Affiliation of authors: Program in Infectious Diseases, School of Public Health, University of California, Berkeley.
Present address: S. M. Philpott, Department of Infectious Disease, Wadsworth Center, New York State Department of Health, Albany.
Correspondence to: Gertrude C. Buehring, Ph.D., Program in Infectious Diseases, School of Public Health, University of California, Berkeley, Berkeley, CA 94720 (e-mail: buehring{at}uclink4.berkeley.edu).
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ABSTRACT |
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INTRODUCTION |
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Most oncogenic retroviruses induce cellular transformation through one of two mechanisms (2). Acutely transforming retroviruses, such as Rous sarcoma virus (RSV), carry a transduced cellular proto-oncogene, giving the virus transcriptional control over the oncogene. Expression of these genes at inappropriate times, at unacceptable levels, or in unsuitable cell types causes rapid induction of tumors through dysfunction of biochemical pathways regulating cellular growth. The poorly transforming nonacute retroviruses, such as avian leukosis virus, induce malignancies by insertional mutagenesis. Proviral DNA integrates close to cellular proto-oncogenes, stimulating excess or inappropriate transcription of neighboring genes and disrupting control of growth processes. Malignancies occur infrequently and only after long latent periods.
HTLV/BLV group viruses do not appear to use either of these methods. These viruses contain a transforming gene, tax, not homologous to cellular proto-oncogenes, and there is no preferred site of integration. The molecular mechanism by which the encoded protein Tax induces cellular transformation is not known, but most studies have focused on its interaction with cellular proto-oncogenes and tumor suppressor genes (3). Tax has been shown to increase expression of cellular genes involved in the regulation of lymphocyte growth, including interleukin 2 and its receptor (4-9), and to interact with putative tumor suppressor proteins like p53 (10-12) and Int-6 (13,14).
Alternatively, we have found that the Tax protein of HTLV/BLV group viruses imbues infected cells with a "mutator phenotype." Cancers develop through a series of well-defined steps, changing progressively from normal cells into premalignant cells and then into localized tumors and metastatic lesions. Even after exposure to strong carcinogens, the mutation rate in normal cells is too low to account for the myriad of alterations that accumulates during tumor progression. Several researchers (15,16) have argued that the earliest step in oncogenesis is a change that increases the cellular mutation rate. When compared with the mutation rate seen in primary cultures of normal human cells, the mutation rate in transformed cells is up to 1000-fold higher (17). This higher mutation rate may be due to decreased replication fidelity or inhibition of DNA damage repair.
Many hereditary disorders involving defective DNA repair are characterized by a higher frequency of certain types of cancers. For example, patients with xeroderma pigmentosum (XP) lack at least one of the enzymes responsible for repairing UV radiation-induced DNA damage. XP patients exhibit 100-fold greater susceptibility to skin cancers (18). Some cancers induced by DNA viruses are also suspected of involving deficient DNA repair because of frequently observed chromosomal instability in infected cells (19-23).
Chromosomal abnormalities are also common in HTLV-1- and BLV-infected leukemia cells (24,25) and may mirror disease severity, suggesting that these changes are important events in the progression to cancer (26). Cells immortalized in vitro with HTLV-1 exhibit similar chromosomal aberrations and enhanced sensitivity to genotoxic chemicals, as measured by increased frequency of micronuclei following treatment with the chemicals (27,28). Finally, expression of human ß-polymerase (an enzyme involved in DNA excision repair) is decreased in HTLV-1-infected cells, suggesting that HTLV-1 might inhibit DNA repair in infected cells (29).
In this article, we examine the hypothesis that one mechanism by which HTLV/BLV group retroviruses may transform infected cells is by inducing genomic instability, most likely by inhibiting cellular repair of spontaneous DNA damage.
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MATERIALS AND METHODS |
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The 38 cell lines used for this study are summarized in Table 1. Cell
lines were maintained as follows: most of the monolayer lines in Dulbecco's modified
high glucose Eagle's medium (Life Technologies, Inc. [GIBCO/BRL],
Gaithersburg, MD); lymphoid-derived suspension lines in RPMI-1640 medium (Life
Technologies, Inc.); Chinese hamster ovary (CHO) lines in alpha-modified minimal essential
medium (Life Technologies, Inc.); and Hep3B line in Eagle's minimum essential medium
(Life Technologies, Inc.) with 0.1 mM nonessential amino acids and 1.0 mM
sodium pyruvate. All media were supplemented with 100 µg/mL streptomycin (Pfizer, New
York, NY), 2 mM L-glutamine, 10 µg/mL insulin, 100 U/mL
penicillin, 50 U/mL polymyxin B, and 5%-10% fetal bovine serum (FBS) (all from
Sigma Chemical Co., St. Louis, MO). All cell lines were grown at 37 °C in a moist
atmosphere of 5% CO2. Culture fluids were changed once or twice per week.
Monolayer cells were passaged at confluence. Suspension cells were passaged to maintain a
density of 0.5-1.0 x 106 cells/mL.
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Six plasmids or modifications thereof were used. Plasmid pRSV-tax1 contains the tax gene of HTLV-1, and plasmid pRSV-tax2 encodes HTLV-2 tax, both under the control of the RSV promoter. Plasmid pBLV-tax contains BLV tax under the control of the simian virus 40 (SV40) promoter (43). Plasmid pK30 contains the whole HTLV-1 provirus (44). Plasmid pRSV-CAT contains the CAT (chloramphenicol acetyltransferase) reporter gene under the control of the RSV promoter (45). Plasmid pBluescribe (Stratagene, La Jolla, CA) contains the gene for ß-galactosidase. Vector-only deletion mutants were made of the tax-containing plasmids by cleaving out the tax insert with EcoRI and converting the plasmid back to closed circular form by use of standard methods (46). Deletion mutants were confirmed for the absence of the tax genes by their size, as determined by agarose gel electrophoresis and by negative polymerase chain reaction (PCR) results by use of primers for regions within the respective tax genes. All plasmids contain an ampicillin-resistance gene for selection during propagation and purification. Plasmids were propagated according to standard methods (46) in XL1-Blue strain of Escherichia coli, and plasmid DNA was purified by use of the Wizard(tm) Maxiprep kit (Promega Corp., Madison, WI).
Cell Transfections
The established H9 T-lymphocytic cell line was transfected with the DNA of various
plasmids by electroporation. Plasmid (10 µg) was added to cells suspended at a
concentration of 1 x 107/mL in 0.3 mL of RPMI-1640 medium
supplemented with 10 mM dextrose (Sigma Chemical Co.) and 1 mM
dithiothreitol (Fisher Scientific Co., Santa Clara, CA). After 10 minutes on ice, the mixture was
subjected to a single electrical pulse of 250 V, 960 µF from a GenePulser Transfection
apparatus equipped with a Capacitance Extender unit (Bio-Rad Laboratories, Richmond, CA)
and maintained on ice another 10 minutes. The mixture was then diluted into 5 mL of medium
supplemented with antimicrobials and 15% FBS and incubated at 37 °C for 48 hours
before analysis. Efficient expression of the transfected tax genes was verified by reverse
transcription-PCR analysis (Fig. 1). Cells transfected with pBluescribe or
vector alone (pRSV-tax1, pRSV-tax2, and pBLV-tax with the inserted tax genes removed) were
used as experimental controls.
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These assays were performed to confirm expression of tax genes in transfected cells and the
absence of tax in plasmid deletion mutants. Assays were done according to standard methods (46). Primers used in these assays were synthesized by Operon (Alameda,
CA). Primer sequences are detailed in Table 2.
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These assays were undertaken to detect micronuclei, small perinuclear bodies containing either chromosomal fragments or entire chromosomes that have failed to attach to the mitotic spindle during cell division. Micronuclei are considered to be a marker of gross chromosomal damage. Thirty-six cell lines were assayed (AA8, UV41, EM9, C72, M23, M26, C127I, MCF7, T47D, H9, HL-60, Tb1Lu, Bat2Cl6, FLK, C37Cl1, C72/BLV, C761, M26/BLV, M26/BLV-Cl1, C8166-45, MT2, MT4, clone 19, FeLV-3281, JLSV5, L691D, GR, CMMT, CE/RSV, KC, XC, ID13, Raji, Hep3B, HeLa, and SV40-infected IMR-90 cells). Cultured cells in the logarithmic phase of growth were treated overnight with 3 µg/mL cytochalasin-B added to the culture medium in order to arrest cells in the process of mitosis, the phase during which micronuclei appear. Cells were washed twice with ice-cold Dulbecco's phosphate-buffered saline (DPBS) (Life Technologies, Inc.), placed in hypotonic solution (0.75% sodium citrate) for 10 minutes to swell cells, then fixed in absolute methanol. Monolayer cells had been grown directly on glass slides, whereas suspension cells were harvested onto glass slides after fixation with the use of a Cytospin-II cytocentrifuge (Shandon Inc., Pittsburgh, PA) at 600 rpm for 6 minutes at room temperature. All cell preparations were dried and stained for 5 minutes with 0.01 µg/mL ethidium bromide. Approximately 1000 cells per slide were examined for the presence of micronuclei under green epifluorescent illumination (450-490 nm), and the number of cells with or without micronuclei was scored. Structures were identified as micronuclei if they were 1) similar in shape and staining to, but smaller than, the normal nucleus, 2) nonrefractile, and 3) not linked to the nucleus by a nucleoplasmic bridge (47). Scoring was blind; i.e., all cell cultures were prepared by one of us (G. C. Buehring) and given to the other (S. M. Philpott) as coded specimens for scoring.
The kinetochore assay was performed as described by Majone et al. (27) to determine if the micronuclei observed were due to whole-chromosome loss or to chromosome fragmentation. The presence of a kinetochore (centromere) within a micronucleus suggests that a whole chromosome is present, whereas the absence of a kinetochore suggests that only a chromosome fragment is present. After methanol fixation on glass slides, cells were incubated with anti-kinetochore monoclonal antibody (Chemicon, Temecula, CA) diluted 1 : 1000 in DPBS, then in biotinylated horse anti-mouse immunoglobulin G (Vector Laboratories, Inc., Burlingame, CA) diluted 1 : 1000 in DPBS, and finally in 5 µg/mL fluoresceinated streptavidin (Vector Laboratories, Inc.). Each incubation was done for 1 hour at room temperature; between each step, slides were rinsed three times with DPBS. After these incubations, cells were stained with propidium iodide in an antifade solution (Sigma Chemical Co.). Cells were scored for micronuclei as described above. When a micronucleus was located, the presence or absence of a kinetochore spot in each micronucleus was determined by switching from green to blue illumination (510-560 nm). The number of kinetochore-positive micronuclei, the number of kinetochore-negative micronuclei, and the total number of cells containing at least one micronucleus were all recorded separately. These samples were not coded.
Apoptosis Assays
Apoptotic cell death was measured in cells cultured in 10% FBS-containing maintenance medium, described above in section entitled "Cell Lines and Cell Culture," as well as under conditions of induced apoptosis, namely, serum withdrawal or the addition of the chemotherapeutic drugs doxorubicin or dexamethasone at a final concentration of 10 µM. Apoptotic cell death in HTLV-1-infected cells (C8166-45, MT2, and MT4), BLV-infected cells (Bat2Cl6), STLV-infected cells (KIA), virally transfected cells (H9 cells transfected with the tax-containing plasmids or whole HTLV-1-containing plasmid), and uninfected cells (H9 and Tb1Lu) was measured three ways as described in detail previously (48). Briefly, trypan blue dye exclusion to detect viable cells was performed daily by counting and plotting viable cells versus time. Apoptosis before and after serum withdrawal was evaluated by electrophoretic analysis of genomic DNA. Extraction of DNA was done by digestion of a DPBS-rinsed cell pellet in 20 µL of 50 mM Tris-HCl (pH 8.0), 10 mM ethylenediamine-tetraacetic acid (EDTA), 0.5% (wt/vol) sodium dodecyl sulfate, and proteinase K (Promega Corp.) (0.5 mg/mL) for 1 hour at 50 °C and then 1 hour with 0.1 mg/mL ribonuclease A (RNase A) (Boehringer-Mannheim, Indianapolis, IN) added. We heated samples to 70 °C and added 10 µL of loading solution (10 mM EDTA, 1% [wt/vol] SeaKem\T low-gelling-temperature agarose [FMC BioProducts, Rockland, ME], 0.25% [wt/vol] bromphenol blue, and 40% [wt/vol] sucrose). Samples were subjected to electrophoresis through a 3% agarose gel for 3 hours at 60 V in Tris-Borate-EDTA buffer (1.1 M Tris and 90 mM boric acid [pH 8.4]). Cells were considered to have undergone apoptotic death if they exhibited a characteristic ladder pattern of genomic DNA degradation. Finally, apoptosis was measured as the amount of fragmented DNA in spent culture media after cells were incubated 0-24 hours in the presence or absence of camptothecin, doxorubicin, or dexamethasone, all anticancer drugs that induce apoptosis in actively dividing cells. A commercially available enzyme-linked immunosorbent assay (ELISA) kit to measure the amount of fragmented DNA released from apoptotic cells (Cellular DNA Fragmentation ELISA; Boehringer-Mannheim) was used according to the manufacturer's directions. Optical density of the final ELISA reaction was measured after different durations of drug exposure. Transfected cells were not subjected to apoptosis studies until 48 hours after transfection, so that those cells injured or killed during transfection could be removed before the assays were begun.
Alkali Denaturation Assay
This assay was used to measure the relative frequency of DNA strand breaks in 29 cell lines (AA8, UV41, EM9, C127I, MCF7, T47D, H9, Tb1Lu, Bat2Cl6, C72/BLV, C761, M26/BLV, C8166-45, MT2, KIA, FeLV-3281, JLSV5, L691D, GR, CMMT, F81, CE/RSV, KC, XC, ID13, Raji, Hep3B, HeLa, and SV40-infected IMR-90 cells). Procedures for performing this type of assay have been described in detail by other investigators (49,50). The analysis was blind; i.e., the person performing the analysis (S. M. Philpott) received coded cells from the person growing the cell cultures (G. C. Buehring). Washed cells were suspended at a concentration of 2.0-5.0 x 105 cells/mL in 5 mL of ice-cold DPBS and transferred as 500-µL aliquots into nine prechilled 12 x 75-mm borosilicate glass tubes. The first three tubes (labeled A) represented the control of undenatured double-stranded DNA. To these was added 1.0 mL of a fresh 1 : 1 mixture of 0.1 N NaOH (denaturing solution) and 0.1 M KH2PO4 (stop buffer) (final pH 7.4); then we added 500 µL of bisbenzamide buffer (indicator) (0.16% N-laroylsarcosine (Sigma Chemical Co.), 0.2 M K4P2O7, 0.04 M EDTA, and 1.0 µg/mL bisbenzamide [Hoechst 33258; Sigma Chemical Co.] [pH 7.4]). Bisbenzamide is a fluorescent dye that binds to double-stranded but not to single-stranded DNA. The next three tubes (labeled B) represented the experimental values. Denaturing solution was added, and the cellular DNA was allowed to unwind for exactly 10 minutes under chilled (4 °C), light-free conditions before stop buffer and indicator were added. This set of tubes contained DNA in partially denatured form. The tubes labeled C represented another control, the maximum unwinding that the DNA is capable of attaining. The DNA in these tubes was allowed to unwind for 2 hours under chilled, light-free conditions before the reaction was stopped. Fluorescence was measured with a fluorometer (excitation, 365 nm; emission, 465 nm; narrow band pass). The fraction (F) of double-stranded DNA remaining after 10 minutes of alkaline denaturation was then calculated as follows: F = (mean fluorescence tubes B - mean fluorescence tubes C)/(mean fluorescence tubes A - mean fluorescence tubes C).
Plasmid Reactivation Assay
This assay, used to measure rates of repair of induced DNA damage, has been described by
others (51,52). Plasmid pRSV-CAT harboring the CAT reporter gene
under control of the RSV promoter was used throughout. For UV damage, 50-100 µL of
plasmid DNA (50 µg/mL in TE buffer [10 mM Tris-HCl at pH 8.0 and 1 mM EDTA]) per well of a sterile 24-well tissue culture plate was exposed to an
unfiltered UV lamp for 15-30 seconds at a distance of 1 inch (approximately 2-3 J/m2 per second). For the preparation of deoxyribonuclease (DNase)-nicked DNA, plasmid
DNA was diluted (50 µg/mL in an ice-cold solution containing 50 mM Tris-HCl
[pH 7.8], 5 mM ß-mercaptoethanol, 5 mM MgCl2,
50 µg/mL BSA, and 0.5 Kunitz unit/mL DNase I [Boehringer-Mannheim])
and incubated on ice for 15 minutes. For the induction of psoralen damage, plasmid DNA was
diluted to 50 µg/mL in 500 µL of sterile ice-cold water and then mixed with 20
µL of 4.6 µM psoralen (furo[2,3-g]coumarin) in ethanol,
incubated at 25 °C for 60 minutes, and exposed for 60 seconds to light from a mercury sun
lamp to trigger the psoralen-DNA cross-linking reaction. For the introduction of damage with
quercetin, plasmid DNA was diluted to 50 µg/mL in sterile ice-cold water, mixed with an
equal volume of a solution of 0.4 mM cupric chloride, 0.4 mM quercetin, and
20 mM Tris-HCl (pH 8.0), and then incubated at 37 °C for 60 minutes. Plasmid
DNA was oxidized by dilution to 50 µg/mL in 3% (vol/vol) H2O2 and incubation at room temperature for 30 minutes. Undamaged control plasmids were
treated with the vehicle solutions without exposure to the damaging agents. After all treatments,
the damaged DNA was purified by ethanol precipitation and resuspended in TE buffer to a final
concentration of 500 µg/mL. To verify the extent of damage, we subjected treated DNA to
electrophoresis on a 1.5% agarose gel and compared it with undamaged DNA (Fig. 2, A, B, and C). Target cells were transfected with 25 µg of damaged
or undamaged pRSV-CAT plasmid DNA by electroporation and were incubated at 37 °C
for approximately 40 hours, the time of maximal enzyme expression in transfected lymphocytes (51). Harvested cells were resuspended in 250 mM Tris-HCl (pH
7.8) and lysed by freeze-thawing three times, the total protein was determined (BCA protein
assay kit; Pierce Chemical Co., Rockford, IL), and DNA repair rates were measured as a function
of CAT enzyme activity quantitatively determined by use of a commercially available
CAT-specific ELISA kit (CAT-ELISA; Boehringer-Mannheim).
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The data presented in Fig. 3, A and B (number of micronuclei and
proportion of single-stranded DNA), were analyzed by three group comparison methods (53): one that assumes normal distribution of each group being compared
(Scheffé's method) and two that are nonparametric (Kruskal-Wallis method and
the median counterpart of Fisher's exact test). The nonparametric analyses were performed
with the use of both Monte Carlo and asymptotic approaches. Because our experimental design
took a fixed-effects approach, the inferences apply only to the cell lines that we used. Data in
Fig. 4
(plasmid reactivation assays) were analyzed by the Dunnett t test (53). This is a multiple comparison method that uses a pooled
standard deviation and allows the comparison of multiple values to a single control. The
minimum significance level was P<.05. All tests were two-sided.
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RESULTS |
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Initially, we investigated the ability of HTLV/BLV group viruses to
induce chromosomal abnormalities by use of micronucleus formation as an
indicator of DNA damage (47). Replicate values were obtained
at different times for 36 cell lines. The mean number of micronuclei
for each line was used for subsequent statistical manipulations (Fig.
3, A). Established cell lines infected with HTLV/BLV group viruses had
statistically significantly higher frequencies of micronuclei per 1000
cells than uninfected cells or established cell lines infected with
other types of oncogenic viruses (Scheffé's method,P<.0001; both Kruskal-Wallis test and Fisher's exact
test, P<.001). When assayed 48 hours after transfection,
cell lines transfected with Tax-encoding plasmids gave similar results
(Fig. 3,
A). Cells transfected with the tax-deleted plasmid vectors did
not show an increase in micronucleus frequency.
Role of Apoptosis and Chromosome Loss on Increased Frequency of Micronuclei
Although micronuclei are generally used as a biomarker for DNA damage in genotoxic studies, micronuclei and micronucleus-like structures can arise through several mechanisms: apoptosis, whole-chromosome loss, or chromosomal breakage (47). We first examined the possibility that the increased rate of formation of micronuclei seen in infected cells was due to apoptosis. When analyzed by trypan blue dye exclusion, agarose gel electrophoresis, or cell death immunoassay, there was no significant difference between infected or transfected cells and uninfected or untransfected cells in the degree of apoptosis demonstrated. Furthermore, there was no increased sensitivity to various apoptosis-inducing treatments (serum starvation and addition of camptothecin) (data not shown) (48).
We also eliminated the possibility that the increased rate of formation of micronuclei seen in infected cells was due to virus-induced mitotic loss of whole chromosomes. A modified micronucleus assay using an anti-kinetochore antibody was carried out to distinguish micronuclei with a kinetochore (marker for a whole chromosome) from micronuclei without (27,28). We saw no increase in the frequency of kinetochore-positive micronuclei in infected or tax-transfected cells (data not shown) (48).
Frequency of DNA Strand Breaks in HTLV- and BLV-Infected/Transfected Cells
To confirm that the micronuclei seen in cells infected with HTLV/BLV group viruses arose from DNA-damaging events, we used an alkali denaturation assay to detect the relative frequency of DNA strand breaks in various infected and uninfected cell lines. Commonly used in radiobiology (50), this method is based on the observation that the unwinding rate of DNA molecules in a mild alkali solution increases with prior exposure to ionizing agents. Since DNA strand breaks are believed to be responsible for this increase, the rate of unwinding can be used as a surrogate measure of strand breakage. This method has been used to study the effect of acute herpesvirus infection on the integrity of host cell DNA (49), and our experiments represent an application of this assay to persistently infected, transformed cells.
The greater the rate of decrease of fluorescence after alkali treatment, the greater the
presumed amount of single-strandedness and DNA damage (or DNA repair intermediates)
present. The mean of triplicate values for each cell line was used for subsequent statistical
manipulations (Fig. 3, B). Cells infected with HTLV/BLV group viruses
showed a statistically significant decrease in fluorescence (increase in single-strandedness due to
DNA damage) when compared with uninfected cell lines or with cells infected with other
oncogenic viruses, including retroviruses known to transform cells by insertional mutagenesis or
a viral oncogene (Scheffé's method, P<.0001; Kruskal-Wallis test and
Fisher's exact test, Monte Carlo approach, P<.001; Fisher's exact test,
asymptotic approach, P<.003). In vitro transfection experiments using
Tax-encoding plasmids gave similar results (Fig. 3,
B). Cells transfected
with tax-deleted plasmid vectors did not show a statistically significant difference from
untransfected cells.
Repair of DNA Damage Caused by UV Light, DNase, Psoralen, Quercetin, or Hydrogen Peroxide in HTLV- and BLV-Infected/Transfected Cells
The micronucleus and alkali denaturation results raised the question of whether HTLV/BLV group viruses themselves cause DNA damage or whether they inhibit repair of spontaneous damage. To test the theory that Tax inhibited normal cellular DNA repair processes, a modified plasmid reactivation assay was used to measure repair rates in uninfected, infected, and tax-transfected cells. Cells will repair lesions in exogenous DNA, so it is possible to measure repair of damaged viral or plasmid DNA transfected into cells in order to assess inherent repair capacity (54). This assay has been used to examine the DNA repair capacity of a variety of normal and malignant cell types, as well as to demonstrate an association between diminished DNA repair proficiency and skin cancer susceptibility in individuals with XP (51,52,55,56).
Gel electrophoresis of CAT reporter plasmids treated with the DNA-damaging agents
indicated that, within the sensitivity of the gel assay, all of the DNA was converted from the
undamaged form (supercoiled) to the damaged forms (linear, nicked, and adducted) with
different mobilities (Fig. 2, A, B, and C). The results of experiments
describing cellular DNA repair by plasmid reactivation analysis are described in Fig. 4,
A, B, C, and D). Our initial
experiments measured repair of UV light-induced DNA damage in HTLV/BLV-infected and
uninfected cell lines. Following transfection with all types of damaged CAT reporter plasmids,
uninfected H9 cells showed CAT enzyme activity equivalent to the level seen in cells transfected
with the undamaged plasmid controls. In contrast, HTLV- and BLV-infected cell lines
transfected with plasmids damaged by UV light, quercetin, and hydrogen peroxide showed a
statistically significant decrease in CAT activity, indicating a diminished ability to repair these
types of DNA damage (Fig. 4,
C and D). There was no statistically
significant decrease in their ability to repair damage induced by DNase or photoactivated
psoralen (Fig. 4,
C and D). The same patterns prevailed for cells
transfected with plasmids containing either the BLV or the HTLV tax gene (Fig. 4,
B). Cells transfected with tax-deleted plasmid vectors did not show
diminished repair of DNA damaged by any of the five agents (data not shown). The mutant CHO
cell line EM9, defective in DNA strand break repair, showed significantly decreased repair rates
of DNA damaged by UV light, quercetin, or hydrogen peroxide as compared with the normal
parental cell line AA8 (Fig. 4,
A). The mutant cell line UV41, defective
in nucleotide-excision repair, was unable to repair psoralen damage as compared with the normal
parental cell line AA8 (Fig. 4,
A). These controls indicated that our
plasmid reactivation system was working correctly.
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DISCUSSION |
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Only two non-HTLV/BLV cell lines (i.e., EM9 and Hep3B) showed significant increases in the frequencies of micronuclei and single-strandedness (decreased fluorescence) when compared with the background levels seen in uninfected cells. The EM9 CHO cell line is known to be defective in DNA strand-break repair (58); it was included as a positive assay control. AA8, the normal parental line of EM9, and UV41, a sister clone, defective in nucleotide-excision repair (58-60) did not show an increase in the frequency of formation of micronuclei or strand breakage. They were included as negative assay controls. Hep3B hepatocarcinoma cells are infected with hepatitis B virus (HBV) (30); our observations, therefore, support recent observations that HBV-infected cells are unusually susceptible to genotoxic damage. One study (20) found that the presence of HBV DNA alone increased the frequency of genetic recombination in hepatocarcinoma cells. Other researchers (61-63) have reported data suggesting that the HBV X protein may be involved in inhibiting endogenous repair pathways.
By using a variety of DNA-damaging agents in the plasmid reactivation assay, we sought to pinpoint the particular DNA repair pathway inhibited by the Tax protein of HTLVs and BLV. Mammalian cells use different biochemical pathways to repair different types of damage, including base-excision repair, nucleotide-excision repair, recombination, and direct reversal of damage by DNA ligase (54). Exposure of plasmid DNA to UV light can cause several types of damage. Most lesions are cyclobutane pyrimidine dimers and [6-4] photoproducts, but strong doses of UV radiation can also cause DNA cross-linking, strand breakage, and oxidative damage (54). To ascertain which of these repair pathways might be inhibited in Tax-transformed cells, we used chemical methods to selectively introduce specific lesions into the CAT reporter plasmid.
DNase was used to introduce nicks into the reporter plasmid. These lesions are sealed by DNA ligase (64). Our data suggest that ligase-directed reversal of DNA damage is not inhibited by the Tax proteins of the HTLV/BLV group.
Lesions that can be repaired by direct reversal of DNA damage are fairly rare; most lesions are repaired by processes requiring excision of damaged DNA. Photoactivated psoralen damages DNA by forming monoadducts and interstrand cross-links; monoadducts are probably repaired by nucleotide excision, while the commonly accepted pathway for cross-link repair involves both nucleotide-excision repair and recombination (54,65). In humans, nucleotide-excision repair is the sole pathway for removing bulky adducts, such as pyrimidine dimers and psoralen adducts (66). This explains why cells from individuals with XP and other hereditary nucleotide-excision repair defects demonstrated decreased repair of psoralen damage in plasmid reactivation studies (54,67,68) similar to ours. Our data suggest that nucleotide-excision repair and recombinatorial repair of DNA damage are not significantly affected by BLV or HTLV Tax. The possibility exists, however, that alternative pathways could be responsible for the repair.
Quercetin is a mutagenic flavonoid that damages DNA by introducing single- and double-stranded breaks (69). Although the mechanics of DNA strand break repair are still poorly understood, several studies (70-72) have suggested that single-stranded breaks are repaired by base excision or nucleotide excision, whereas double-stranded breaks are repaired by homologous or illegitimate recombination. Quercetin is a mutagen only under aerobic conditions, suggesting involvement of reactive oxygen intermediates. H2O2 reacts with transition metal ions to generate hydroxyl radicals, which then react with the nitrogenous bases or the backbone-forming sugars of DNA to produce oxidized pyrimidines, oxidized purines, and single-stranded breaks (73). Free-radical quenchers inhibit quercetin-induced DNA scission, thus supporting this model (69). Lesions induced by free radicals are corrected by base-excision repair.
Our data on reactivation of plasmids damaged with quercetin and H2O2 suggest that the DNA repair pathway inhibited in cells infected with viruses of the HTLV/BLV group of retroviruses is normal base excision of oxidative lesions. Results showing no inhibition of psoralen-induced damage in these cells imply, but do not rule out, that nucleotide-excision repair and recombinatorial mechanisms are not affected. The transforming Tax protein characteristic of these viruses seems to be the primary determinant of the DNA repair inhibition, although further experiments (such as an in vitro acellular system to measure DNA repair in the absence and presence of recombinant Tax) are needed to confirm this observation. To our knowledge, this is the first report of inhibition of DNA repair by any retrovirus and offers a new explanation for the riddle of the mechanism of transformation by the HTLV/BLV group of retroviruses. These results open the door to future research that could focus on delineating the molecular mechanisms of Tax inhibition of DNA repair. Tax-induced decrease of ß-polymerase production is one possibility for a mechanism (29). Alternatively, Tax might not directly inhibit base-excision repair. Rather, it might saturate (i.e., overwhelm) this pathway by somehow stimulating overproduction of metabolic oxidants. In future studies, it will perhaps be possible to identify exactly which Tax domain(s) might be responsible for the DNA repair inhibition that we have discovered and what cellular factors they might interact with.
Most studies of retroviral oncogenesis have focused on the role of oncogenes, examining increased or aberrant expression of these genes by insertional mutagenesis or viral transduction. Although our study represents a dramatic departure from this approach, a DNA repair inhibition model does not exclude other models of Tax-induced cellular transformation. Our model simply questions whether the transformation process rests entirely on growth-regulatory mechanisms, as proposed by transactivation and tumor suppressor models. Instead, we believe that gross genomic alterations are also necessary for progression to a neoplastic state and may, in fact, be the initial event. Disturbance of oncogene and/or tumor suppressor gene function might represent a later step. Inhibition of DNA repair and interleukin 2-driven autostimulation could also work synergistically to render infected cells susceptible to multiple mutagenic changes. This model could also explain why HTLV- and BLV-induced malignancies occur in only about 1% of infected individuals and only after a long latent period. Inhibition of DNA repair might prime virally infected cells for secondary mutagenic events, but such events would be rare and would accumulate only over a long period of time.
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NOTES |
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We thank G. Niermann, J. Xiao, A. Kilani, S. Linn, and K. Radke for technical advice and critical comments. We also thank B. Weiser and H. Burger for reviewing manuscript drafts and M. Tarter for advice on statistics. We are grateful to G. Firestone for providing the GR cell line and the pRSV-CAT plasmid, W. Waschman for the pRSV-tax1 and pRSV-tax2 plasmids, L. Willems for the pBLV-tax plasmid, P. Duesberg for the Rous sarcoma virus-infected chick fibroblasts, T. Forte for the Hep3B cell line, M. McGrath for the M23 and M26 cell lines, and K. Radke for the Bat2Cl6 and Tb1Lu cell lines. Other cell lines were obtained from the former Naval Biosciences Laboratory (Oakland, CA) and the American Type Culture Collection (Manassas, VA). The following reagent was obtained through the AIDS Research and Reference Reagent Program, Division of Acquired Immunodeficiency Syndrome, National Institute of Allergy and Infectious Diseases, National Institutes of Health: human T-cell leukemia virus-1 K30 DNA (plasmid pK30) from T. Kindt.
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Manuscript received June 30, 1998; revised March 24, 1999; accepted April 5, 1999.
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