Somatic Mutation in Human T-Cell Leukemia Virus Type 1 Provirus and Flanking Cellular Sequences During Clonal Expansion In Vivo

Franck Mortreux, India Leclercq, Anne-Sophie Gabet, Arnaud Leroy, Eric Westhof, Antoine Gessain, Simon Wain-Hobson, Eric Wattel

Affiliations of authors: F. Mortreux, I. Leclercq, Unité 524 Institut National de la Santé et de la Recherche Médicale (INSERM), Institut de Recherche sur le Cancer de Lille, and Unité d'Oncogenèse Virale, Centre Oscar Lambret, Lille, France; A.-S. Gabet, E. Wattel, Unité 524 INSERM, Institut de Recherche sur le Cancer de Lille, Unité d'Oncogenèse Virale, Centre Oscar Lambret, and Unité d'Oncogenèse Virale, UMR5537 Centre National de la Recherche Scientifique (CNRS)-Université Claude Bernard, Centre Léon Bérard, Lyon, France; A. Leroy, Unité d'Oncogenèse Virale, Centre Oscar Lambret; E. Westhof, Institut de Biologie Moleculaire et Cellulaire-CNRS, Strasbourg, France; A. Gessain (Unité d'Epidémiologie des Virus Oncogènes), S. Wain-Hobson (Unité de Retrovirologie Moléculaire), Institut Pasteur, Paris, France.

Correspondence to: Eric Wattel, M.D., Ph.D., Unité d'Oncogenèse Virale, UMR5537-CNRS-Université Claude Bernard, Centre Léon Bérard, 28, rue Laënnec 69373 Lyon cedex 08, France (e-mail: wattel{at}lyon.fnclcc.fr).


    ABSTRACT
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 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Background: Human T-cell leukemia virus type 1 (HTLV-1), the causative agent of adult T-cell leukemia/lymphoma, shows intrapatient genetic variability. Although HTLV-1 can replicate via the reverse transcription of virion RNA to a double-stranded DNA provirus (the conventional manner for retroviruses), its predominant mode of replication is via the clonal expansion (mitosis) of the infected cell. This expansion is achieved by the viral oncoprotein Tax, which keeps the infected CD4 T lymphocyte cycling. Because Tax also interferes with cellular DNA repair pathways, we investigated whether somatic mutations of the provirus that occur during the division of infected cells could account for HTLV-1 genetic variability. Methods: An inverse polymerase chain reaction strategy was designed to distinguish somatic mutations from reverse transcription-associated substitutions. This strategy allows the proviral sequences to be isolated together with flanking cellular sequences. Using this method, we sequenced 208 HTLV-1 provirus 3' segments, together with their integration sites, belonging to 29 distinct circulating cellular clones from infected individuals. Results: For 60% of the clones, 8%–80% of infected cells harbored a mutated HTLV-1 provirus, without evidence of reverse transcription-associated mutations. Mutations within flanking cellular sequences were also identified at a frequency of 2.8 x 10-4 substitution per base pair. Some of these clones carried multiple discrete substitutions or deletions, indicating progressive accumulation of mutations during clonal expansion. The overall frequency of somatic mutations increased with the degree of proliferation of infected T cells. Conclusions: These data indicate that, in vivo, HTLV-1 variation results mainly from postintegration events that consist of somatic mutations of the proviral sequence occurring during clonal expansion. The finding of substitutions in flanking sequences suggests that somatic mutations occurring after integration, presumably coupled with selection, help move the cellular clones toward a transformed phenotype, of which adult T-cell leukemia/lymphoma is the end point.



    INTRODUCTION
 Top
 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Human T-cell leukemia virus type 1 (HTLV-1), the first human pathogenic retrovirus isolated, is the etiologic agent of a malignant CD4 T lymphoproliferation (adult T-cell leukemia/lymphoma [ATLL]) (1,2) and of a chronic progressive neuromyelopathy (tropical spastic paraparesis [TSP]/HTLV-1-associated myelopathy [HAM]) (3,4). Furthermore, this virus has been associated, although to a lesser extent, with the development of a variety of inflammatory diseases (510). Among the 15–25 million individuals infected worldwide, approximately 3%–5% will develop ATLL, depending on as yet unknown cofactors. Proviral loads during the asymptomatic phase of infection are generally lower than in TSP/HAM, although they can be as high as one infected cell in 25 peripheral blood mononuclear cells (PBMCs) (11) The remarkable genetic stability of HTLV-1 led to the suggestion that the virus replicated in concert with cell mitosis (12). This hypothesis was confirmed for all cases of HTLV-1 infection, notably asymptomatic carriers, patients with TSP/HAM, and patients with ATLL (13). In the latter case, tumor cell clones were identified in a general background of oligoclonal or polyclonal expansion of infected but nontransformed cells (14). This mode of replication results from the effects of the viral protein Tax on T-cell proliferation. Tax protein induces the expression of numerous genes involved in the differentiation and the proliferation of T cells (15,16). Furthermore, through the functional inactivation of P16INKA4 (17) and the activation of cyclin D2 (18) and cyclin D3 (19), Tax intervenes directly in the pathway controlling T-cell proliferation.

In addition to its positive effect on cell cycling, the Tax protein negatively interferes with some DNA repair functions of the host cells. Indeed, Tax represses the expression of the human {beta} polymerase gene (20) and disrupts other prominent cellular DNA repair pathways (21,22). Via p53, Tax influences the transition from G1 to S phase and impairs the DNA-damage sentinel at this junction (23). Furthermore, Tax functionally inhibits the human mitotic checkpoint protein Mad1 (24). These functional characteristics of Tax may explain its mutagenic effect on cellular chromosomal DNA, as evidenced in vitro (25).

Considered together, these data indicate that HTLV-1 replicates mainly through persistent host cell proliferation (26) in the context of a Tax-induced genetic instability that should be detectable in vivo. In this study, we attempt to investigate the genetic variability of HTLV-1 that might be resulting from a previously undescribed mechanism and to ascertain whether the intrapatient genetic variability of the 3' RU5 region of the long terminal repeats (LTRs) is a consequence of somatic mutations in the proviral sequence rather than of reverse transcription. Mutations within flanking cellular sequences, including one integration site that corresponded to the {alpha}-enolase gene (27), a housekeeping gene, were also observed in both symptomatic and asymptomatic infected individuals.


    MATERIALS AND METHODS
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 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Samples

The DNA samples from PBMCs of six HTLV-1-seropositive individuals were studied: two patients with TSP/HAM (P1 and P2), two asymptomatic carriers (P3 and P4), and two patients with ATLL (P5 and P6). The age and sex of the subjects were as follows: P1, a 40-year-old female; P2, a 46-year-old female; P3, a 32-year-old female; P4, a 57-year-old female; P5, a 55-year-old male; and P6, a 40-year-old male. The DNA samples from PBMCs from five HTLV-1-seronegative individuals were used as the negative controls. According to French law, the six HTLV-1-infected individuals and the five seronegative subjects gave their informed consent before the collection of the blood samples.

Molecular Detection and Semiquantitative Analysis of HTLV-1 Integration

Inverse polymerase chain reaction (PCR) analysis of the HTLV-1 integration sites was done for the detection of infected cellular clones (13,14,26,28,29). This method consists of the amplification of the 3' extremities of the proviruses together with their flanking sequences. It allows the detection of circulating clones of HTLV-1-bearing T cells. In addition, as detailed in the "Results" section, sequencing cloned inverse PCR products makes it possible to arrange proviral sequences into cellular clones according to their cellular flanking sequences. Accordingly, for a given sample, aligning the proviral sequences flanked by distinct integration sites allows for the detection of reverse transcription-associated substitutions. By contrast, comparing proviral sequences flanked by identical integration sites helps to detect somatic mutations.

Because there is a stochastic component to the detection of HTLV-1 integration sites within the range of 100–1000 copies per clone with the use of inverse PCR, samples from the six individuals were analyzed in quadruplicate (28). In this study, the last 399 base pairs (bp) of HTLV-1 3'-LTR, together with the cellular flanking sequences, were amplified.

Two micrograms of DNA was digested by 20 U NlaIII (New England Biolabs, Montigny-Le-Bretonneux, France) in 1x NlaIII buffer for 3 hours at 37 °C. The completion of the digestion was controlled by 1% agarose gel electrophoresis. DNA was extracted with phenol–chloroform (1 : 1) and precipitated with 100% ethanol. One microgram of digested DNA was circularized for 14 hours at 16 °C with 20 U of T4 DNA ligase (New England Biolabs) in 600 µL of 1x T4 DNA ligase buffer and 1 mM adenosine triphosphate. DNA was extracted with phenol–chloroform (1 : 1) and precipitated with 100% ethanol. Samples were analyzed through quadruplicate experiments: 4 x 500 ng of circularized DNA was amplified for 30 cycles with the use of 200 µM of primer pair BIO6 (5'-CTCCTGCTAGTTTATTGAGCCATA-3') at position 8621–8598 and LTR1 (5'-TCGCATCTCTCCTTCACGCG-3') at position 8657–8675 (nucleotide coordinates are numbered according to the HTLV-1 reference sequence on ATK-1) (30). Amplifications were performed with the use of the proofreading Pfu DNA polymerase (Stratagene Cloning Systems, La Jolla, CA), which has one of the lowest error rate (1.3 x 10-6 error per base per duplication) (31). Amplifications were performed according to the instructions of the enzyme manufacturers. Thermal cycling parameters were as follows: 96 °C for 10 minutes and 30 times at 96 °C for 60 seconds, 58 °C for 60 seconds, and 72 °C for 3 minutes, followed by a final elongation step of 10 minutes at 72 °C. Quadruplicate inverse PCR analysis of a DNA sample from patient P5 allowed the detection of a dominant clone on a background of less abundant forms, whereas five abundant clones were detected in the DNA sample from patient P6 (see the "Results" section). To detect the most abundant clone within this sample, we performed inverse PCR with 250, 100, 150, 10, 6.6, 1, and 0.1 ng of tumor DNA from patient P6 that was diluted in DNA from PBMCs of a noninfected individual.

Runoff Analysis of Amplified Products

The length polymorphism generated by PCR amplification of HTLV-1 flanking sequences was analyzed by making a runoff, as described previously (13,14,26,28). This method consists of the linear PCR amplification of both the 3' extremity of the provirus and its flanking sequence. Two microliters of amplified product was submitted to 10 cycles of linear PCR with 2 µM of 5'-32P-radiolabeled primer BIO5 (5'-TGGCTCGGAGCCAGCGACAGCCCAT-3') (position 8995–9020), 1 U of the Stoffel fragment of the Taq DNA polymerase (Perkin-Elmer Applied Biosystems, Courtaboeuf, France), and 200 µM of each deoxynucleoside triphosphate in a final volume of 20 µL. The thermal cycling parameters were as follows: 95 °C for 10 minutes and 10 times at 95 °C for 60 seconds, 58 °C for 60 seconds, and 72 °C for 3 minutes, followed by a final elongation step of 10 minutes at 72 °C. After boiling in deionized water, 2 µL of runoff products was analyzed on a 6% sequencing gel.

Southern Blot Analysis

Approximately 10 µg of high-molecular-weight DNA originating from PBMCs from patients P5 and P6 was digested with EcoRI or PstI and then subjected to electrophoresis through a 0.6% agarose gel. After Southern blotting to a nylon membrane, the filter was hybridized with the randomly primed 32P-labeled PMT-2–3 probe, which corresponds to the 1.7-kilobase (kb) fragment of the gag-pol region obtained after digestion of the MT2 cell line DNA by PstI.

Cloning and Sequencing HTLV-1 3' RU5 Sequences Together With Their Integration Site

Purified products from inverse PCR experiments were phosphorylated by the T4 polynucleotide kinase (Pharmacia, Uppsala, Sweden) and then ligated with the SmaI-digested (Pharmacia) and dephosphorylated M13mp18 replicative-form DNA (New England Biolabs), as described previously (12,32). After transformation of Escherichia coli XL1 by electroporation, recombinant M13 plaques were screened by hybridization with the HTLV-1 LTR-specific 32P-labeled oligonucleotide BIO5. Single-stranded templates were sequenced with the use of fluorescent dideoxynucleotides (Perkin-Elmer Applied Biosystems). The products were resolved on a 377A DNA sequencer (Perkin-Elmer Applied Biosystems) with 377A software (Perkin-Elmer Applied Biosystems). Sequence alignments were performed with Sequence Navigator Software (Perkin-Elmer Applied Biosystems).

Control PCR

To check the accuracy of the inverse PCR and the absence of PCR-associated recombination, we used as controls three cloned 3' HTLV-1 RU5 sequences flanked by their integration sites and harboring distinct mutations. Two hundred fifty copies of each of these three cloned sequences were mixed with 1 µg of uninfected DNA. Five hundred nanograms of the DNA mixture was amplified for 30 cycles with the use of 200 µM of the BIO6 and LTR1 primer pair under the same conditions as those used in the analysis of DNA samples from patients. Purified PCR products were cloned and sequenced as described below. Forty-one molecular clones were obtained, sequenced, and then analyzed by CLUSTAL alignment with Sequence Navigator Software.

PCR amplification of the{alpha}-enolase gene fragment was performed by classical PCR. Five hundred nanograms of DNA from patient P1 was amplified by Pfu DNA polymerase with the primer pair HA-ENO-S (5'-GGGGTTAAGGAAGAAAAGCA-3') at position 1038–1058 and HA-ENO-AS (5'- TTGGAACTGGAATTTCACACA-3') at position 1404–1383 (nucleotide coordinates are numbered according to the {alpha}-enolase GenBank reference: HSENOAL1) (27). Amplification and cycling were performed as described for inverse PCR. After the PCR products were cloned and sequenced, seven sequences were analyzed by CLUSTAL alignment with Sequence Navigator Software.

Quantification of HTLV-1 Proviral Load

The HTLV-1 proviral load was assessed by real-time quantitative PCR with the use of a dual-labeled fluorescent probe (ABI PRISM 7700 Sequence Detection System; Perkin-Elmer Applied Biosystems). Standard curves for the albumin and HTLV-1 tax genes were generated with the use of DNA extracted from HTLV-1-negative PBMCs for the former and an HTLV-1 plasmid for the latter. It was estimated that 10 ng of high-molecular-weight DNA (equivalent to roughly 1500 PBMCs) would contain 3000 copies (two copies per PBMC) of the albumin gene. The primer set for the HTLV-1 tax gene was PXF (5'-GAAACCGTCAAGCACAGCTT-3') positioned at 7163–7182 and PXR (5'-TCTCCAAACACGTAGACTGGGT-3') positioned at 7385–7364. The primer set for albumin was 5'-GCTGTCATCTCTTGTGGGCTGT-3' positioned at 16283–16304 and 5'-ACTCATGGGAGCTGCTGGTTC-3' positioned at 16442–16421 (nucleotide coordinates are numbered according to the albumin GenBank reference: HUMALBGC) (33). The TaqMan probe consisted of an oligonucleotide with a 5'-reporter dye and a 3'-quencher dye. The fluorescent reporter dye 6-carboxy-fluorescein was covalently linked to the 5' end of the oligonucleotide. The reporter was quenched by 6-carboxy-tetramethyl-rhodamine at the 3' end. The probe for the HTLV-1 tax gene was PXT (5'-TTCCCAGGGTTTGGACAGAGTCTTCT-3') positioned at 7331–7355; the probe for albumin was ALB (5'-CCTGTCATGCCCACACAAATCTCTCC-3') positioned at 16340–16366. TaqMan amplification was carried out in reaction volumes of 50 µL, with the use of the TaqMan PCR core Reagent Kit (Perkin-Elmer Applied Biosystems). Each reaction contained the following: 1x TaqMan buffer; 300 nmol/L of each primer; 200 nmol/L of each corresponding fluorescent probe; 3.5 mmol/L MgCl2; 200 µmol/L deoxyadenosine triphosphate, deoxycytidine triphosphate, and deoxyguanosine triphosphate; 400 µmol/L deoxyuridine triphosphate; 1.25 U of AmpliTaqTM Gold (Perkin-Elmer Applied Biosystems); and 0.5 U AmpEraseTM uracil N-glycosylase (Perkin-Elmer Applied Biosystems). Each sample was analyzed in triplicate with the use of 500 ng of DNA in each reaction. Thermal cycling was initiated with a 2-minute incubation at 50 °C, followed by a first denaturation step of 10 minutes at 95 °C and then by 45 cycles at 95 °C for 15 seconds and 58 °C for 1 minute (for tax) or 60 °C for 1 minute (for albumin).

Rex-Responsive Element and Two-Dimensional Structure Analysis

The secondary structure analysis of the Rex-responsive element (RXRE) variants was performed with ESSA-SAPSSARN software, as described previously (34,35). The probabilities of the pairing of each base with each other were computed with McCaskill's program (36).


    RESULTS
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 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Characterization of Samples Studied

DNA samples were obtained from six HTLV-1-infected individuals—two patients with TSP/HAM (P1 and P2), two asymptomatic carriers (P3 and P4), and two patients with ATLL (P5 and P6). The frequency of HTLV-1 DNA-positive PBMCs was estimated by real-time quantitative PCR to be 1% for the asymptomatic carriers; 3% and 2% for patients P1 and P2, respectively (TSP/HAM); and 41% and 62% for patients P5 and P6, respectively (ATLL). This distribution reflects the general finding of proviral load in the order ATLL > TSP/HAM > asymptomatic carriers. Fig. 1Go represents the pattern of HTLV-1-infected T-cell clones circulating in the peripheral blood of all six individuals as determined by inverse PCR. Because there is a stochastic element to inverse PCR amplification of low-frequency HTLV-1 integration sites (28), quadruplicate inverse PCR analysis was performed (4 x 0.5 µg, approximately 4 x 75 x 103 cell equivalents). A signal present in all four samples corresponded to a clonal frequency of one or more in 150 cells, while a single positive amplification would correspond to a frequency of one or fewer in 1500 (13,28).



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Fig. 1. Inverse polymerase chain reaction (PCR) analyses of 3' human T-cell leukemia virus type 1 (HTLV-1) integration sites. For each sample, 0.5 µg of digested and circularized DNA was submitted to quadruplicate PCR analyses. The molecular weight marker (M) was the DNA MWM X (i.e., molecular weight marker X) (Boehringer Mannheim Biochemicals, Indianapolis, IN). Patients are identified by their unique patient number. For each sample, the percentage of infected peripheral blood mononuclear cells (PBMCs), as estimated by quantitative PCR, is given in parentheses. No signal was detected after inverse PCR analysis of uninfected DNA (NC1 and NC2). As described previously, the abundance of detected clones, i.e., the number of infected cells that constitute these clones, is not proportionate to the intensity of the observed signal but is proportionate to the frequency of their detection after quadruplicate experiments (28). Signals identified by the numbered horizontal arrows correspond to the 29 sequences obtained after cloning and are detailed in Fig. 2Go. The sequencing allowed the determination of the length of these sequences (RU5 region and integration site). From the length of runoff products, we could infer the corresponding signals on the gels. A) Runoff analysis of inverse PCR products of samples from patients P5 and P6 with adult T-cell leukemia/lymphoma (ATLL). Patient P5 had an acute ATLL with 25 x 107 ATLL cells/L and a clear band after Southern blot analysis, meaning that the tumor cells were monoclonal with respect to the provirus integration of PBMC DNA. The pattern of clonally expanded T cells revealed one major clone (P5-1) with approximately 50 fewer abundant clones. Patient P6 had an acute subtype of ATLL with 9.5 x 105 ATLL cells/L. Five clones were found four times after runoff analysis. B) Runoff analysis of inverse PCR products of samples from patients P2 and P1 with tropical spastic paraparesis/HTLV-1-associated myelopathy. One clone from patient P2 (clone P2–4) was detected four times, while the remaining clones were detected at a lower frequency. C) Runoff analysis of inverse PCR products of samples from asymptomatic carriers P4 and P3. For both samples, all of the signals were detected less than four times and, therefore, corresponded to a clonal frequency of fewer than one in 150 infected PBMCs.

 
DNA from tumor cells in peripheral blood from patient P5 displayed a single dominant clone (clone P5–1) in a background of approximately 50 oligoclonally expanded, infected cells (Fig. 1Go, A). Five clones of a frequency of more than one in 150 infected PBMCs were detected in the DNA from circulating tumor cells derived from patient P6 (Fig. 1Go, A). Dilution of DNA from this sample was analyzed subsequently by inverse PCR (37), and only the signal at approximately 250 bp, which corresponds to clone P6-4, remained detectable at the concentrations tested (10 ng, not shown). As described previously, the abundance of detected clones, i.e., the number of infected cells that constitute these clones, is not proportionate to the intensity of the observed signal but is proportionate to the frequency of their detection after quadruplicate experiments (28). This finding indicated that a single clone harboring one integrated provirus dominated this sample. Therefore, both tumor PBMC samples from the two patients with acute ATLL displayed a malignant clone harboring a single integrated provirus in a background of polyclonally expanded, HTLV-1-positive CD4 T cells. The patterns of integration sites for the four remaining nonmalignant samples are given in Fig. 1Go, B and C. Both the proviral load and the number and the abundance of circulating clones were higher in the samples from the two patients with TSP/HAM than in the samples from the two asymptomatic carriers (Fig. 1Go), which is in agreement with the results of previous studies (11,13,14,37).

HTLV-1 Genetic Variability: Somatic Mutations Versus Reverse Transcription-Associated Errors in the Provirus

The primary aim of this study was to test the hypothesis that somatic mutations of the provirus could account for the HTLV-1 genetic variability. To examine in detail the possible HTLV-1 mutation process in vivo, we cloned inverse PCR products from the six patient samples without size selection. A total of 208 clones were sequenced, encompassing the 3' RU5 sequences of HTLV-1 (379 bp, approximately 80 kb of data) together with their flanking cellular sequences (mean, 119 bp; range, 7–362 bp; approximately 23 kb of data). The sequences could be arranged into 29 cellular clones based on cellular flanking sequences. The HTLV-1 regions of 29 cellular clones were aligned with respect to clone P1-1C, which was taken arbitrarily as the reference (Fig. 2Go).



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Fig. 2. Somatic mutation of the human T-cell leukemia virus type 1 (HTLV-1) 3' RU5 sequence in vivo. Overall, 29 distinct HTLV-1 3' integration sites were isolated; the corresponding flanking cellular sequence hexameric repeats, generated during integration, are given on the right. RU5 sequences were aligned according to patient P1 RU5 sequence on which the eight consensus mutations that differentiate all of the six patient RU5 consensus sequences are represented in boldface (top). Deletions are denoted by a dash. Coordinates of the sequence are those of the ATK-1 sequence (30). Those sequences harboring mutations in the flanking sequences are denoted by an asterisk. Patients are identified by their unique patient number (UPN). Each cluster of RU5 sequences sharing a common integration site (and, therefore, belonging to a unique clone of expanded T cells) is identified by its cellular clone number. Horizontal dark bar separates the clusters of sequences derived from each of the DNA samples from the six patients. Consensus RU5 sequences of patients are represented as gray lanes. For each cellular clone, the number of non-unique RU5 consensus sequences is indicated in parentheses. TSP = tropical spastic paraparesis; ATLL = adult T-cell leukemia/lymphoma.

 
Fig. 2Go represents the 208 RU5 sequences that were assorted into 29 distinct cellular clones. These clones were identified by the corresponding clusters of identical flanking sequences represented on Fig. 2Go by the hexameric repeats duplicated during integration. Not all RU5 sequences derived from the same cellular clone were identical—in fact, 18 (approximately 60%) of 29 clones were not homogeneous (Fig. 2Go). The number of variants per RU5 sequence ranged from zero to four. A total of 38 single-base substitutions and eight single-base deletions were scored. Some sequences harbored up to three to four mutations (i.e., clone P5-1 sequence C9, two transversions and two deletions; clone P6-4 sequence C90, three substitutions and one deletion; and clone P2-4 sequence C565, one transition and two transversions). Such clones clearly put these sequences well beyond PCR error artifacts. The PCR error was estimated for this region to be fewer than one per 18.5 kb sequenced (see the "Materials and Methods" section). Since recombination can occur during PCR (38), it was possible that some mutations could be ascribed as being derived somatically when, in fact, they could have occurred from recombination. Accordingly, 250 copies each of DNA from three molecular clones of different provirus : cell junctions (Fig. 2Go, P1-1 C109, P1-4 C253, and P1-3 C344) were mixed in equimolar ratio and amplified (see the "Materials and Methods" section). Small sequence differences in the RU5 region allowed the three proviruses to be distinguished. PCR products were cloned, and 41 molecular clones were sequenced. No recombinants were observed, indicating that the recombination frequency was 2.4% or less.

Fig. 3Go summarizes the different steps at which a mutation can occur during the synthesis of the HTLV-1 provirus. Accordingly, the distribution of the 46 RU5 mutations indicates that they did not correspond to minus-strand, synthesis-associated reverse transcription errors, which are expected to result in a homogeneous population of RU5 sequences (Fig. 3Go, A). Similarly, the presence of at least one patient consensus sequence within each of the 29 clones demonstrated that none of the 46 RU5 substitutions corresponded to a plus-strand, synthesis-associated mutation corrected before the newly infected host cell divided. These substitutions, all of which were harbored by only a subset of sequences within clones, might have theoretically resulted from somatic mutations or from reverse transcription-associated substitutions during a single cycle of the plus-strand synthesis of the provirus, in the absence of DNA mismatch repair before the first division of the newly infected CD4+ T cell (Fig. 3Go). However, two aspects of our results rule out the second possibility. First, the actual RU5 mutation frequency is incompatible with such reverse transcription-associated errors. Indeed, if the mutations shown in Fig. 2Go corresponded to plus-strand, synthesis-associated errors, they all would necessarily be the result of a single cycle of reverse transcription. Accordingly, the average RU5 mutation frequency was an approximately 4.2 x 10-3 substitution per replication cycle per base (46 distinct substitutions in 29 RU5 sequences of 379 bp), which is about 600 times higher than that for the HTLV-1 reverse transcription (39) and 100 times higher than that for the human immunodeficiency virus reverse transcription (40). Second, some RU5 sequences harbored two to four substitutions, a mutation frequency not observed to date for a single step of reverse transcription-mediated DNA elongation. Therefore, the majority of substitutions appear to result from somatic mutations during cellular replication and not from plus-strand, synthesis-associated reverse transcription errors or from PCR artifacts or recombinations. Fig. 2Go shows that, for about 60% of the HTLV-1-positive clones, 8%–80% of the infected cells harbored a somatically mutated HTLV-1 provirus. The 46 somatic mutations are detailed in Table 1Go. As can be seen, the ratio of the transition to the transversion was one, while substitutions from C and G were more frequent (which is in keeping with the high GC content of the RU5 region [approximately 60%]).



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Fig. 3. Human T-cell leukemia virus type 1 (HTLV-1) 3' RU5 sequence mutations at different stages of HTLV-1 replication. PBS = primer binding site; PPT = polypurine tract; – = minus strand; + = plus strand. Different steps in HTLV-1 RU5 sequence replication that may affect the retroviral mutation rates are illustrated. RNA is represented as a thin line, whereas DNA is represented as a thick line. The first six horizontal lines represent the synthesis of the provirus. Line 7 represents the integrated provirus flanked with its two integration sites represented as black boxes. NlaIII restriction sites are represented on both the provirus and the 3' cellular flanking sequence. Lines 8 and 9 represent the first two mitoses of the infected cells that harbor the integrated provirus. For each cell, the RU5 sequence and the 3' flanking sequence encompassed by the 2 NlaIII restriction sites (i.e., the sequences obtained after inverse polymerase chain reaction [PCR]) are represented. Line 10 represents the sequences obtained after inverse PCR, cloning, and sequencing. A) Reverse transcription-associated substitution during minus-strand synthesis. Open circle represents a mutation that has occurred during the synthesis of the 5' RU5 of the minus strand. As shown in lines 1–6, this substitution appears to be harbored by both strands of the two long terminal repeats of the integrated provirus. Accordingly, all of the infected cells from the corresponding clone (identified by their common integration site) harbor a provirus with the same substitution (see lines 8 and 9). As a consequence, all of the sequences from this clone obtained after inverse PCR harbor the same mutation at the same position. B) Reverse transcription-associated substitution during plus-strand 3' RU5 synthesis, in the absence of DNA mismatch repair before the first division of the newly infected CD4+ T cell. Such substitution is harbored by only one strand of the integrated provirus (line 7). Accordingly, after the first mitosis and in the absence of DNA repair (such repair being expected to result in the absence of mutation or in a pattern identical to that shown in A), such substitution results in two populations of infected cells: One population harbors the RU5 substitution, and the other harbors the native RU5 sequence. As a consequence, two populations of sequences are obtained after inverse PCR (line 10). C) Somatic mutation of the 3' RU5 sequence in the absence of reverse transcription-associated substitution. In this case, no substitution occurs during the provirus synthesis. After integration, a 3' RU5 substitution that has occurred during the second mitosis is represented on line 9. As for the plus-strand substitution, such somatic mutation results in a double population of RU5 sequences (line 10). The means by which it was possible to distinguish somatic mutations from plus-strand-associated substitutions are explained in the text.

 

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Table 1. Analyses of somatic mutations in RU5 region of human T-cell leukemia/lymphoma virus type 1 provirus and flanking cellular sequences*
 
Somatic Mutation of Cellular Sequences

Somatic mutations were also found in the flanking sequences of two clones (Fig. 4Go). For the first example, P5-1 from an ATLL patient (Fig. 1Go), the cellular DNA sequence did not correspond to anything in the current databases (sequence was analyzed as described by comparison to the nonredundant human sequence database, the human complementary DNA [dbEST] database, and the MONTH [June 2000] database by use of BLASTN with Search Launcher, FASTA, and Repeat Masker) (32). Three sequences encoded four single-base substitutions with respect to the most abundant sequence, which was assumed not to be somatically mutated (Fig. 4Go, A). For clone P1-7, derived from a TSP/HAM sample, the proviral flanking sequences showed virtual identity with the promoter region of the human {alpha}- (or non-neuronal) enolase gene (27). Proviral integration occurred 146 bp 5' to the TATA box and, as a result, uncoupling the TATA box from a number of transcriptional motifs, such as the CCAAT box, AP1, and PEA2 sites. As Fig. 4Go, B, shows, all five sequences harbored a T->C transition at position 1217 with respect to the published sequence, while one carried two additional purine–purine transitions. The T1217C transition could represent a single nucleotide polymorphism in the patient's P1 {alpha}-enolase gene or a somatic mutation acquired during clonal expansion.



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Fig. 4. Effects of human T-cell leukemia virus type 1 (HTLV-1) infection on the genomic sequence of infected cells belonging to two circulating clones. HTLV-1 3' flanking sequences are aligned relative to the first base of the hexameric repeat sequences flanking the provirus (NlaIII site underlined). The number of sequences is given in parentheses. Distinct sequences are identified by different number of asterisk marks associated with the number (n) of clones. A) Three of the 14 flanking sequences that derived from the malignant adult T-cell leukemia/lymphoma clone P5-1 harbored four substitutions. B) Clone P1-7 harbored a provirus that disrupted the first exon of the {alpha}-enolase gene for non-neuronal enolase (HSENOAL) (27) at position 1034. In addition, sequencing five HSENOAL sequences allowed the detection of three distinct substitutions. The T->C transition at position 1217 was present in all five sequences, while sequence P1-7* harbored two additional substitutions, indicating that these mutations were accumulated in a stepwise manner. Numbering is that of HSENOAL entry.

 
To distinguish between the two, we amplified DNA from whole PBMC DNA by using oligonucleotides encompassing position 1217. To avoid the amplification of residual P1-7 sequences, the 5' oligonucleotide was complementary to a region of the {alpha}-enolase gene 5' to the integration site. All seven recombinants sequenced were identical and had T at position 1217. Therefore, the T1217C substitution was restricted to clone P1-7, indicating that it represented a somatic mutation. Accordingly, sequence P1-7* (Fig. 4Go, B), with a further two substitutions at positions 1160 and 1365, was somatically derived from P1-7. PBMC DNA from this patient was amplified on two occasions, 4 and 6 years later. The clone with the T1217C substitution was identified on both occasions (not shown), indicating that the somatically mutated clone was indeed persisting over time.

A total of seven substitutions (Table 1Go) were found in approximately 24 kb of cellular DNA, representing a frequency of 2.8 x 10-4/bp sequenced. This frequency is remarkably similar to that for the HTLV-1 RU5 region (5.8 x 10-4/bp sequenced [Table 1Go]), particularly in light of the very different base composition of the two regions (HTLV-1, 40% AT [i.e., adenosine and thymidine]; cellular, 59% AT). These values are mutation frequencies, not mutation rates, because it is not possible to estimate the average number of mitoses in any lineage. If the same mutation frequency applied across the whole genome, one would predict approximately 1.7 x 106 mutations per diploid cell (approximately 2.8 x 10-4 x 6 x 109), or approximately one mutation per 3.5 kb (a remarkable mutation load).

Distribution of Somatic Mutations Within RU5

The mutations appear to be generally distributed randomly across the HTLV-1 RU5 region, with no evidence of hot spots (Fig. 5Go). About 70% of all CpG dinucleotides in the genome of vertebrates are methylated (41), and the HTLV-1 provirus has been found to be heavily methylated in cell lines (42). Five of 14 G->A and C->T transitions (36%, positions 8708, 8781, 8797, 8813, and 8842) occurred within CpG dinucleotides, suggesting that they might have arisen as a result of deamination of 5mCpG.



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Fig. 5. Random distribution of the somatic mutations along the human T-cell leukemia virus type 1 (HTLV-1) 3' long terminal repeat RU5 sequence. RXRE = Rex-responsive element; ATLL = adult T-cell leukemia/lymphoma; TSP/HAM = tropical spastic paraparesis/HTLV-1-associated myelopathy.

 
RXRE is a cis-acting RNA element required for Rex function and maps to the U3R region of the LTR (4347). HTLV-1 Rex protein acts at the post-transcriptional level to induce the appearance of unspliced and singly spliced viral messenger RNA in the cytoplasm (48). The action of Rex requires both the overall secondary structure intrinsic to the RXRE and the specific sequences from one small region of this large structure (44,49,50), which forms a protein-binding site for Rex. Mutational analyses of RXRE have demonstrated that Rex binding in vitro is related to the function of Rex in vivo (45,50).

Of the 46 mutations, 22 mapped to 21 sites in the RXRE. Some mapped to the Rex-binding site or else disrupted RNA secondary structures (Fig. 6Go), which was evidenced by de novo calculations using the variant sequences (not shown). Because RXRE plays a crucial role in the expression of HTLV-1, it is likely that a number of these variants will have impaired expression of viral proteins.



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Fig. 6. Somatic mutations within the Rex-responsive element (RXRE) secondary structure. The RXRE corresponds to the consensus sequence of patient P1. *Position 1 corresponds to position 8615 of the ATK sequence (30). The three Rex-binding motifs are enclosed by rounded-triangular outlines at the right side (4347). Fig. represents the 22 substitutions together with their corresponding sequence number as represented in Fig. 2Go. del = deletion.

 
Somatic Mutation Frequency and Number of Infected Cells Within Clones

It has been clearly established that genomic instability is characteristic of some cancers (51). Accordingly, tumor clones P5-1 and P6-4 from the two ATLL patients harbored a high mutation frequency (Fig. 2Go). However, Fig. 2Go shows that somatic mutations were not restricted to ATLL samples. If somatic mutation were associated with DNA replication, then its extent should be generally related to the number of rounds of mitosis, which may be reflected in the clonal frequency in vivo. It is possible to get an approximate estimate of clonal frequencies from quadruplicate inverse PCR (compare Fig. 1Go) (14,28). Fig. 7Go, A, shows that the average number of mutations per kilobase in 57 RU5 sequences belonging to six HTLV-1 clones with a detection frequency of one or more per 150 PBMCs was more than twice that of the remaining 151 sequences derived from 23 clones with a detection frequency of fewer than one per 150 PBMCs. This difference was statistically significant (two-sided P = .037; Student's t test for unpaired samples). Indeed, since the frequency of abundant, circulating clones is in the order ATLL > TSP/HAM > asymptomatic carriers, the number of acquired somatic mutations was higher in HTLV-1-associated disease than in the virus carriers. As Fig. 7Go, B, shows, PBMC DNA from ATLL patients, TSP/HAM patients, and asymptomatic carriers harbored a mutation frequency of 0.71, 0.52, and 0.22 substitution per kilobase of sequence (RU5 sequences plus integration sites), respectively. The difference between asymptomatic carriers and ATLL patients was statistically significant (two-sided P = .048; Student's t test for independent samples). However, the somatic mutation frequency of the 45 sequences derived from the six nonmalignant ATLL clones was identical to that of TSP/HAM sequences: 0.5 substitution per kilobase of sequence (RU5 sequences plus integration sites).



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Fig. 7. Increased somatic mutations in RU5 and flanking sequences associated with disease. A) Frequency of somatic mutations in RU5 sequences according to the abundance of circulating clones. The detection frequency of circulating human T-cell leukemia virus type 1 (HTLV-1)-positive clones was estimated by quadruplicate inverse polymerase chain reaction analysis (Fig. 1Go). B) Distribution of the frequency of somatic mutations along both the RU5 and flanking sequences according to the clinical status. kb = kilobase; PBMCs = peripheral blood mononuclear cells; ATLL = adult T-cell leukemia/lymphoma; TSP/HAM = tropical spastic paraparesis/HTLV-1-associated myelopathy.

 

    DISCUSSION
 Top
 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Hitherto, the generator of retroviral mutations has always been considered to be the reverse transcriptase. From the time that clonal expansion of cells bearing retroviral proviruses was identified (12) in the context of Tax-associated DNA repair impairment (2022,24,25), mutation accrued during mitosis could be considered as another possible source. The data given above indicate that somatic mutation of HTLV-1 and of cellular sequences occurs, even within clones derived from asymptomatic carriers. That the frequency of somatic mutation increases with the HTLV-1 clonal frequency strongly suggests that there is a progressive accumulation of mutations as opposed to a single hypermutational event (Figs. 4 and 6GoGo). Obviously, mutations in p53, p16 deletions, and chromosomal rearrangements associated with ATLL might have undergone the same process as that described in our study for both the provirus and its flanking sequences.

The average mutation frequency of HTLV-1 flanking sequences is remarkably high, approximately 1.7 x 106 mutations per diploid cell (approximately 2.8 x 10-4 x 6 x 109), or approximately one mutation per 3.5 kb. For some clones, the mutation frequency is even higher. Since mutations are accumulated in a stepwise manner (Fig. 4Go, B), it is not possible to estimate the rate of somatic mutation per mitosis. However, a few approximate calculations may be made for that number. Since the host genome mutation rate is something of the order of 10-10 to 10-9 per base per mitosis, attaining a mutation load of several million substitutions per genome would require an unrealistic number of rounds of replication for any one clone—approaching 106. Note that this calculation is based on mutation rates. Because a fraction of the mutations are probably deleterious, the number of rounds of replication is underestimated. However, if some arms of mismatch repair were inactivated, then this number could be reduced considerably. Given the data showing that the HTLV-1 Tax protein interferes with the DNA mismatch repair systems in vitro (21,22), it may be assumed that the Tax protein may do the same thing in vivo.

Given a novel source of mutation, to what extent is this exploited, if at all, by the infected cell? It is probably safe to assume that, with such a mutation pressure, there is considerable negative selection, although the associated cell loss might well be masked by clonal expansion. Expression of HTLV-1 proteins marks out the cell as non-self and a target for cell-mediated immunity, which is particularly intense (52). Although deletions of only single nucleotides were noted, if they occurred in the viral open reading frames, they could considerably limit protein expression. Negative selection will ensure maintenance of the Tax+ phenotype. Because Tax is a target for cytotoxic T lymphocytes (CTLs), it is possible that some mutations in human leukocyte antigen (HLA) class I-restricted CTL epitopes are compatible with Tax function, which, therefore, allows the cellular clone to escape cell-mediated immunity (53). This mutation in HLA class I-restricted epitope that is compatible with Tax function may occur as long as most of the other HTLV-1 proteins were inactivated first by somatic mutations. This speculation follows from the observation that cases of CTL escape seem to occur only when the cellular immune responses are highly focused on a single target (54).

It has been demonstrated that Tax expression is observed frequently in CD4+-infected cells in vivo (55). By contrast, HTLV-1 infection in vivo is characterized by a very low viremia, which is usually undetectable. One can explain this paradoxical association of Tax expression with the absence of viremia by the fact that the genetic organization of HTLV- 1 (especially the robust inactivation of Tax expression by Rex) precludes an intense expression of Tax that is needed for viral production. By contrast, the level of Tax expressed in the cell appears to be sufficient to interfere with the numerous cellular pathways leading to clonal expansion. Is it possible that some of the mutations are incorporated into packaged virion RNA? Since RNA viruses manifest mutation rates of one to two per genome per cycle (56), which can only be slightly increased by chemical mutagenesis (57), it seems, by extrapolation, that an HTLV-1 provirus could absorb between one and two substitutions without suffering irreversible loss. However, the low fixation rate of amino acid substitutions—of the order of 0.1% per century (58)—would suggest that the majority of somatically mutated proviruses are without virus progeny.

In view of the similarities among the HTLV/bovine leukemia virus (BLV) group of retroviruses, somatic mutations are expected to be identified for simian T-cell leukemia virus, HTLV-2, and BLV. It is of note that both a replication via mitosis and a defective DNA repair in infected cells have been evidenced for HTLV-2 and BLV (22). It will be interesting to see if high-frequency somatic mutations pertain to other viruses having a replication cycle that can include integration and host cell proliferation. In this context, it may be noted that the hepatitis B virus X protein has been found to disrupt cellular DNA repair (59).

In conclusion, the in vivo HTLV-1 genetic variability results predominantly from somatic mutations of the proviral sequence rather than from reverse transcription-associated substitutions. Furthermore, it appears that somatic mutation accompanies clonal expansion. The degree of somatic mutation is so great that it is consistent with the in vitro findings of a Tax-associated mutator phenotype. Not all somatic mutations were deleterious to the HTLV-1-bearing cellular clone, for there was evidence of sequential mutations. These findings suggest that somatic mutations after integration, presumably coupled with selection, help move the cellular clones toward a transformed phenotype, of which ATLL is the end point. A conundrum would appear to remain: Given the mutation pressure due to the high level of somatic mutations, why is the lifetime risk of ATLL as low as 3%–5%? The expression of HTLV-1 proteins would mark the infected cell as non-self. Furthermore, the frequency of neoantigen formation could be higher than that typically found in other malignancies. Together, these features may allow robust control by host cell-mediated immunity. In support of this hypothesis, clinical observations and experimental investigations have shown that suppression of HTLV-1-specific cellular immune response led to the development of ATLL (6064).


    NOTES
 
I. Leclercq and A.-S. Gabet contributed equally to this work.

Present address: I. Leclercq, Université Mons-Hainaut, Service de Biologie Moléculaire-Pentagone aile A 3ème étage, Mons, Belgium.

Supported by grants from the Association pour la Recherche sur le Cancer, from the Fondation Contre la Leucémie, and from the Comité Départemental du Rhône de la Ligue Nationale Contre le Cancer. I. Leclercq and F. Mortreux were supported by bursaries from the Ministère de l'Enseignement Supérieur et de la Recherche. A.-S. Gabet was supported by a grant from the Centre Léon Bérard.

We thank Claudine Pique, Ali Saib, and Renaud Mahieux for their critical review of this article. We also thank Pierre Wattre and collaborators in whose laboratories we conducted the DNA extraction, digestion, ligation, and polymerase chain reaction analysis. We are grateful to Marie-Dominique Reynaud for assistance with the preparation of this manuscript and to Dr. Michel Crépin (from the plateau technique de séquençage du Centre Hospitalier Régional Universitaire de Lille) for technical assistance.


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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Manuscript received July 26, 2000; revised December 29, 2000; accepted January 10, 2001.


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