Introduction

The fact that the DNA of cancer cells is different from the DNA of normal cells indicates that carcinogenesis involves substantial errors in DNA replication, deficits in DNA repair, and alterations in chromosomal segregation.¹ Given the rarity of mutations in normal cells and the large numbers of mutations observed in human cancers, it has been proposed that the spontaneous mutation rate in normal cells is not sufficient to account for the number of mutations found in human tumors. In other words, cancer is assumed to result from an increased somatic mutations frequency, ie from a mutator phenotype. DNA damage, combined with loss of DNA repair and clonal selection, is thought to help move the normal cell towards such a phenotype.

The main human tumor viruses include human T cell leukemia viruses, Epstein–Barr virus, human papilloma virus, Kaposi sarcoma-associated herpes virus and human hepatitis B and C viruses. These virus provide exceptional systems to study the mechanisms underlying human cancers. Among these, human T cell leukemia viruses type 1 (HTLV-1), the first human pathogenic retrovirus isolated, is the etiologic agent of the malignant CD4 T lymphoproliferation adult T cell leukemia/lymphoma (ATLL).^2,3 Furthermore, this virus has been associated with the development of a chronic progressive neuromyelopathy (tropical spastic paraparesis (TSP)/HTLV-1-associated myelopathy (HAM)),⁴ and, to a lesser extent, to a variety of inflammatory diseases.^5,6,7,8,9

Among the 15–25 million individuals infected worldwide, approximately 3% to 5% will develop ATLL, depending on as yet unknown cofactors. ATLL harbors different clinical features resulting in a division of the spectrum of the disease into four clinical subtypes^10,11 referred to as acute, lymphoma, chronic and smoldering subtypes. Infection early in life is crucial in the development of ATLL.¹² The prolonged premalignant asymptomatic phase that precedes leukemia is characterized by the persistent oligoclonal expansion of HTLV-1 infected cells.^{13,14,15,16,17,18,19,20,21} This results from the action of the HTLV-1 Tax protein that triggers T cell proliferation^22,23 and negatively interferes with the main cellular pathways involved in the maintenance of genomic integrity.^{22,23,24,25,26,27,28,29,30} The mutagenic effect of Tax generates a hitherto undescribed mechanism of retrovirus genetic variability that results from somatic mutations of the proviral sequence rather than from reverse transcription.¹⁸ In fact, HTLV-1 genetic variability and infected cell genetic instability go hand in hand, thereby establishing a pro-cancer genotype.

Reverse transcription is an error-prone process

RNA viruses are unique in that their polymerases, although clearly marked by the same basic quaternary structure of the polymerization domain common to all nucleic acid polymerases, lack a particular domain encoding a 3' exonuclease.^31,32 Furthermore, host exonucleases are not incorporated into specific virions. Consequently, any polymerization error becomes immortalized. Not surprisingly, the highest mutation rates observable for microbes are to be found in RNA viruses.³³ Higher mutation rates – referred to hypermutations – can theoretically and practically be encountered. However, there comes a threshold beyond which the mutation rate is deleterious to the survival of the virus.³³ Retroviruses are somewhat schizophrenic: on the one hand they undertake DNA polymerization, while on the other hand their reverse transcriptase is devoid of any 3' exonucleolytic activity. Thus the genetic variability typical of RNA viruses becomes part of their way of life. For some reason that is not yet evident, the average mutation rate of retroviruses is approximately 20-fold lower than that of an ensemble of familiar RNA viruses.³⁴

HTLV-1 is genetically stable

The first complete HTLV-1 sequence emerged in 1983.³⁵ Since then, a forcible accumulation of sequence data has shown that HTLV-1 is extraordinarily stable genetically. While some segments are slightly more variable than others, geographically widely dispersed viruses are <10% divergent at the nucleic acid level. This includes isolates from tribes and aboriginal populations isolated from the mainstream of human movement for thousands of years.³⁶ Intrapatient variability is usually <0.5%, a value significantly lower than that observed with lentiviruses, such as HIV-1. The intrapatient variation of the HIV-1 envelope protein 5 years post infection is greater than that of all HTLV-1 envelope proteins to date.³⁷ Even antigenically stable RNA viruses, such as measles and hepatitis A virus, have, in this PCR era, shown more variability than expected. Thus HTLV-1, and its close or distant cousins, HTLV-2 and bovine leukemia virus (BLV) respectively, constitute remarkable exceptions, unique among the genetically promiscuous RNA viruses.

High circulating proviral loads characterize HTLV-1 infection

The non-malignant neurological disorder TSP/HAM is accompanied by very high proviral loads, sometimes approaching 1/5 peripheral blood mononuclear cells.^{38,39,40,41,42,43} Consequently, substantial proportions of the CD4 T cells in the periphery may be infected. Proviral loads during the long asymptomatic phase of infection are lower than in TSP/HAM, although they may be as high as 1/25 PBMCs.^{38,39,40,41,42,43} Here, there is evidence of an extensive replication, yet HTLV-1 is genetically more stable than any other RNA virus. HIV proviral loads in late-stage disease, AIDS, may reach up to 1/100 to 1/30 in the periphery. In other words HIV-1 also shows signs of extensive viral replication, although the maximum HIV-1 proviral load is lower than that observed for HTLV-1. Yet HIV-1 varies considerably, as one would expect.³⁷ By contrast, HTLV-1 remains stable while displaying very high proviral loads. How is this possible? The misincorporation rate of HTLV-1 reverse transcriptase turns out to be on the lower side (7 10^-6/base/cycle),^44,45 while that of HIV-1 is on the upper side (3.5 10^-5/base/cycle).⁴⁶ Only one log discriminates between them. Of all the components underlying viral variation, the mutation rate is among the least important while the number of successive cycles of replication is probably the most important.⁴⁷ How can the virus remain stable while replicating intensely?

Persistent clonal expansion of infected cells, a way of life for HTLV-1

The paradoxical combination of an elevated proviral load with a remarkable genetic stability suggested that the virus replicated in concert with cell mitosis (Figure 1).¹⁹ This was confirmed for all cases of HTLV-1 infection, notably asymptomatic carriers, TSP/HAM and ATLL.¹³ In ATLL, tumor cell clones were identified against a general background of oligo- or polyclonal expansion of infected, but non-transformed cells.¹⁴ The number of clones detected varied considerably but remained between five and 60 clones per sample (Figure 2). The fact that there may be hundreds or more clones within an individual clearly indicates that HTLV-1 does replicate via reverse transcription in vivo. However the bulk of the proviral load is made up of the most abundant clones, which are relatively few (<10–15) in each sample. Consequently we conclude that HTLV-1 in vivo replication predominantly occurs via mitosis rather than via reverse transcription. Analyzing the pattern of HTLV-1 replication over time revealed that the Tax-driven expansion of T cells may persist for considerable periods of time, despite strong cellular immunity. Hence, most circulating HTLV-1-positive clones persist over time. However, the abundance of these persistent clones, ie the number of cells within clones, is shown to fluctuate over time in some infected individuals, suggesting that the degree of infected T cell proliferation (or elimination) varies over time. Interestingly, in asymptomatic carriers, infected cells accumulate within circulating clones, since detected clones are more abundant in persons with longstanding infection.¹³

Figure 1.

Schematic representation of HTLV-1 replication in vivo.

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Figure 2.

Distribution of HTLV-1 infected clones in two patients with TSP/HAM (patients P1 and P2). DNA from PBMCs was analyzed by IPCR. Each signal on the gel corresponds to a cluster of >100 proviruses with the same flanking sequence, ie belonging to the same cellular clone. Quadruplicate experiments permitted estimation of the abundance of cells within circulating clones, most abundant cells being the most frequently detected after four experiments. M, molecular weight marker; NC, negative control (adapted from Ref. 18).

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The HTLV-1 proviral load of TSP/HAM patients may draw near 10% of PBMCs, or approximately 3 10⁹ copies (the proviral burden in secondary lymphoid organs is unknown). Assuming a minimum of 10 HTLV-1-driven proliferating clones and that proliferation is not unduly abnormal, ie the clones divide approximately every 24 h, it would be possible to attain such a proviral load after approximately 28 divisions. Even if the proviral load was 10-fold greater, only 31 divisions would be needed. Clearly, as only few HTLV-1 carriers ultimately develop non-malignant disease after 20–40 years of infection, clonal expansion must be actively restrained, otherwise CD4 lymphocytosis would be obvious within weeks post infection.

As clonal expansion is held in check, the question arises of how. Anti-HTLV-1 cellular immunity is the obvious explanation. It is present in asymptomatic carriers⁴⁸ and is particularly intense among TSP/HAM patients⁴⁹ in whom the proviral load is much greater than in the former group. Indeed, anti-tax activity is especially intense in TSP/HAM patients, easily detectable in direct CTL assays. Although tax expression directly induces the cell to proliferate and up-regulates a number of cytokines and transcription factors, it is antigenically non-self, which is detrimental to clonal survival.

Two-step nature of HTLV-1 primary infection in vivo

As there is no evidence for cell-free virus in vivo, HTLV-1 transmission must involve infected allogeneic cells. Reverse transcription must follow soon, as allogeneic cells are ephemeral. However, once a few rounds of reverse transcription are over, the clonal expansion of infected T cells predominates. The squirrel monkey (Saimiri sciureus) constitutes an interesting model for investigating the tempo of HTLV-1 replication during primary infection. After experimental infection of squirrel monkeys (Saimiri sciureus) with HTLV-1 infected cells, the virus appeared to be transcribed only transiently into the animal circulating blood, spleen and lymph nodes.⁵⁰ A stable disappearance of the viral expression occured at 2–3 weeks from inoculation, coinciding with the development of the anti-HTLV-1 immune response and the persistent detection of the provirus in PBMCs. Analyzing the HTLV-1 replication pattern over time in PBMCs and various organs from HTLV-1 experimentally infected monkeys revealed that PBMCs and lymphoid organs constitute the major reservoirs for HTLV-1.^50,51,52,53 A pattern of persistent clonal expansion of infected cells, identical to that observed in humans, was evidenced in the PBMCs of experimentally infected animals.⁵¹ The dissemination of the virus throughout the different body compartments appeared to result from the cellular transport of the integrated provirus. The circulating proviral burden increased as a function of time in one animal studied over a period of 4 years. The high proviral loads observed in the last samples resulted from the accumulation of infected cells via the extensive proliferation of a restricted number of persistent clones on a background of polyclonally expanded HTLV-1-positive cells. Therefore, the animal model revealed that HTLV-1 primary infection would be a two-step process that includes a transient phase of reverse transcription followed by the persistent multiplication of infected cells. As represented in Figure 3, the degree of T cell proliferation may vary between animals. These results suggest that when attempting to block HTLV-1 replication the choice of the target might depend on the stage of infection.

Figure 3.

Schematic representation of HTLV-1 replication over time. The gray curve represents the early and transient step of vertical replication through reverse transcription and integration that allows allogeneic cells to infect the new organism. Black curves represent the persistent clonal expansion of newly infected cells. The degree of expansion varies between clones and/or between infected organisms.

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Pathogenic and clinical implications of HTLV-1-infected T cell proliferation

Although flower cells, typical of ATLL, could be detected on blood smears, hyperlymphocytosis is a very rare event in HTLV-1-infected individuals with no malignancy. Indeed, the number of infected cells within circulating clones ranges from 1/3000 to 1/150 PBMCs. As proviral loads are higher in TSP/HAM than in asymptomatic carriers, both the mean number of clones and the abundance of cells within theses clones have been found to be significantly higher in TSP/HAM.

In the spinal cord of patients with HAM/TSP, infiltrating CD4-positive lymphocytes seem to be the major reservoir for the virus.⁵⁴ This is thought to play a central role in the pathogenesis of the disease. An oligoclonal proliferation of these HTLV-1-infected CD4 lymphoid T cells is present in the cerebrospinal fluid of the patients.⁵⁵ Furthermore, clonal populations of HTLV-1-infected lymphocytes sharing identical HTLV-1 proviral flanking sequences (ie integration sites in the cellular DNA) and thus derived from a single HTLV-1-infected progenitor, are detected, for a given patient, in both the cerebrospinal fluid and the peripheral blood. This indicates that HTLV-1 crosses the blood–brain barrier by way of the migration of HTLV-1-infected lymphocytes in vivo. Similarly, the dissemination of the virus throughout the different body compartments has been evidenced in humans as in animal models (Figure 4).

Figure 4.

Quadruplicate linker-mediated PCR analysis of HTLV-1 integration sites in DNA samples derived from a cell line and from various organs of an experimentally infected squirrel monkey. DNA was analyzed by quadruplicate LMPCR. Amplified products were submitted to run-off analysis with an HTLV-1 3' LTR specific oligonucleotide. Run-off products were resolved on a sequencing gel. M, molecular weight marker. E1540 cell line was used for experimental infection. Note that common clones are present in different body compartments (adapted from Ref. 51).

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Monitoring the clonality of malignant cells over time in patients treated for ATLL has been found to be a useful tool to assess treatment efficacy.^14,56 Figure 5 represents the clonality pattern of HTLV-1-infected cells over time in a 60-year-old woman treated for smoldering ATLL with skin infiltration. Southern blotting showed that this infiltration corresponded to tumor cells bearing a single HTLV provirus. Local chemotherapy induced a near complete remission of skin lesions. A second skin biopsy of a remaining lesion was performed. Figure 5 shows the HTLV-1 integration patterns of both samples. A single, intense band at 365 bp dominated the tumor samples. Nonetheless, there were 38 additional bands of lower frequency, some of which only came up once. The number of detectable bands in the post-therapy sample was much lower. Only nine were detected, although the 365 bp band, presumably corresponding to the tumor cells, was found in 3/4 samples, suggesting a frequency of 1/300 of all cells recovered. Six courses of systemic chemotherapy resulted in complete clinical remission. A PBMC sample taken at remission was analyzed, revealing clonally expanding HTLV-1 T cells. The 365 bp band was also present at a frequency of 4/4 samples or 1/150 PBMCs.

Figure 5.

Run-off analysis of IPCR products of tumor cell DNA from skin biopsies of a patient with ATLL (A) and after local chemotherapy (B). Lane C: analysis of IPCR products derived from the PBMC sample taken 6 months after the second skin biopsy. After local chemotherapy, the number of clones detected decreased from 39 to nine. The major integration site, identified by a horizontal arrow, was detected 4/4 times in the first sample; 2/4 replicas were still detected in the second biopsy and third sample, respectively (adapted from Ref. 14).

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Analyzing additional patients under treatment revealed that the persistence of tumor clones at high frequencies (>1/300 PBMCs) was commonly observed, even in complete responders. This persistence was invariably correlated with relapse and/or poor outcome.⁵⁶ A fluctuating frequency of some tumor clones was observed, with evidence for clonal change under treatment in one patient, indicating that treatment of ATLL can result in the selection of resistant clones. Finally, allogeneic bone marrow transplantation (BMT) using an HTLV-1-infected sibling as donor was found to be associated with the long-lasting disappearance of tumor clones and a possible cure of the disease. A long-term persistent clonal expansion of circulating HTLV-1 bearing T cells, deriving from the donor bone marrow, was evidenced in this patient. Data indicate that the variable success of ATLL treatments is probably due to the clonal heterogeneity of malignant cells that results in the selection of resistant clones. Semi-quantitative assessment of infected T cell clonality should be able to predict treatment failure. Accordingly, additional therapy may be tailored to the clonality pattern observed after first-line therapy.

Tax triggers T cell proliferation and genetic instability

There is ample literature documenting the influence of the HTLV-1 Tax protein on the up-regulation of transcription factors and lymphokines and their receptors (for recent review, see Ref. 57). In a recent work, Hanon et al⁵⁸ demonstrated that Tax expression was the norm for infected lymphocytes that derived from infected individuals without malignancy. Therefore, the pleiotropic function of Tax is thought to cooperate in promoting the proliferation of infected T cell. Indeed, Tax protein induces the expression of numerous genes involved in the differentiation and the proliferation of T cells.^59,60 Through the functional inactivation of P16^INKA4,⁶¹ and the activation of cyclin D2⁶² and cyclin D3,⁶³ Tax intervenes directly in the pathway controlling T cell proliferation. Furthermore, the N terminus of Tax has recently been found to directly and specifically interact with CDK4, resulting in a Tax/CDK holoenzyme complex that capably phosphorylates the Rb protein.⁶⁴

In addition to its positive effect on cell cycling, Tax protein negatively interferes with some DNA repair functions of the host cells. Indeed, Tax inactivates in trans the promoter of the human polymerase²² and disrupts other prominent cellular DNA repair pathways.^24,26 By competing with p53 for p300/CBP binding, Tax influences the transition from G1 to S phase and impairs the activity of the DNA-damage sentinel at this junction.^64,65 Furthermore, Tax functionally inhibits the human mitotic checkpoint protein MAD1.²³ These functional characteristics of Tax may explain its mutagenic effect on cellular chromosomal DNA, as recently evidenced in vitro.²⁵

In addition to Tax, the more recently described accessory proteins of HTLV-1 might also account for the pathogenesis of the infection.⁶⁶ Indeed, open reading frames (ORFs) pX-I and pX-II encode proteins produced from both single- and double-spliced transcripts. The double-spliced pX-I and pX-II transcripts encode Rof and Tof proteins, respectively, whereas the single-spliced pX-I and pX-II RNAs encode the p12^I protein consisting of the last 98 residues of Rof, and the p13^II protein corresponding to the last 87 residues of Tof, respectively. Neither pX-I nor pX-II proteins are required for virus replication in vitro.⁶⁷ However, both are important in vivo, since the HTLV-1 p12^I protein and the Tof protein of HTLV-2 are required for the establishment of a persistent infection in rabbits.^68,69 In humans, circulating specific pX-I and pX-II CD8⁺ T lymphocytes are regularly observed, whatever the clinical status.⁷⁰ This indicates that pX-I and pX-II are synthesized in vivo. Recent works have demonstrated that by activating nuclear factor of activated T cells, p12^I is critical for viral infectivity in quiescent primary lymphocytes.^71,72,73

Somatic mutations of the provirus

The data summarized above indicate that in vivo, HTLV-1 replicates mainly through persistent host cell proliferation in a context of Tax-induced genetic instability. In a recent work, we investigated whether this could account for HTLV-1 genetic variability in vivo.¹⁸ To this end, we used a strategy able to distinguish reverse transcriptase (RT)-associated mutations from somatic substitutions. Figure 6 represents two clones of HTLV-1-infected cells. All the cells of a given clone derive from a single infected progenitor and therefore share the same integration site. As there is no preferential target site for HTLV-1 integration in vivo, the characterization of an HTLV-1 integration site can be used as the specific and unique signature of a given clone. Consequently, the comparison of HTLV-1 proviral sequences bordered by different flanking sequences reveals RT-associated mutations, while the comparison of HTLV-1 proviral sequences sharing the same integration site, ie belonging to the same clone, reveals somatic mutations.

Figure 6.

RT-associated mutation versus somatic mutation. Two clones of HTLV-1-infected cells are represented. Bottom: two 3' extremities of the HTLV-1 provirus flanked by their integration sites (see text for details).

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An inverse polymerase chain reaction (IPCR) strategy was designed to distinguish somatic mutations from RT-associated substitutions (Figure 7). The proviral sequences were isolated, together with flanking cellular sequences. A total of 208 clones encompassing the 3' RU5 sequences of HTLV-1 (379 bp, 80 kb of data) and their flanking cellular sequences (mean, 119 bp; range 7 to 362 bp, 23 kb of data) were sequenced. Sequences obtained could be arranged into 29 cellular clones, depending on their cellular flanking sequences. For 60% of the clones, 8% to 80% of infected cells harbored a mutated HTLV-1 provirus, without evidence of reverse transcription-associated mutations. Typical results of sequence alignment corresponding to a DNA sample derived from a patient with TSP/HAM are shown in Figure 8. Overall, not all RU5 sequences derived from the same cellular clone were identical – 18 of 29 clones (60%) were not homogeneous. The number of variants per RU5 sequence ranged from 0 to 4. A total of 38 single-base substitutions and eight single-base deletions were scored. Some sequences harbored up to 3–4 mutations.

Figure 7.

LMPCR and IPCR analyses of HTLV-1 integration and clonal expansion.

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Figure 8.

Somatic mutations of the HTLV-1 3' RU5 sequence in vivo in a patient with TSP/HAM. Seven distinct HTLV-1 3' integration sites were isolated from this patient; the corresponding flanking cellular sequence hexameric repeats, generated during integration, are given on the right. RU5 sequences were aligned according to patient RU5 sequence. Deletions are denoted by a dash (-). Coordinates of the sequence are those of the ATK-1 sequence.³⁵ 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. For each cellular clone, the number of non-unique RU5 consensus sequences is indicated between brackets (adapted from Ref. 18).

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Figure 9 summarizes the different steps at which a mutation can occur during the synthesis of HTLV-1 provirus. Accordingly, the distribution of the 46 RU5 mutations described above indicates that those mutations did not correspond to minus-strand synthesis-associated RT errors, which are expected to result in a homogeneous population of RU5 sequences (Figure 9B). 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 RT-associated substitutions occurring 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 (Figure 9). However, two aspects of our results rule out the second possibility. First, the actual RU5 mutation frequency is incompatible with those RT-associated errors. By using a HTLV-1 single cycle replication assay for mutation detection, the in vivo mutation rate for HTLV-1 was determined by Mansky et al to be 7 10^-6 mutations per target base pair per replication cycle.⁴⁴ Indeed, if the mutations shown in Figure 8 corresponded to plus-strand synthesis-associated errors, they would necessarily be the result of a single cycle of reverse transcription. Accordingly, the average RU5 mutation frequency was 4.2 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 for HTLV-1 RT⁴⁴ and 100 times higher than for HIV RT.⁴⁶ Second, some RU5 sequences harbored two to four substitutions, a mutation frequency not observed to date for a single step of RT-mediated DNA elongation. Therefore, it appears that most substitutions are not due to plus-strand synthesis-associated RT errors nor to PCR artifacts or recombinations. They result from somatic mutations during cellular replication. These data indicate that HTLV-1 in vivo variation results mainly from post-integration events that consist of somatic mutations of the proviral sequence occurring during clonal expansion.

Figure 9.

HTLV-1 3' RU5 sequence mutations at different stages of HTLV-1 replication. Different steps in HTLV-1 RU5 sequence replication that may affect retroviral mutation rates are illustrated. (A) Reverse tanscription and clonal expansion without mutation. (B) Reverse transcriptase (RT)-associated substitution during minus-strand synthesis. The plain circle represents a mutation that has occurred during the synthesis of the 5' RU5 of the minus strand. This substitution appears to be harbored by both strands of the two LTR of the integrated provirus. Accordingly, all infected cells from the corresponding clone (identified by their common integration site) harbor a provirus with the same substitution. As a consequence, all the sequences from this clone obtained after inverse PCR harbor the same mutation at the same position. (C) RT-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 (black square). 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 of (B)), such substitution results into two populations of infected cells: one harbors the RU5 substitution, the other one harbors the native RU5 sequence. As a consequence, two sequence populations are obtained after inverse PCR (line 10). (D) Somatic mutation of the 3' RU5 sequence in the absence of RT-associated substitution. In this case, no substitution occurs during provirus synthesis. After integration, a 3' RU5 substitution that has occurred during the second mitosis is represented at the bottom. As is the case for plus-strand substitution, such somatic mutation results in a double population of RU5 sequences. The distinction between somatic mutations and plus-strand-associated substitutions is explained in the text.

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Consequences of somatic mutations for the virus

Mutations were found to be generally distributed in a random manner across the HTLV-1 RU5 region, with no evidence of hot spots. The rex responsive element (RXRE) is a cis-acting RNA element required for Rex function that maps to the U3R region of the LTR.^{74,75,76,77,78} HTLV-1 Rex protein acts at the post-transcriptional level to induce the appearance of unspliced and singly spliced viral mRNA in the cytoplasm.⁷⁹ Rex action requires both the overall secondary structure intrinsic to the RXRE and specific sequences from one small region of this large structure^74,80,81 which forms a binding site for the protein. Mutational analyses of RXRE have demonstrated that in vitro Rex binding correlates with in vivo Rex function.^76,81

Of the 46 mutations, 22 mapped to 21 sites in the RXRE. As can be seen from Figure 10A, some mapped to the Rex binding site or else disrupted RNA secondary structures, as evidenced by de novo calculations using the variant sequences (Figure 10B). As RXRE plays a crucial role in the expression of HTLV-1, it is likely that a number of these variants will have impaired the expression of viral structural proteins. It will be interesting to confirm these results by testing the efficiency of present RXRE sequences for exporting mRNA.

Figure 10.

Somatic mutations within the RXRE secondary structure. (a) RXRE corresponds to consensus sequence of patient P1 (see Figure 8). Position 1 corresponds to position 8615 of the ATK sequence.³⁵ The three Rex binding motifs are boxed^{74,75,76,77,78} (adapted from Ref. 18). The Figure represents the 22 substitutions (del, deletion). (b) Mutated RXRE RNA secondary structures, as predicted by de novo calculations with ESSA-SAPSSARN software^101,102 using the variant sequences.

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Somatic mutation of cellular sequences

Frequent somatic mutations were also found in the flanking sequences, suggesting that both the provirus and the cellular genome undergo the same process of genetic instability. A total of seven substitutions were found in 24 kb of cellular DNA, representing a frequency of 2.8 10^-4 per bp sequenced. This is remarkably similar to results obtained for the HTLV-1 RU5 region (5.8 10^-4 per bp sequenced), particularly in light of the very different base composition of the two regions (HTLV-1 40% AT, cellular 59% AT). These values are mutation frequencies, not mutation rates, as 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 a remarkable mutation load of 1.7 10⁶ mutations per diploid cell (2.8 10^-4–6 10⁹), or approximately one mutation per 3.5 kb.

A cellular clone derived from a TSP/HAM sample resulted in five sequences; the proviral flanking sequences showed virtual identity with the promoter region of the human -, or non-neuronal, enolase gene.⁸² Proviral integration occurred 146 bp 5' to the TATA box so uncoupling it from a number of transcriptional motifs such as CCAAT box, AP1 and PEA2 sites. All five sequences harbored a TC transition at position 1217 with respect to the published sequence, while one carried two additional purine–purine transitions. The T1217C transition was found to be restricted to clone P1-7, indicating that it represented a somatic mutation. 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, indicating that the somatically mutated clone persisted over time.

Somatic mutation frequency is proportionate to the degree of infected T cell proliferation

It has been clearly established that genomic instability is characteristic of some cancers.¹ However, somatic mutations are not restricted to ATLL samples. When a somatic mutation is associated with DNA replication, its extent generally depends on the number of rounds of mitosis, which may be reflected by the clonal frequency in vivo. It is possible to get an approximate estimate of clonal frequencies from quadruplicate inverse PCR.^14,83 Figure 11a shows that the average number of mutations per kilobase in 57 RU5 sequences belonging to six HTLV-1 clones with a detection frequency of 1 per 150 PBMCs was more than twice that of the remaining 151 sequences derived from 23 clones with a detection frequency of <1 per 150 PBMCs. This difference was statistically significant (P = 0.037). Indeed, since the frequency of abundant circulating clones varies in the order ATLL > TSP/HAM > asymptomatic carriers, the number of acquired somatic mutations was higher in HTLV-1-associated disease than in virus carriers. As shown in Figure 11b, PBMC DNA from ATLL, TSP/HAM, and asymptomatic carriers harbored a mutation frequency of 0.71, 0.52 and 0.22 substitution per kb of sequence (RU5 sequences plus integration sites), respectively. The difference between asymptomatic carriers and ATLL patients was statistically significant (P = 0.048). However, the frequency of somatic mutation in the 45 sequences derived from the six non-malignant ATLL clones was identical to that of TSP/HAM sequences: 0.5 substitution/kb of sequence (RU5 sequences plus integration sites).

Figure 11.

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 HTLV-1 positive clones was estimated by quadruplicate inverse PCR analysis (Figure 2). (b) Distribution of the frequency of somatic mutations along both the RU5 and flanking sequences according to the clinical status (from Ref. 18).

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HTLV-1 infection as a model of premalignant condition

Figure 12 summarizes the data detailed above. The link between mutation frequency and the degree of infected T cell proliferation fits well with a somatic process. As shown in the figure, both result from the effect of Tax, which is also the main target for the robust cellular immune response that characterizes HTLV-1 infection in vivo. The proliferation of infected cells increases the proviral load and allows the virus to disseminate through body compartments such as the CNS in the case of TSP/HAM. Somatic mutations account for HTLV-1 genetic variability and infected cell genetic instability. HTLV-1 variations might modify Tax epitope and thereby generate mutants with a proliferative advantage. Such mutants were recently isolated in the tumor DNA from ATLL patients.⁸⁴ Cellular genetic instability constitutes a propitious background for the accumulation of critical mutations leading to malignancy. The proposed model implies that any factor impairing cellular immune response triggers both infected T cell proliferation and genetic instability, thereby increasing the risk of developing ATLL. This is supported by clinical observation and experimental investigation showing that the suppression of HTLV-1-specific cellular immune response leads to the development of ATLL.^{85,86,87,88,89} In a more general manner, any factor that stimulates T cell proliferation in asymptomatic carriers may constitute a cofactor of ATLL. Recent works suggest that Strongyloides stercoralis might be one of these factors.

Figure 12.

Pathogenesis of HTLV-1 replication in vivo.

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Strongyloides stercoralis as a cofactor of ATLL

Strongyloides stercoralis (Ss) causes strongyloidiasis, a chronic, usually asymptomatic, gastrointestinal infection,^90,91 found in the same endemic area as HTLV-1.⁹² The incubation period that precedes ATLL is significantly shortened in HTLV-1 carriers with Ss infection, suggesting that Ss could be a cofactor of ATLL. We recently investigated the effect of Ss infection on infected T lymphocytes in HTLV-1 asymptomatic carriers in vivo. After real-time quantitative PCR, the mean circulating HTLV-1 proviral load was more than five-fold higher in HTLV-1 carriers with strongyloidiasis than in HTLV-1+ individuals without Ss infection (P < 0.009). This increased proviral load was found to result from the extensive proliferation of a restricted number of infected clones, ie from oligoclonal expansion, as evidenced by the semiquantitative amplification of HTLV-1 flanking sequences. The positive effect of Ss on clonal expansion was reversible under effective treatment of strongyloidiasis in one patient with parasitological cure, whereas no significant modification of the HTLV-1 replication pattern was observed in an additional case with strongyloidiasis treatment failure. Therefore, Ss stimulates the oligoclonal proliferation of HTLV-1-infected cells in HTLV-1 asymptomatic carriers in vivo. Since the degree of infected T cell proliferation correlates with the frequency of somatic mutations, this finding is thought to account for the increased risk of ATLL and the shortened period of latency observed in patients with strongyloidiasis.

The positive effect of a parasitic infection on the development of a lymphotropic virus-associated lymphoma is reminiscent of the well-known synergetic association between Epstein–Barr virus (EBV) infection and malaria in the genesis of Burkitt lymphoma (BL).^93,94 Just as ATLL and HTLV-1, BL has a complex, multistep pathogenesis, in which EBV is the best documented, but almost certainly not the only, contributing factor. Epidemiologically, hyperendemic malaria increases the incidence of BL by a factor of 20 in Africa.⁹³ Concerning HTLV-1, the incidence of strongyloidiasis seems to be higher in ATLL patients than in asymptomatic HTLV-1-infected individuals. In Martinique, the French West Indies, the prevalence of Ss infection in the general population averages 2%, whereas 20% of Ss-infected individuals are coinfected by HTLV-1.⁹⁵ In the same area, the prevalence of Ss infection in ATLL patients is 42% versus 23% in patients with other T cell lymphomas.⁹⁶ Pathogenically, EBV-associated lymphomagenesis is assumed to result from the malaria-associated chronic stimulation of germinal center cell proliferation, together with a possible acquired EBV-specific T cell immune deficiency. In the case of ATLL, present results show that Ss increases the proliferation of HTLV-1 infected T cells by a mechanism which remains to be elucidated.

Present data emphasize the need to assess the effect of other endemic pathogens on the clonality pattern of HTLV-1-infected cells, especially in areas where ATLL incubation period is short. Strongyloidiasis is asymptomatic in about 50% of the cases.^90,91 Future experiments are needed in order to confirm that the positive effect of Ss on clonal expansion is reversible under effective treatment of Ss infection. This will help confirm that the down-regulation of both HTLV-1 proviral load and infected T cell proliferation under treatment results from parasite clearance. Validating this hypothesis will allow strongyloidiasis treatment as an effective prophylaxis of ATLL in HTLV-1 carriers.

Conclusions and perspectives

During normal somatic cell mitosis, the parental cell duplicates its genome, containing approximately 6 10⁹ nucleotides, then partitions its DNA equally between the two newly generated daughter cells. Errors occurring in the course of these processes result in mutations, which is infrequent since normal human somatic cells accurately replicate their DNA every time they divide.¹ The overall mutation rate of somatic human cells has been estimated at 1.4 10^-10 nucleotides/cell/division or 2.0 10^-7 mutations/gene/cell division.⁹⁷ The mean mutation frequency of 10⁶ per HTLV-1 infected lymphocytes¹⁸ might account for the oncogenic potential of the virus. Yet, a conundrum remains: given the mutational pressure of elevated levels 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. Besides, the frequency of neo-antigen formation could be higher than usually found in other malignancies. Both features might allow robust control by host cell-mediated immunity. As detailed above, suppressing the anti-HTLV-1 cellular immune response contributes to the development of ATLL, whereas restoring it might have a curative effect in animal models,⁹⁸ as in humans.⁵⁶ How could somatic mutations help an infected cell escape CTL response? The generation of Tax escape mutants is the first explanation that comes to mind. The expression of HTLV-1 proteins marks out the cell as non-self and as a target for cell-mediated immunity, which is particularly intense.⁴⁹ Only deletions of single nucleotides were noted. However, if they occurred in the viral open reading frames they would considerably limit protein expression. Negative selection then ensures the maintenance of the Tax+ phenotype. As Tax is a target for cytotoxic T lymphocytes, some mutations in HLA-class I-restricted CTL epitope are likely to be compatible with Tax function, thus allowing the cellular clone to escape cell-mediated immunity.⁹⁹ This is possible as long as most other HTLV-1 protein sequences are inactivated first, as revealed by the observation that cases of CTL escape seem to occur only when high cellular immune responses are focused on a single target.¹⁰⁰ Indeed, tax sequences corresponding to escape mutants have been observed in the tumor DNA from ATLL patients,⁸⁴ suggesting that, in these cases, the tumor clone would have derived from an infected cell having escaped CTL response. In addition to the generation of Tax escape mutants, the huge excess of somatic cellular mutations might alter the expression of genes involved in antigen presentation. Indeed, one can speculate that the CTL pressure can select clonally expanded cells with acquired mutations impairing the proteasome, the transport of the peptide or MHC–peptide interactions. Future studies allow exploration of this question.

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Acknowledgements

This work was supported by grants from the Association pour la Recherche sur le Cancer, the Fondation Contre la Leucémie, the Comités Départementaux du Rhône, de la Saone et Loire, de l'Ardèche et de la Savoie de la Ligue Nationale Contre le Cancer, and from the Programme de Recherche Fondamentale en Microbiologie et Maladies Infectieuses et Parasitaires of the French Ministry of Health. FM was the recipient of bursaries from the Ministère de l'Enseignement Supérieur et de la Recherche, from the Fondation pour la Recherche Médicale and from the Fond National pour la recherche scientifique (Belgium). ASG is supported by a grant from the Centre Léon Bérard. We thank Christiane Pinatel, Agnes Lançon, Céline Paul and Marie-Dominique Reynaud for assistance.