Analysis of Efficiency and Fidelity of HIV-1
(+)-Strand DNA Synthesis Reveals a Novel Rate-limiting Step during
Retroviral Reverse Transcription*
Matthias
Götte
§¶,
Masanori
Kameoka
,
Nathan
McLellan
,
Luciano
Cellai**, and
Mark
A.
Wainberg
From the
McGill University AIDS Centre, Lady Davis
Institute-Jewish General Hospital, Montréal, Québec H3T
1E2, Canada, the Departments of § Experimental Medicine and
Microbiology and Immunology, McGill University, Montréal,
Québec, Canada, and the ** Istituto di Strutturistica
Chimica, Consiglio Nazionale delle Ricerche, I-00016 Monterotondo
Stazione, Rome, Italy
Received for publication, October 5, 2000, and in revised form, November 20, 2000
 |
ABSTRACT |
We have analyzed the efficiency and
accuracy of polymerization at several different stages during the
initiation of human immunodeficiency virus type 1 (HIV-1) (+)-strand
DNA synthesis. This reaction is of particular interest, as it involves
the recruitment by reverse transcriptase of an RNA primer that serves
as substrate for both the polymerase and RNase H activities of the
enzyme. We found that the correct incorporation of the first two
nucleotides was severely compromised and that formation of mismatches
was completely absent at this stage of initiation. Although the
fidelity of incorporations decreased concomitantly with ensuing
polymerization, the elongation of mispaired primers was literally
blocked. Instead, mispaired primer strands initiated a switch from
active synthesis of DNA to premature RNase H-mediated primer removal.
These findings suggest the existence of a fragile equilibrium between
these two enzymatic activities that is shifted toward RNase H cleavage
once the polymerization process is aggravated. Our data show that the initiation of HIV-1 (+)-strand DNA synthesis differs significantly from
reactions involving other primer/template combinations, including tRNA-primed (
)-strand DNA synthesis.
 |
INTRODUCTION |
Alterations in the availability of deoxynucleoside triphosphates
(dNTPs)1 can profoundly
affect retroviral reverse transcription in several ways (1, 2). Low
concentrations of dNTPs have been shown to cause arrest of reverse
transcription of human immunodeficiency virus type 1 (HIV-1) RNA in
non-dividing macrophages and in quiescent T lymphocytes (3). This
arrest resulted in incomplete reverse transcripts that were identified
in the early G1 phase of the cell cycle. DNA synthesis was
eventually re-initiated at later stages, as concentrations of dNTPs
increased (4, 5). Endogenous reverse transcriptase (RT)
reactions revealed that diminished or biased dNTP pools might also
force the insertion of incorrect nucleotides (6, 7). The high error
rate of HIV-1 RT results in about one mistake per round of replication
(8). Together with the lack of proofreading, this is one of the major
reasons for the enormous variability of HIV. This genetic diversity is also linked to the problem of drug resistance since the prolonged use
of available RT inhibitors, including both nucleoside and non-nucleoside analogs, gives rise to mutant enzymes that confer resistance to the administered antiviral drug (9).
It is poorly understood whether fluctuations in dNTP concentrations
affect reverse transcription indiscriminately or whether specific
events in the reaction are particularly sensitive to such variations.
Reverse transcription is complex and involves a variety of different
stages in which the RT enzyme has to accommodate different
primer/template substrates (10, 11). Reverse transcription of HIV-1 is
primed by human tRNA
, which binds
via its 3' terminus to the primer-binding site of the RNA genome.
Synthesis of the first DNA strand, (
)-strand DNA, is initiated from
the binary tRNA/primer-binding site complex and proceeds toward the
5'-end of the template, to yield so-called (
)-strand strong-stop DNA
(12-14). The transcribed RNA of the newly formed RNA/DNA hybrid is
degraded by the RT-associated RNase H activity. RNase H degradation can
be seen at a fixed distance from the primer terminus, in temporal
coordination with DNA synthesis, and, as well, in the absence of DNA
synthesis, independent of the precise location of the 3'-end of the
primer (15-19). This facilitates the release of the strong-stop DNA,
which is transferred to the 3'-end of the RNA template to allow
continuation of (
)-strand DNA synthesis (20). During this process, a
purine-rich fragment near the 3'-end of the genomic RNA,
i.e. polypurine tract (PPT), remains resistant to RNase H
degradation and serves as a primer for (+)-strand DNA synthesis
(21-24). Following its selection, the PPT primer is initially
extended, and specific RNase H cleavage at the RNA-DNA junction removes
the RNA fragment from newly synthesized DNA. (+)-Strand DNA synthesis
is a discontinuous process that may require initiation from a central
polypurine tract as well (25, 26). At a later stage of (+)-strand DNA
synthesis, specific RNase H cleavage must also remove the tRNA from
(
)-strand DNA to facilitate a second strand transfer, which is
necessary for completion of synthesis of proviral DNA. Precise removal
of both primers is important in regard to subsequent steps in the
retroviral life cycle, as these cuts define the ends of the
double-stranded DNA that is integrated into the genome.
Each of these various stages during reverse transcription may be
affected by fluctuations of available dNTP pools in a different fashion. However, the initiation of retroviral (+)-strand DNA synthesis
is unique in that both the polymerase and RNase H activities recruit
the same strand as substrate. Hence, at this stage of the reaction, the
RT-associated RNase H activity may directly influence the
polymerization process.
We have recently studied the interplay between both active sites under
saturating dNTP concentrations and demonstrated that a specific pausing
site at position +12 correlated precisely with the emergence of RNase H
cleavage at the RNA-DNA junction that removes the RNA primer
from newly synthesized DNA (23). In the present study, we have
investigated the impact of diminished and biased dNTP pools on
efficiency and accuracy of the polymerization process and the
consequences regarding the interdependence between the two active sites
of RT. Steady-state kinetics revealed severely diminished rates of
nucleotide incorporation and unusually high fidelity during the initial
steps of the reaction. Limited concentrations of dNTPs promoted a
switch from active DNA synthesis to RNase H activity and caused
premature removal of the primer before the enzyme reached the pause
site at position +12. As a result, the lower limits of dNTP
concentrations required to effectively and accurately initiate
(+)-strand DNA synthesis are significantly higher compared with those
identified in reactions that involve DNA/DNA primer/template substrates.
 |
EXPERIMENTAL PROCEDURES |
Nucleic Acids and HIV-1 RT--
The oligonucleotides used
in this study were chemically synthesized using the phosphoramidite
method. We utilized a PPT-derived RNA primer (17R), its DNA counterpart
(17D), and chimeric DNA-RNA primers (1D-17R and 3D-17R) that correspond
to different stages during synthesis of (+)-strand DNA;
"nD" and "nR" refer to numbers of DNA and
RNA residues, respectively. The sequences of all oligonucleotides used
are as follows (RNA residues are in italics, and the region of the
template (57D) that provides complementarity to the RNA primer is in
boldface): 17R, 5'-UUAAAAGAAAAGGGGGG-3'; 1D-17R, 5'-UUAAAAGAAAAGGGGGGA-3'; 3D-17R,
5'-UUAAAAGAAAAGGGGGGACT-3'; 17D,
5'-UUAAAAGAAAAGGGGGG-3'; 20D, 5'-UUAAAAGAAAAGGGGGGACT-3'; and
57D,
5'-CGTTGGGAGTGAATTAGCCCTTCCAGTCCCCCCTTTTCTTTTAAAAAGTGGCTAAGA-3'.
The crude oligonucleotides were purified on 7 M urea and
12% polyacrylamide gels containing 50 mM Tris borate (pH
8.0) and 1 mM EDTA. 5'-End labeling of oligonucleotides was
performed with [
-32P]ATP and T4 polynucleotide kinase
(Life Technologies, Inc.). 3'-End labeling was performed with
[
-32P]dATP using the RNase H-deficient RT-E478Q
mutant. For this purpose, 100 ng of the pure RNA primer, pre-annealed
to the template, was incubated at 37 °C for 30 min with
[
-32P]dATP and 1 µg of enzyme. The radiolabeled
products were again electrophoretically purified to obtain
homogeneously labeled nucleic acids. Wild-type HIV-1 RT and the RNase
H-deficient mutant enzyme were purified as described (27).
Primer Extension Reactions--
Km and
kcat values were determined using gel-based
assays according to procedures previously described (28). In all
experiments, primer/template sequences were heat-annealed prior to
reaction initiation. The prehybridized nucleic acid substrate (100 nM) was preincubated with HIV-1 RT (10 nM) in
buffer containing 50 mM Tris-HCl (pH 7.8), 50 mM NaCl, and appropriate dNTP combinations, which allowed
us to monitor the efficiencies of correct and incorrect nucleotide
insertions at positions +1, +2, +3, and +6. Kinetic parameters for
position +1 were determined in "standing-start" experiments, in
which the correct dNTP or the incorrect nucleotide was added at
different concentrations to the reaction mixture. All other positions
were monitored in "running-start" experiments, which measure the
efficiency of nucleotide incorporations at template positions distant
from the primer terminus (28).
Concentrations of correct dNTP that were required to allow DNA
synthesis just before the template position of interest has been
reached were kept constant between 10 and 50 µM. These
concentrations were high enough to ensure efficient incorporation, but
did not exceed the critical limit that might give rise to misinsertions at the following position. For instance, 50 µM was
required for efficient dATP incorporation at position +1 using the pure
RNA primer, whereas 10 µM was sufficient in the case of
the DNA primer. These conditions allowed us to monitor correct and
incorrect nucleotide insertions at position +2 using varying
concentrations of dCTP or dTTP (see Fig. 1A). Reactions were
initiated by the addition of MgCl2 (6 mM) and
carried out for 8 min at 37 °C. These experiments were repeated at
least twice. DNA synthesis was stopped by the addition of EDTA at a
final concentration of 40 mM. Following precipitation with
ethanol, reaction products were visualized on 7 M urea and
15% polyacrylamide gels. Data analysis was as described by Boosalis
et al. (28).
The lower limits of dNTP concentrations required to initiate (+)-strand
DNA synthesis were determined in a similar fashion. Three dNTPs were
used at a high saturating concentration of 100 µM,
whereas the concentration of the fourth dNTP of interest was varied
between 0 and 10 µM. Reactions were performed using an excess of enzyme (200 nM) over the primer/template
substrate (100 nM) to maximize efficiency of DNA synthesis.
Polymerization reactions were stopped after 60 min and analyzed as
described above. Time course experiments were also performed using a
2-fold excess of enzyme over primer/template to monitor appearance of
products during a 60-min period. For this purpose, pre-annealed
primer/template substrate (100 nM) was incubated with 200 nM enzyme. DNA synthesis and RNase H degradation were then
initiated by the addition of MgCl2 at a final concentration
of 6 mM. Reactions were performed at 37 °C and stopped
at different times by adding 1-µl aliquots of the reaction mixture to
9 µl of 95% formamide containing 40 mM EDTA.
Pyrophosphorolysis--
To monitor pyrophosphorolysis, the
pre-annealed RNA/DNA or DNA/DNA substrates (100 nM) were
preincubated for 5 min at 37 °C with the RNase H-deficient mutant
enzyme (30 nM) in buffer containing 50 mM
Tris-HCl (pH 7.8), 50 mM NaCl, and 6 mM
MgCl2. Pyrophosphorolytic cleavages were initiated by
adding various concentrations of PPi to the reaction
mixture. Reactions were performed at 37 °C and stopped after 45 min
by adding 1-µl aliquots of the reaction mixture to 9 µl of 95%
formamide containing 40 µM EDTA. Samples were analyzed as
described above.
 |
RESULTS |
Experimental Design--
We recently devised an in
vitro system that allows the study of structure-function
relationships of HIV-1 RT at various stages of initiation of (+)-strand
DNA synthesis through use of select primer/template combinations (23).
This system has now been used to determine kinetic parameters regarding
the efficiency of correct and incorrect nucleotide additions. The
PPT-derived RNA primer (17R) has allowed us to study steady-state
kinetics at early stages of the reaction, whereas the chimeric
DNA-RNA primer (3D-17R), which corresponds to the addition of
the first three nucleotides to the PPT primer, was employed to study
nucleotide insertions as the enzyme accommodates increased numbers of
DNA residues (Fig. 1A).
Utilizing gel-based assays, we determined Km and
kcat values with respect to nucleotide
incorporations at positions +1, +2, +3, and +6. Reactions were
monitored with 5'-end-labeled primer strands; and unless otherwise
indicated, the primer/template substrate was used at a 10-fold excess
over RT to ensure that reaction conditions fulfilled steady-state
criteria. To minimize the complexity of the assay, we initially used an RNase H-deficient mutant RT, i.e. RT-E478Q (29), to avoid
cleavage of the extended RNA primer. These experiments were also
performed with homologous DNA primers for purpose of comparison.

View larger version (28K):
[in this window]
[in a new window]
|
Fig. 1.
Reverse transcriptase kinetics at early
stages during initiation of (+)-strand DNA synthesis.
A, use of RNA/DNA primer/template combinations and
experimental design. The pure RNA (17R) and the chimeric DNA-RNA primer
(3D-17R) were employed to determine kinetic parameters at positions +1,
+2, +3, and +6 as described under "Experimental Procedures." All
experiments were conducted with the RNase H-deficient RT-E478Q mutant
enzyme. B, efficiency of pyrophosphorolysis monitored with
primers 3D-17R (left panel) and 20D (right
panel). The excision of terminal residues was analyzed at
concentrations of 10, 50, 100, 500, and 1000 µM
PPi (lanes 1-5, respectively). Lanes
C represent the control in the absence of PPi.
C, incorporation of incorrect nucleotides at position +1
using varying concentrations of dGTP. Reactions with the DNA primer 17D
are shown in the left panel, and the corresponding reactions
with the RNA primer 17R are shown in the right panel.
Lanes C indicate controls in the absence of dNTPs.
Lanes 1-9 show reactions in the presence of 10, 20, 50, 100, 200, 400, 1000, 2000, and 4000 µM dGTP,
respectively. D, misinsertions at positions +1 and +2
using increasing concentrations of enzyme. The primer/template was
incubated with relatively high concentrations of dGTP (500 µM) and increasing concentrations of the RNase
H-deficient RT-E478Q mutant enzyme to monitor incorrect nucleotide
insertions at position +1 (left panel). Position +2 was
analyzed by incubating the preformed complex with the correct
nucleotide (dATP, 50 µM) and 500 µM dTTP,
i.e. an incorrect nucleotide in regard to the second
position (right panel). Lanes 1-6 represent
reactions performed at the following ratios of nucleic acid to RT:
1:0.2, 1:0.5, 1:1, 1:2, 1:3, and 1:5, respectively. C lanes
indicate controls in the absence of dNTPs. Reactions were allowed to
proceed for 20 min.
|
|
Steady-state Kinetics of Correct and Incorrect Nucleotide
Incorporations--
The results of our measurements are summarized in
Table I, and important examples are shown
in Fig. 1. We found that the efficiency of incorporation
(kcat/Km) of the correct nucleotide at position +1 was diminished by ~50-fold when the RNA
primer was used (Table I). The influence of the RNA primer was not
restricted to the terminal RNA moiety since the diminution in rate of
nucleotide incorporation was also seen at position +2. Following these
initial steps, the efficiencies of correct nucleotide insertions
increased significantly at position +3, and both the RNA- and
DNA-primed reactions had similar
kcat/Km ratios once the
enzyme had reached position +6 (Table I). A diminution in usage of the
RNA primer was also noted when monitoring pyrophosphorolysis, i.e. the reverse reaction of nucleotide incorporation (Fig.
1B). The presence of PPi solely facilitated the
excision of the terminal DNA residue of the chimeric primer 3D-17R.
Shorter products were not even seen at a high concentration of
PPi (1 mM) that transferred ~40% of the
3D-17R primer into the 2D-17R fragment (Fig. 1B, left panel, lane 5). In contrast, relatively lower
concentrations of PPi (100-500 µM) were
sufficient to convert 40-60% of the homologous 20D DNA primer into
shorter fragments that contained 19, 18, or 17 residues (Fig.
1B, right panel, lane 4).
View this table:
[in this window]
[in a new window]
|
Table I
Average kinetic parameters for correct and incorrect nucleotide
insertions
Data were determined from experiments shown in Fig. 1. , no mispair
formation; ND, not determined.
|
|
The nature of the nucleic acid substrate has even more pronounced
effects as the enzyme is forced to incorporate incorrect nucleotides.
We studied the formation of G:T mismatches, the easiest mispair to
form, and found that the addition of dGMP was completely blocked at
position +1 when using the RNA primer (Fig. 1C). In contrast, DNA-primed reactions permitted the incorporation of dGMP not
only at position +1 (G:T mispairs), but also at position +2, which
created a second adjacent mispair (G:G). This result shows, in
agreement with previous studies, that mismatches are easily formed and
extended, provided that the primer is composed of DNA (30, 31). To
determine whether the lack of mismatch incorporation, seen with the RNA
primer, is restricted to the first position and might be attributable
to low enzyme concentrations used in these experiments, we next
increased the concentration of RT and analyzed the formation of G:T
mispairs at positions +1 and +2. Although the incorrect nucleotide was
used at a relatively high concentration of 500 µM and
despite a 5-fold excess of enzyme over the primer/template substrate
and the long incubation time (20 min), the formation of mismatches at
these two positions was not observed (Fig. 1D). Thus,
accuracy of DNA synthesis at the first two positions of the initiation
reaction was unusually high. Fidelity decreased to values similar to
those observed with the homologous DNA primer at later stages of DNA
synthesis. Differences in rates of incorrect nucleotide insertions,
with either RNA/DNA or DNA/DNA primer/template combinations, were
almost negligible when reactions were monitored at position +6 (Table
I).
Fidelity during Synthesis of Full-length DNA--
The above
findings suggest that the initiation of (+)-strand DNA synthesis may be
sensitive to decreased or biased dNTP pools. Although our data show
that formation of mismatches may occur at later stage of DNA synthesis,
it is not clear whether such mispaired replication intermediates can be
effectively extended. Therefore, we determined minimum concentrations
of dNTPs needed to allow DNA synthesis to continue beyond these
critical template positions by incrementally increasing the
concentrations of one of the four nucleotides while using the other
three dNTPs at saturating concentrations. For this purpose, we employed
a chimeric primer that represents the PPT-derived RNA primer extended
by a single nucleotide (1D-17R), which enabled us to monitor DNA
synthesis at both initial and later stages. To determine the lower
limits of dNTP concentrations required for full-length DNA synthesis, we used conditions that gave rise to high yields of the product under
saturating nucleotide concentrations. Thus, reactions were allowed to
proceed for 1 h using an excess of the RT-E478Q mutant enzyme
over primer/template (Fig.
2A).

View larger version (17K):
[in this window]
[in a new window]
|
Fig. 2.
Lower limits of dNTP concentrations required
to generate full-length DNA products. A, the
concentrations of three dif- ferent dNTPs were kept constant at saturating levels of 100 µM, whereas the fourth dNTP was used at varying
concentrations between 0 and 10 µM. Lanes
1-11 represent reactions with the following concentrations of the
limiting nucleotide: 10, 5, 2.5, 1.3, 0.63, 0.31, 0.16, 0.08, 0.04, 0.02, and 0 µM, respectively. Reactions proceeded for
1 h using the enzyme in excess over primer/template. DNA synthesis
was monitored using the 3'-end-labeled chimeric primer 1D-17R, which
allowed us to study a broader spectrum of positions after the initially
slower steps. The majority of abortive reaction products are seen at
the level of initiation between positions +2 and +12. Nucleotide
concentrations required to bypass these positions to yield 50% of the
full-length DNA product are summarized in Table II. These data were
obtained from the graphs shown in B. For each of the four
dNTPs, concentrations of 5-10 µM were saturating. The
graphs in C point to differences in product formation seen
in the absence of one of the four dNTPs at low concentrations between 0 and 2 µM.
|
|
Consistent with steady-state kinetics, we detected a striking
difference in regard to the yield of full-length DNA in reactions that
compared nucleotide limitations and positions +2 and +3. First, a
relatively high concentration of dCTP, i.e. >1
µM, was required to bypass position +2, which is the
first incorporation event in this experiment (Fig. 2B and
Table II). In contrast, 0.1 µM dTTP was sufficient to bypass position +3 and to yield 50% of the full-length product compared with reactions conducted under
saturating dNTP concentrations of 10 µM (Fig.
2B and Table II). In either case, the majority of abortive
reaction products were found at these initial steps of DNA synthesis.
Later positions were relatively easy bypassed, unless the limiting
nucleotide affected three or more consecutive bases. For instance, dGTP
was required at a relatively high concentration of ~0.8
µM since low levels of dGTP had the potential to
aggravate DNA synthesis at multiple positions, i.e.
positions 4, 5, and 8-10. In contrast, dATP, which is required at
positions +6 and +7, was needed only at a relatively low concentration
of 0.1 µM to achieve 50% full-length DNA synthesis. The
full-length product was even detected in the absence of either dATP or
dTTP, indicating that mispaired primers can, in principle, be extended,
although the yield was relatively poor.
View this table:
[in this window]
[in a new window]
|
Table II
Concentrations of dNTPs required to initiate synthesis of HIV-1
(+)-strand DNA
Reactions were monitored with the 3'-end-labeled chimeric 1D-17R primer
in the presence of three dNTPs at a final concentration of 100 µM each; the fourth dNTP was added at variable
concentrations (Fig. 2).
|
|
Primer Removal Is Faster than Mismatch Formation and/or
Extension--
Next, we increased the complexity of these assays and
compared results obtained with the RT-E478Q mutant enzyme and with
wild-type RT containing an intact RNase H domain. To analyze the
influence of diminished dNTP pools on the interplay between the
polymerase and RNase H activities of RT, we generated a 3'-end-labeled
chimeric primer corresponding to the addition of a single radiolabeled DNA residue to the PPT primer, as in the experiments shown in Fig. 2.
This type of labeling facilitates the analysis of RNase H-mediated
processing of the newly synthesized DNA strand since the size of the
processed reaction product directly points to the stage of DNA
synthesis at which the RNA primer is removed (23). The temporal
relationship between the polymerase and RNase H active sites of RT was
monitored in a time course reaction in which one of the four
nucleotides was omitted (Fig. 3,
A and B).

View larger version (17K):
[in this window]
[in a new window]
|
Fig. 3.
Effects of biased dNTP pools on interactions
between the polymerase and RNase H active sites of RT.
A, the 3'-end-labeled chimeric primer 1D-17R was employed to
study the effects of RNase H cleavage on polymerization in the absence
of one of the four dNTPs. B, reactions were monitored in
time course experiments using the wild-type enzyme in a 2-fold excess
over primer/template. Arrows on the right point to reaction
products representing processed and unprocessed primer strands.
Lanes 1-8 represent reactions performed for 1, 3, 6, 10, 15, 22, 30, and 45 min, respectively. C, shown is the
DNA-primed reaction in the presence of dCTP, dTTP, and dGTP and in the
absence of dATP. Reactions were stopped after 1, 3, 6, 12, 20, 40, and
60 min (lanes 1-7, respectively). Lane C is a
control in the absence of MgCl2.
|
|
DNA synthesis was completely blocked in the absence of dCTP,
i.e. the correct nucleotide at the second template position
(Fig. 3B, panel 1). Thus, misincorporations did
not take place at this stage of DNA synthesis. However, a diminution in
the chemical step and decreased binding of the incoming incorrect dNTP
are not the only possible factors that block DNA synthesis under these conditions. The time course shows that the RT-associated RNase H
activity prematurely cleaved at the RNA-DNA junction, resulting in
release of a single nucleotide containing the radiolabel. The same
result was obtained with reaction mixtures lacking dTTP or dGTP (Fig.
3B, panels 2 and 3). Here, effective
DNA synthesis required incorporation of incorrect nucleotides at
positions +3 and +4, respectively. DNA synthesis was again blocked even
before the enzyme had reached these critical positions, and major RNase H cleavage products were found to correspond to short DNA fragments containing two and three residues, respectively. Thus, in these cases,
polymerization stopped before an incorrect nucleotide was added, which
finally resulted in premature removal of the primer.
A similar sequence of events was also seen when dATP was omitted from
the reaction mixture (Fig. 3B, panel 4). However,
under these conditions, we did observe incorrect insertions at position +6, although the misaligned primer was not further extended. In this
case, the short cleavage fragment corresponded to a sequence of six DNA
residues, indicating that primer removal was initiated after mismatch
formation at position +6. Collectively, these data demonstrate that the
RT-associated RNase H activity can seriously affect the initiation of
(+)-strand DNA synthesis as the availability of dNTP pools is limited.
The requirement for dATP incorporation at position +7 may be considered
as an additional obstacle that compromises elongation of the mismatched
intermediate. However, the homologous DNA primer was efficiently
extended under the same reaction conditions (Fig. 3C). The
RT enzyme efficiently bypassed critical, consecutive template positions
at which mismatches were formed, which is consistent with previously
published data (30-34) and demonstrates that high rates of
misinsertions and extensions occur when DNA/DNA serves as substrate.
Switching from Active DNA Synthesis to RNase H
Activity--
To further assess the biological significance of
these observations, we next determined the effects of fluctuations of
dNTP concentrations on initiation of (+)-strand DNA synthesis. The time
course in Fig. 4 (left panel)
shows formation of the processed full-length product when dATP was used
at a saturating concentration of 10 µM. In good agreement
with our previous data (23), the first product that appeared in the
time course corresponds to 12 DNA residues and therefore identifies the
primer removal reaction after the enzyme paused at position +12. (The
paused product was not seen under these conditions since the enzyme was
used in excess over primer/template.) The short fragment disappeared
with time as DNA synthesis resumed and the processed full-length
product emerged. In contrast, 100 nM dATP, which was
sufficiently high to produce the full-length product with the RT-E478Q
mutant (Fig. 2 and Table II), blocked DNA synthesis entirely when using
the wild-type enzyme (Fig. 4, right panel). Instead, two
major abortive DNA species were identified. One of these corresponded
to newly synthesized DNA that was cleaved from the RNA primer after RT paused at position +12. The shorter product, which contains six DNA
residues, indicates premature removal of the primer before the enzyme
has successfully passed position +6 (compare with Fig. 3). Another
short product contains five DNA residues and thus indicates RNase H
cleavage before the mispair was formed. Hence, both the natural pausing
site at position +12 and template positions that require incorporation
of the limited nucleotide represent obstacles that can block DNA
synthesis and induce removal of the primer prematurely.

View larger version (27K):
[in this window]
[in a new window]
|
Fig. 4.
Nucleotide concentrations that promote a
switch from active DNA synthesis to premature removal of the
primer. Reaction conditions were as described in the legend to
Fig. 3, except that dATP concentrations were 10 µM
(left panel) and 0.1 µM (right
panel), whereas the other three dNTP concentrations were kept
constant at 100 µM. Lanes 1-9 represent
reactions performed for 1, 3, 6, 10, 15, 22, 30, 45, and 60 min,
respectively.
|
|
Secondary Initiation Events Can Rescue DNA Synthesis--
The
observation that the RNA primer is prematurely cleaved from newly
synthesized DNA does not necessarily mean that DNA synthesis is blocked
in irreversible fashion. We have recently demonstrated that
chain-terminated DNA synthesis can be rescued since RT is capable of
recognizing the 3'-end of the cleaved RNA and eventually initiates a
second round of DNA synthesis (23). Therefore, we analyzed whether
biased dNTP pools may likewise initiate such secondary priming events.
For this purpose, we used the 5'-end-labeled chimeric DNA-RNA
primer (3D-17R) and initiated DNA synthesis in the presence of only
dATP and dGTP (Fig. 5A). The
time course shows that DNA synthesis specifically stopped at position
+10 (Fig. 5, A and B, schematic). The
next template position would then require insertion of an incorrect
nucleotide, which did not take place under these conditions. Instead,
the intensity of this band decreased as RNase H cleavage products
emerged. An additional band that migrated just above the largest
cleavage product appeared at longer reaction times. This is reminiscent
of secondary initiation reactions observed in the presence of
chain-terminating stop nucleotides. DNA synthesis was rescued from the
primary and secondary products, as the missing nucleotides,
i.e. dCTP and dTTP, were included in the reaction mixture.
Extension of the primary product is most likely more efficient since
re-initiation from the RNA primer eventually involves displacement of
the first DNA strand. However, these secondary reactions may be
important, as the efficiency of extension of the primary product is
diminished through incorrect nucleotide incorporations, e.g.
as shown for position +6.

View larger version (21K):
[in this window]
[in a new window]
|
Fig. 5.
Secondary initiation reactions at limiting
concentrations of dNTPs. A, reactions were
initiated from the 5'-end-labeled chimeric primer 3D-17R using only two
of the four dNTPs (dATP and dGTP, 10 µM each). The
omission of dCTP and dTTP requires misinsertions at positions +11 and
+12, respectively, which is not seen in this time course. Instead, the
presence of dATP allows extension of the processed primer by a single
nucleotide to yield a secondary product, designated 1D-17R. Lanes
1-7 represent reactions performed for 1, 3, 6, 10, 12, 22, and 30 min, respectively. Both primary and secondary products
were extended by another nucleotide, as dCTP was added after 30 min to
the reaction mixture. Reactions were allowed to proceed for another 10 and 30 min (lanes 6' and 7', respectively). The
runoff product and the paused product at position +12 were also both
observed after the addition of the fourth dNTP (dTTP) after 30 min
(lanes 6" and 7"). C lanes indicate a
control in the absence of dNTPs. B, shown is a schematic
representation of the results shown in A.
|
|
It is difficult to assess the extent to which such secondary initiation
reactions may contribute to rescue of DNA synthesis, as RNase H
cleavage continues into the RNA fragment (Fig. 5A). These
cleavage events give rise to 3'-truncated PPT fragments that initiate
DNA synthesis with diminished efficiency (35). A second round of DNA
synthesis may be relevant when DNA synthesis from longer primary
products is aggravated by a mispaired primer terminus, as shown for
position +6. It remains to be seen whether such mechanisms might
contribute to maintenance of the highly conserved sequence downstream
from the PPT that is recognized by the viral integrase.
 |
DISCUSSION |
Structural differences among double-stranded DNA, double-stranded
RNA, and DNA/RNA primer/template substrates can modulate reverse
transcription in multiple ways, which include both overall efficiency
of polymerization as well as the accuracy of this process. In this
report, we investigated the efficiency and fidelity of the initiation
of HIV-1 (+)-strand DNA synthesis, a stage of reverse transcription at
which RT accommodates an RNA/DNA primer/template. Our data show that
the efficiency of initial incorporation events was severely compromised
compared with subsequent events when the active site interacts with the
newly synthesized DNA duplex. This finding is reminiscent of recent
biochemical studies showing slow rates of RNA-primed initiation of
(
)-strand DNA synthesis and a sharp transition to a faster and more
processive mode of polymerization once the sixth nucleotide has been
incorporated (36, 37). Both RNA-primed reactions may therefore be
considered as important rate-limiting steps during reverse
transcription that respond to low concentrations of available dNTP
pools in a particular sensitive fashion. Despite this analogy, our
study also highlights significant functional differences between the two initiation events and reactions that involve double-stranded DNA or
DNA/RNA primer/templates.
Insertions of Correct Nucleotides--
Several lines of evidence
suggest a functional distinction between the first two nucleotide
additions and the ensuing polymerization events. Differences
regarding the relative efficiencies of DNA- versus
RNA-primed reactions, i.e.
(kcat/Km)DNA/(kcat/Km)RNA, were significantly less pronounced beyond position +2 (Table I). The low efficiency of nucleotide incorporation at position +2 with the
RNA primer is consistent with our recent observation of an RNA-specific
pause site after a single nucleotide extension (23). The relatively
sharp transition from initially low catalytic efficiencies to a more
efficient mode of DNA synthesis at position +3 is seen with limited
concentrations of enzyme as well as with an excess of RT over
primer/template.
These data suggest a modulation of interactions between RT and its
nucleic acid substrate that appears to specifically involve the first
two steps of DNA synthesis. Our recent measurements of the numbers of
base pairs that fit between the polymerase and RNase H active sites
have indeed identified structural differences between complexes
containing RNA/DNA and DNA/DNA (23). These variations most likely
reflect differences in helical parameters of the bound nucleic acid
substrates. However, it seems unlikely that such global structural
alterations can explain the observed high Km values
for initial nucleotide incorporations. In fact, despite a retained
difference in the number of base pairs (23), the efficiency of
nucleotide addition at position +6 was literally identical with both
RNA and DNA primers (Table I).
The sharp functional transition between positions +2 and +3 rather
suggests that structural differences between both substrates may be
critical, as they affect interactions in the vicinity of the active
site. This interpretation is consistent with mutational analysis of the
RT "primer grip," which is implicated in interactions with the
first one to approximately three nucleotides of the primer strand (38,
39). Some amino acid substitutions were shown to selectively impair
RNA-primed DNA synthesis, whereas the same mutant enzymes were capable
of efficiently recruiting the homologous DNA primer. These results,
along with the kinetic data presented in this work, strongly suggest
that the primer grip of RT plays a dominant role in substrate
discrimination between RNA/DNA and DNA/DNA.
The so-called "minor groove binding track" (MGBT), which is located
in the thumb subdomain of HIV-1 RT (40), is presumably less important
in this regard, although particular mutations in this motif were shown
to alter efficiency and specificity of the primer removal reaction
(41). The crystal structure of HIV-1 RT cross-linked to a DNA duplex
via residues of the MGBT shows contacts with the minor groove at a
distance of ~5-7 base pairs upstream from the polymerase active site
(42). Thus, the MGBT still interacts with the RNA/DNA segment as DNA
synthesis proceeds relatively efficiently beyond position +2. In
contrast, the location of this motif correlates well with the increased
rates of polymerization seen after incorporation of the sixth
nucleotide during RNA-primed (
)-strand DNA synthesis, suggesting that
the MGBT might play a role in discriminating between RNA/RNA and
DNA/RNA substrates (36, 37).
Specific structural features of the distinct chimeric replication
intermediates that are generated during (
)- and (+)-strand DNA
synthesis may provoke altered interactions with RT and may, in turn,
influence formation of a catalytically competent ternary complex. The
NMR structure of a model substrate that mimics the chimeric duplex
formed during initiation of (
)-strand DNA synthesis suggests an
unusual narrow minor groove around the tRNA-DNA junction (43). The
narrow minor groove and also the rigidity of the RNA/RNA segment are
likely to be considered as structural parameters that alter optimal
interactions with the MGBT (14, 43). In analogy, a compression of the
minor groove correlates with termination of (+)-strand DNA synthesis
after the second strand transfer (44). The unusual narrow minor groove
of the chimeric replication intermediate generated during tRNA-primed
(
)-strand DNA synthesis may also help to explain frequent pausing of
RT, seen at positions +3 and +5 (45). In contrast, the NMR structure of
the PPT-derived substrate that is accommodated during (+)-strand
initiation points to relatively wide minor groove dimensions (46); and
in fact, pausing during initiation of HIV-1 (+)-strand DNA synthesis is
restricted to position +1 (23). This isolated initial pausing site
suggests again a dominant role of the RT primer grip in discriminating between RNA/DNA and DNA/DNA substrates. However, specific pausing at
position +12 shows that the structure of the PPT-derived hybrid regains
importance at a later stage of DNA synthesis, presumably through
altered interactions with the RNase H domain.
Fidelity--
The efficiencies with which incorrect nucleotides
are incorporated during the initiation of (+)-strand DNA synthesis
follow a pattern similar to that seen in regard to correct nucleotides. The result is more extreme, as misinsertions were never seen at positions +1 and +2. Fidelity of DNA synthesis at position +6 was
similar to that seen with the homologous DNA primer, but the extension
of misaligned primers was still diminished compared with DNA-primed
reactions. When the two initial steps were bypassed, DNA synthesis
could still be detected, even in the absence of one of the three
nucleotides. Considering the high error rates reported for other types
of substrates, the initiation of (+)-strand DNA synthesis, particularly
the first two nucleotide incorporation events, is unusually accurate.
These data, together with previous biochemical studies (30-34),
suggest that fidelity increased, in order of dependence on the
primer/template substrate, as follows: DNA/DNA < DNA/RNA < RNA/RNA
RNA/DNA. The availability of data regarding the
initiation complex that contains the tRNA/RNA duplex is relatively
limited. However, experiments with only two or three of the four dNTPs
revealed that misincorporations were rarely observed, similar to early
steps during initiation of (+)-strand DNA synthesis (47, 48). The high
fidelity of both RNA-primed reactions may be advantageous for viral
replication, considering that the initiation of (
)- and (+)-strand
DNA synthesis and the corresponding primer removal reactions define the
ends of the pre-integrative DNA that serves as substrate of the viral
integrase (11).
The fidelity of the initiation of (+)-strand DNA synthesis is further
increased, as reactions were conducted with the wild-type enzyme, which
contains an intact RNase H domain. Nucleotide concentrations of ~100
nM, which allowed 50% full-length DNA synthesis with the RNase H-deficient RT-E478Q mutant, were not sufficient to bypass critical template positions when using the wild-type enzyme. Instead, RNase H cleavage at the newly formed RNA-DNA junction was seen, as
these positions become involved in DNA synthesis. These data point to a
fragile equilibrium between the polymerase- and RNase H-competent modes
that is shifted toward RNase H degradation as the polymerase process is
diminished. As for the natural pausing site at position +12, diminished
or biased dNTP levels decrease the efficiency of DNA synthesis by an
extent sufficient for initiation of RNase H activity. These data
strongly support the existence of two competing binding modes of the RT
enzyme on its PPT-derived substrate (23, 49), i.e.
polymerase-competent binding mode, which involves specific interactions
between the polymerase active site and the 3'-end of the primer
terminus, and an RNase H-competent binding mode, in which the RNase H
active site is positioned over the RNA-DNA junction in the same
orientation as seen during synthesis of (
)-strand DNA. Thus, the
concentration of available dNTP pools may also be considered as an
important parameter that influences specificity for PPT primer usage.
It is conceivable that intracellular dNTP concentrations are not high
enough to favor a polymerase-competent binding mode with an RNA/DNA
substrate that involves non-PPT primers (26, 49).
Taken together, our data reveal several unique features of the
initiation of HIV-1 (+)-strand DNA synthesis that distinguish this
stage of reverse transcription from other RNA- and DNA-primed reactions. The results point to an important rate-limiting step in
reverse transcription since relatively high dNTP concentrations were
required to overcome three distinct aspects of (+)-strand initiation.
These are as follows: 1) the high fidelity, yet inefficient incorporation of the first two nucleotides into the (+)-strand; 2) the
fragile equilibrium between the polymerase- and RNase H-competent binding modes; and 3) the specific pausing site at position +12 that
aggravates DNA synthesis even after primer removal (Fig. 4). Further
studies should now analyze the efficiency of (+)-strand DNA synthesis
using mutant enzymes that display diminished dNTP-binding capacities,
e.g. as described for amino acid substitutions at Tyr115 (50, 51). In this context, it is important to note
that the majority of amino acid substitutions in nucleoside
analog-resistant RT molecules affect either directly or indirectly the
nucleotide-binding pocket (42, 52). It is thus conceivable that this
type of amino acid substitution may cause significant reductions in the efficiency of RNA-primed reactions that require per se
relatively high concentrations of dNTPs. This may help to explain the
attenuated replication of some drug-resistant viruses that can also be
correlated, in some cases, with the diminished processivity displayed
by the corresponding mutant enzymes (53-56).
 |
ACKNOWLEDGEMENT |
We thank Dr. S. F. J. Le Grice
for Escherichia coli strains expressing wild-type
HIV-1 RT as well as the RNase H-deficient mutant enzyme.
 |
FOOTNOTES |
*
This work was supported by grants from the Medical Research
Council of Canada.The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
¶
To whom correspondence should be addressed: McGill AIDS
Centre, Lady Davis Institute-Jewish General Hospital, 3755, chemin Côte-Ste-Catherine, Montréal, Québec H3T 1E2, Canada.
Tel.: 514-340-8222 (ext. 3299); Fax: 514-340-7537; E-mail:
mgoette@ldi.jgh.mcgill.ca.
Published, JBC Papers in Press, November 28, 2000, DOI 10.1074/jbc.M009097200
 |
ABBREVIATIONS |
The abbreviations used are:
dNTPs, deoxynucleoside triphosphates;
HIV-1, human immunodeficiency virus type
1;
RT, reverse transcriptase;
PPT, polypurine tract;
MGBT, minor groove
binding track.
 |
REFERENCES |
1.
|
Meyerhans, A.,
Vartanian, J.-P.,
Hultgren, C.,
Plikat, U.,
Karlsson, A.,
Wang, L.,
Eriksson, S.,
and Wain-Hobson, S.
(1994)
J. Virol.
68,
535-540[Abstract]
|
2.
|
O'Brian, W. A.,
Namazi, A.,
Kalhor, H.,
Mao, S.-H.,
Zack, J. A.,
and Chen, I. S. Y.
(1994)
J. Virol.
68,
1258-1263[Abstract]
|
3.
|
Zack, J. A.,
Haislip, A. M.,
Krogstad, P.,
and Chen, I. S. Y.
(1992)
J. Virol.
66,
1717-1725[Abstract]
|
4.
|
Korin, Y. D.,
and Zack, J. A.
(1998)
J. Virol.
72,
3161-3168[Abstract/Free Full Text]
|
5.
|
Kootstra, N. A.,
Zwart, B. M.,
and Schuitemaker, H.
(2000)
J. Virol.
74,
1712-1717[Abstract/Free Full Text]
|
6.
|
Zhang, H.,
Dornadula, G.,
and Pomerantz, R. J.
(1996)
J. Virol.
70,
2809-2824[Abstract]
|
7.
|
Vartanian, J.-P.,
Plikat, U.,
Henry, M.,
Mahieux, R.,
Guillemot, L.,
Meyerhans, A.,
and Wain-Hobson, S.
(1997)
J. Mol. Biol.
270,
139-151[CrossRef][Medline]
[Order article via Infotrieve]
|
8.
|
Bebenek, K.,
and Kunkel, T. A.
(1993)
in
Reverse Transcriptase
(Skalka, A. M.
, and Goff, S. P., eds)
, pp. 85-102, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
|
9.
|
Schinazi, R. F.,
Larder, B. A.,
and Mellors, J. W.
(2000)
Int. Antiviral News
8,
65-91
|
10.
|
Gilboa, E.,
Mitra, S. W.,
Goff, S. P.,
and Baltimore, D.
(1979)
Cell
18,
93-100[Medline]
[Order article via Infotrieve]
|
11.
|
Telesnitsky, A.,
and Goff, S. P.
(1997)
in
Retroviruses
(Coffin, J. M.
, Hughes, S. H.
, and Varmus, H. E., eds)
, pp. 121-160, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
|
12.
|
Marquet, R.,
Isel, C.,
Ehresmann, C.,
and Ehresmann, B.
(1995)
Biochimie (Paris)
77,
113-124[CrossRef][Medline]
[Order article via Infotrieve]
|
13.
|
Mak, J.,
and Kleiman, L.
(1997)
J. Virol.
71,
8087-8095[Free Full Text]
|
14.
|
Götte, M.,
Li, X.,
and Wainberg, M. A.
(1999)
Arch. Biochem. Biophys.
365,
199-210[CrossRef][Medline]
[Order article via Infotrieve]
|
15.
|
Schatz, O.,
Mous, J.,
and Le Grice, S. F.
(1990)
EMBO J.
9,
1171-1176[Abstract]
|
16.
|
Furfine, E. S.,
and Reardon, J. E.
(1991)
J. Biol. Chem.
266,
406-412[Abstract/Free Full Text]
|
17.
|
Gopalakrishnan, V.,
Peliska, J. A.,
and Benkovic, S. J.
(1992)
Proc. Natl. Acad. Sci. U. S. A
89,
10763-10767[Abstract]
|
18.
|
Götte, M.,
Fackler, S.,
Hermann, T.,
Perola, E.,
Cellai, L.,
Gross, H.-J.,
Le Grice, S. F. J.,
and Heumann, H.
(1995)
EMBO J.
14,
833-841[Abstract]
|
19.
|
Götte, M.,
Maier, G.,
Gross, H.-J.,
and Heumann, H.
(1998)
J. Biol. Chem
273,
10139-10146[Abstract/Free Full Text]
|
20.
|
Peliska, J. A.,
and Benkovic, S. J.
(1992)
Science
258,
1112-1118[Medline]
[Order article via Infotrieve]
|
21.
|
Huber, H. E.,
and Richardson, C. C.
(1990)
J. Biol. Chem.
265,
10565-10573[Abstract/Free Full Text]
|
22.
|
Fuentes, G. M.,
Rodríguez-Rodríguez, L.,
Fay, P. J.,
and Bambara, R. A.
(1995)
J. Biol. Chem
270,
28169-28176[Abstract/Free Full Text]
|
23.
|
Götte, M.,
Maier, G.,
Mochi Onori, M.,
Cellai, L.,
Wainberg, M. A.,
and Heumann, H.
(1999)
J. Biol. Chem.
274,
11159-11169[Abstract/Free Full Text]
|
24.
|
Champoux, J. J.
(1993)
in
Reverse Transcriptase
(Skalka, A. M.
, and Goff, S. P., eds)
, pp. 103-118, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
|
25.
|
Charneau, P.,
Alizon, M.,
and Clavel, F.
(1994)
J. Mol. Biol.
241,
651-662[CrossRef][Medline]
[Order article via Infotrieve]
|
26.
|
Klarman, G. J., Yu, H.,
Chen, X.,
Dougherty, J. P.,
and Preston, B. D.
(1997)
J. Virol.
71,
9259-9269[Abstract]
|
27.
|
Le Grice, S. F.,
and Grüninger-Leitch, F.
(1990)
Eur. J. Biochem.
178,
307-314
|
28.
|
Boosalis, M. S.,
Petruska, J.,
and Goodman, M. F.
(1987)
J. Biol. Chem.
262,
14689-14696[Abstract/Free Full Text]
|
29.
|
Schatz, O.,
Cromme, F. V.,
Grüninger-Leitch, F.,
and Le Grice, S. F.
(1989)
FEBS Lett.
257,
311-314[CrossRef][Medline]
[Order article via Infotrieve]
|
30.
|
Preston, B. D.,
Poiesz, B. J.,
and Loeb, L. A.
(1988)
Science
242,
1168-1171[Medline]
[Order article via Infotrieve]
|
31.
|
Roberts, J. D.,
Bebenek, K.,
and Kunkel, T. A.
(1988)
Science
242,
1171-1173[Medline]
[Order article via Infotrieve]
|
32.
|
Boyer, J. C.,
Bebenek, K.,
and Kunkel, T. A.
(1992)
Proc. Natl. Acad. Sci. U. S. A
89,
6919-6923[Abstract]
|
33.
|
Kati, W.,
Johnson, K. A.,
Jerva, L. F.,
and Anderson, K. S.
(1992)
J. Biol. Chem
267,
25988-25997[Abstract/Free Full Text]
|
34.
|
Kerr, S. G.,
and Anderson, K. S.
(1997)
Biochemistry
36,
14056-14063[CrossRef][Medline]
[Order article via Infotrieve]
|
35.
|
Powell, M. D.,
and Levin, J. G.
(1996)
J. Virol.
70,
5288-5296[Abstract]
|
36.
|
Lanchy, J.-M.,
Keith, G.,
Le Grice, S. F.,
Ehresmann, B.,
Ehresmann, C.,
and Marquet, R.
(1998)
J. Biol. Chem.
273,
24425-24432[Abstract/Free Full Text]
|
37.
|
Thrall, S. H.,
Krebs, R.,
Wöhrl, B. M.,
Cellai, L.,
Goody, R. S.,
and Restle, T.
(1998)
Biochemistry
37,
13349-13358[CrossRef][Medline]
[Order article via Infotrieve]
|
38.
|
Gosh, M.,
Williams, J.,
Powell, M. D.,
Levin, J. G.,
and Le Grice, S. F.
(1997)
Biochemistry
36,
5758-5768[CrossRef][Medline]
[Order article via Infotrieve]
|
39.
|
Powell, M. D.,
Gosh, M.,
Jacques, P. S.,
Howard, K. J.,
Le Grice, S. F. J.,
and Levin, J. G.
(1997)
J. Biol. Chem.
272,
13262-13269[Abstract/Free Full Text]
|
40.
|
Bebenek, K.,
Beard, W. A.,
Dargen, T. A.,
Li, L.,
Prasad, R.,
Luton, B. A.,
Gorenstein, D. G.,
Wilson, S. H.,
and Kunkel, T. A.
(1997)
Nat. Struct. Biol.
4,
194-197[Medline]
[Order article via Infotrieve]
|
41.
|
Powell, M. D.,
Beard, W. A.,
Bebenek, K.,
Howard, K. J.,
Le Grice, S. F.,
Darden, T. A.,
Kunkel, T. A.,
Wilson, S. H.,
and Levin, J. G.
(1998)
J. Biol. Chem.
274,
19885-19893[Abstract/Free Full Text]
|
42.
|
Huang, H.,
Chopra, R.,
Verdine, G. L.,
and Harrison, S. C.
(1998)
Science
282,
1669-1675[Abstract/Free Full Text]
|
43.
|
Szyperski, T.,
Götte, M.,
Billeter, M.,
Perola, E.,
Cellai, L.,
Heumann, H.,
and Wüthrich, K.
(1999)
J. Biomol. NMR
13,
343-355[CrossRef][Medline]
[Order article via Infotrieve]
|
44.
|
Lavigne, M.,
and Buc, H.
(1999)
J. Mol. Biol.
285,
977-995[CrossRef][Medline]
[Order article via Infotrieve]
|
45.
|
Isel, C.,
Lanchy, J.-M.,
Le Grice, S. F. J.,
Ehresmann, C.,
Ehresmann, B.,
and Marquet, R.
(1996)
EMBO J.
15,
917-924[Abstract]
|
46.
|
Fedoroff, O. Y,
Ge, Y.,
and Reid, B. R.
(1997)
J. Mol. Biol.
269,
225-239[CrossRef][Medline]
[Order article via Infotrieve]
|
47.
|
Lanchy, J.-M.,
Ehresmann, C.,
Le Grice, S. F. J.,
Ehresmann, B.,
and Marquet, R.
(1996)
EMBO J.
15,
7178-7187[Abstract]
|
48.
|
Oude Essink, B. B.,
and Berkhout, B.
(1999)
J. Biomol. Sci.
6,
121-132
|
49.
|
Palaniappan, C.,
Kim, J. K.,
Wisniewski, M.,
Fay, P. J.,
and Bambara, R. A.
(1998)
J. Biol. Chem
273,
3808-3816[Abstract/Free Full Text]
|
50.
|
Martin-Hernadez, A. M.,
Domingo, E.,
and Menendez-Arias, L.
(1996)
EMBO J
15,
4434-4442[Abstract]
|
51.
|
Boyer, P. L.,
Sarafianos, S. G.,
Arnold, E.,
and Hughes, S. H.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
3056-3061[Abstract/Free Full Text]
|
52.
|
Sarafianos, S. G.,
Das, K.,
Ding, J.,
Boyer, P. L.,
Hughes, S. H,
and Arnold, E.
(1999)
Chem. Biol.
6,
137-146[CrossRef]
|
53.
|
Boyer, P. L.,
and Hughes, S. H.
(1995)
Antimicrob. Agents Chemother.
39,
1624-1628[Abstract]
|
54.
|
Back, N. K. T.,
Nijhuis, M.,
Keulen, W.,
Boucher, C. A. B.,
Oude Essink, B. B.,
van Kuilenburg, A. B. P.,
van Gennip, A. H.,
and Berkhout, B.
(1996)
EMBO J
15,
4040-4049[Abstract]
|
55.
|
Quan, Y.,
Inouye, P.,
Liang, C.,
Rong, L.,
Götte, M.,
and Wainberg, M. A.
(1998)
J. Biol. Chem
273,
21918-21925[Abstract/Free Full Text]
|
56.
|
Sharma, P. L.,
and Crumpacker, C. S.
(1999)
J. Virol.
73,
8448-8456[Abstract/Free Full Text]
|
Copyright © 2001 by The American Society for Biochemistry and Molecular Biology, Inc.