.4-kb product.



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Fig. 2. Sequencing approach for the human leukocyte antigen (HLA)-A24 gene. Schematic representation of the HLA class I gene, including the 5' flanking region (FR) and intron-exon organization (sizes in base pairs [bp] are indicated). The approximate scale is illustrated at the bottom. The functional domains of the HLA class I molecule are mentioned above the corresponding encoding gene segments (rectangular boxes). A schematic drawing of the complete HLA class I polypeptide assembled with ß2-microglobulin (ß-2m) on a cell is shown in the square box above. Dashed arrows = primers for polymerase chain reaction amplification of HLA-A24-specific sequencing templates =; solid arrows = Texas Red-labeled sequencing primers; E = exon; i = intron; CRC = HLA class I regulatory complex; LP = leader peptide; TM = transmembrane domain; cyt. = cytoplasmic.

 
For HLA-A24-specific amplification of a 217-bp product around codon 89, the following -21 M13-tailed or untailed primers were used for forward and reverse M13-based sequencing: A24e2F (5'-AGAGAACCTGCGGATCGC-3'; Tm = 58 °C), located in exon 2, and Ai2R (5'-GGCCTAAACTGAAAATGAAAC-3'; Tm = 60 °C), located in intron 2. Similarly, a 210-bp product around codon 279 was amplified with -21 M13-tailed or untailed primers Ae4F (5'-CCATGTGCAGCATGAGGGT-3'; Tm = 60 °C), located in exon 4, and A24e5R (5'-GATGCCCACGATGGGGAC-3'; Tm = 60 °C), located in exon 5.

Amplification was performed in a total volume of 100 µL containing PCR buffer (50 mM KCl, 10 mM Tris–HCl [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 2Go) 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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Table 2. Texas Red-labeled oligonucleotide primers used for human leukocyte antigen-A sequencing*

 
Oligonucleotide Dot Blot Hybridization

The sequence-specific biotinylated oligonucleotide probes used are shown in Fig. 3.Go For each dot blot hybridization, 5–10 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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Fig. 3. Insertion of four nucleotides in human leukocyte antigen (HLA)-B15*21 is detected in cell line CSCC-7 and in a primary tumor but not in a cervical intraepithelial neoplasia (CIN) III lesion. A) Sequence (codons 30-35) is read from the bottom to the top of the gel (arrow). Heterozygous positions in the primary tumor sample are denoted as follows: K = G or T; S = C or G; and M = A or C. wt = wild-type; ins = four-nucleotide insertion. B) Biotinylated(circled B) oligonucleotide probes specific for the wild-type sequence (B*1521 WT) or the insertion (B*1521 INS) at codon 32 were hybridized to polymerase chain reaction-amplified products of DNA from primary tumor or tumor-free (normal) tissue, autologous lymphoblastoid control cells (LCL-7), tumor cell line CSCC-7, and a CIN III lesion. The inserted nucleotides are underlined.

 
Loss of Heterozygosity (LOH) Analysis

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.


    RESULTS
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
Identification of the HLA-B15 Gene Defect in CSCC-7 Cells and Primary Tumors

The failure of one of a series of HLA-B15-specific PCRs fortuitously defined the probable site of mutation in CSCC-7 cells (Fig. 1Go). We designed an HLA-B15-specific sequence reaction around codons 29-35 in exon 2 (Fig. 1Go) 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,Go 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,Go 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 {alpha}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 1Go). 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,Go 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,Go A). Thus, with the apparently normal chromosome 6 karyotype of the CC11 cell lines (Fig. 4,Go 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. 2Go).




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Fig. 4. A) Schematic representation of chromosome 6 with relative positions of human leukocyte antigen (HLA) class I genes and microsatellite markers (46) used in CC11+ and CC11- cell lines and clinical samples. Illustrative measurements for two markers on 6p and 6q, respectively, are shown to the right. Numeric results for each marker represent ratios calculated by dividing the ratio of the allelic peak heights (p1/p2) measured for the normal sample (N) by the ratio of peak heights measured for test samples. The inverse ratio is taken for values less then 1. Complete loss of one of the two alleles represents true loss of heterozygosity (LOH), and ratios of 1.3 or less represent retention of heterozygosity (ROH). Ratios of 1.7 or more represent allelic imbalance (a.i.) (47). Prim.T = primary tumor; meta = metastasis; sm = size marker; nt = not tested; CIN = cervical intraepithelial neoplasia. B) Chromosome 6 karyogram. Standard cytogenetic analysis was performed on CpG-banded chromosomes in metaphase spreads of CC11- and CC11+ cells. The karyotype of both cell lines showed two chromosome 6 copies.

 
The HLA-A24 gene revealed complete sequence identity for the 2650-bp sequence from the flanking region through intron 5 in both cell lines, with two exceptions, shown in Fig. 5.Go In the HLA-A24-negative cell line CC11-, de novo heterozygosity was observed at two distinct nucleotide positions, one at codon 89 (GAG in exon 2) and the other at codon 279 (CAG in exon 5). In both cases, the codon's first wild-type base was replaced with thymine, resulting in a TAG stop codon.



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Fig. 5. Substitutions in exons 2 and 5 in CC11+ and CC11- cells. Sequencing with Texas Red-labeled sequencing primers (SP) on the polymerase chain reaction-amplified templates was performed with several overlapping primers (Fig. 4Go). Sequence ambiguities between CC11- and CC11+ cells were validated if at least three sequencing primer reactions showed the same result. Forward reactions are presented for SP2 (Fig. 2Go) in exon 2 and SP7 (Fig. 2Go) in exon 5. Sequence gels are read from bottom to top (arrows). Heterozygous positions in CC11- cells at exons 2 and 5 are denoted as K = G or T and Y = C or T (underlined). The substitutions result in heterozygous presence of TAG stop codons at residues 89 and 279.

 
Assignment of the HLA-A24 Exon 2/Exon 5 Mutations to Different Chromosome 6 Copies

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 {alpha}1-domain or the transmembrane domain, respectively (Fig. 2Go). 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,Go 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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Fig. 6. A) Each chromosome 6 copy in CC11- cells exclusively contains a substitution in either exon 2 or 5 of the human leukocyte antigen (HLA)-A24. Primers specific for the wild-type (wt) and substituted (subst) sequences in exon 2 (e2) and in exon 5 (e5) were used in different combinations to amplify a 1.6-kilobase (kB) product spanning the region between both heterozygous positions in exons 2 (forward, F) and 5 (reverse, R). Polymerase chain reaction (PCR) A used primers e2-wt-F (5'-CGCTACTACAACCAGAGCG-3'; melting temperature [Tm] = 62 °C) and e5-wt-R (5'-GATGGGGACGGTGGC*CTG-3'; Tm = 62 °C; * = G -> C mismatch deliberately introduced to increase 3' wt specificity), PCR B used primers e2-subst-F (5'-CGCTACTACAACCAGAGCT-3'; Tm = 58 °C) and e5-wt-R. PCR C applied e2-wt-F and e5-subst-R (5'-GATGGGGACGGTGGGCTA-3'; Tm = 60 °C). In D, e2-subst-F was used with e5-subst-R. B) Both substitutions are detected in primary tumor tissue. wt and/or substitution-containing sequences in exons 2 and 5 were amplified from DNA obtained from cell lines and clinical tissue samples. A fraction of the squamous carcinoma component (scc) of the fresh-frozen tumor was microdissected on the basis of HLA-A24 antigen heterogeneity (+/-) observed in this small segment by anti-HLA-A24 monoclonal antibody staining, as reported elsewhere (13). + = HLA-A24 positive. Not shown are the negative results for the following clinical samples: adenocarcinoma component, a cervical intraepithelial neoplasia III lesion, and a metastatic (adenocarcinoma) lesion. PCR I (with primers e2-wt-F and Ai2R) and PCR II (with primers e2-subst-F and Ai2R) amplified a 156-base-pair (bp) product; PCR III (with primers Aex4F and e5-wt-R) and PCR IV (with primers Aex4F and e5-subst-R) amplified a 160-bp product. Ai2R and Aex4F are described in the text. The PCR profiles were the same as those described for A24-specific amplification on behalf of M13-based sequencing. A 10-µL sample of product was applied to 1.5% agarose gel. sm = size marker Phix 174 RF DNA HaeIII digest (New England Biolabs, Beverly, MA).

 
Analysis of the HLA-A24 Exon 2/Exon 5 Mutations in CC11- Primary Lesions

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,Go 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).


    DISCUSSION
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
The identification of mechanisms that contribute to the frequent loss of HLA class I expression in tumors is central to our understanding of immune escape and to the development of suitable immunotherapeutic strategies. Structural genetic alterations, in contrast to regulatory defects, may not be overcome by the appropriate immunomodulatory stimuli. In the multistep process of tumorigenesis, LOH or chromosomal rearrangements may involve the HLA region on 6p (20). Smaller mutations may also target HLA class I genes and thus lead to loss of allelic expression, but to our knowledge this has not been demonstrated directly in primary tumors.

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 1Go). 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. 3Go). This error and G -> T and C -> T substitutions in exons 2 and 5 of CC11- cells (Fig. 5Go) generated stop codons. These changes result in truncation of the HLA-B15 polypeptide at the {alpha}2-domain and of the HLA-A24 molecule at the {alpha}1-domain and at the transmembrane segment, respectively (Fig. 2Go). 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,Go 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. 4Go; Table 1Go). 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 1Go). 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. 4Go). 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.


    NOTES
 
We thank Jacqueline Anholts for technical assistance, S. L. van Zelderen-Bhola for cell line karyotyping, and Professor Els Goulmy for critically reviewing the manuscript.


    REFERENCES
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 

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Manuscript received March 19, 1999; revised July 26, 1999; accepted August 3, 1999.


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