From the Department of Biochemistry, University of Iowa, Iowa City, Iowa 52242
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ABSTRACT |
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Drosophila factor 2, an RNA polymerase II transcript release factor, exhibits a DNA-dependent ATPase activity (Xie, Z., and Price D. H. (1997) J. Biol. Chem. 272, 31902-31907). We examined the nucleic acid requirement and found that only double-stranded DNA (dsDNA) effectively activated the ATPase. Single-stranded DNA (ssDNA) not only failed to activate the ATPase, but suppressed the dsDNA-dependent ATPase. Gel mobility shift assays showed that factor 2 formed stable complexes with dsDNA or ssDNA in the absence of ATP. However, in the presence of ATP, the interaction of factor 2 with dsDNA was destabilized, while the ssDNA-factor 2 complexes were not affected. The interaction of factor 2 with dsDNA was sensitive to increasing salt concentrations and was competed by ssDNA. In both cases, loss of binding of factor 2 to dsDNA was mirrored by a decrease in ATPase and transcript release activity, suggesting that the interaction of factor 2 with dsDNA is important in coupling the ATPase with the transcript release activity. Although the properties of factor 2 suggested that it might have helicase activity, we were unable to detect any DNA unwinding activity associated with factor 2.
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INTRODUCTION |
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A common mechanism employed to control the potential of RNA
polymerase II to synthesize full-length transcripts is through an early
elongation block. This process, referred to as abortive elongation or
premature termination has been observed in various transcription
systems (1-5), as well as in our study of Drosophila RNA
polymerase II transcription (6, 7). We observed that two distinct
classes of complexes were formed after initiation. The predominant
class undergoes abortive elongation which only gives rise to short
transcripts, whereas the second class overcomes early blocks and
carries out productive elongation. Based on our results, a model has
been proposed for the control of elongation by RNA polymerase II which
highlights the function of both negative and positive factors (6, 7).
According to the model, the action of negative transcription elongation
factors (N-TEF)1 was
responsible for the abortive elongation. The transition from abortive
elongation to productive elongation was mediated by the action of
positive transcription elongation factors (P-TEF). P-TEFb, one of the
components of P-TEF, has recently been purified and identified as a
5,6-dichloro-1--D-ribofuranosylbenzimidazole-sensitive kinase that can phosphorylate the C-terminal domain (CTD) of the largest subunit of RNA polymerase II (8, 9) and is required for
Tat-transactivation of transcription from the HIV-LTR (10, 11).
One of the components of N-TEF, factor 2, was originally identified due to its ability to suppress the appearance of shorter than full-length transcripts during transcription in vitro (12). Factor 2 is a 154-kDa protein that associates stably with early elongation complexes under low salt conditions, but dissociates in 1 M KCl (13). Factor 2 causes the release of transcripts by RNA polymerase II in an ATP-dependent manner and results in premature termination (13). Further characterization of the transcript release activity of factor 2 revealed that the process requires ATP hydrolysis. Non-hydrolyzable analogs of ATP did not support factor 2 mediated transcript release (14). The intrinsic ATPase activity of factor 2 was activated by dsDNA and did not require other protein cofactors (14).
All information gathered on factor 2 function suggests that ATP hydrolysis is a key reaction for the transcript release activity of factor 2, but how it associates with elongation complexes and how the ATP hydrolysis is coupled to the transcript release event is not clear. Escherichia coli termination factor rho has an RNA-dependent ATPase activity that allows the translocation of rho along nascent transcripts to track elongation complexes and unwind RNA-DNA hybrids in transcription bubbles (15-17). RNA polymerase III termination factor La also has an ATPase activity that is dependent on RNA binding (18). Like rho, La causes the release of nascent transcripts by unwinding DNA-RNA hybrid using energy from ATP hydrolysis (18, 19). On the other hand, E. coli transcription repair coupling factor (TRCF) that is able to release nascent transcripts from stalled RNA polymerase lacks a helicase activity (20, 21). Instead, TRCF interacts with DNA and may associate with elongation complexes through its interaction with DNA and RNA polymerase and cause the release of RNA polymerase and nascent transcripts by hydrolyzing ATP (20, 21). Though it is not clear how factor 2 associates with elongation complexes, the finding that dsDNA is able to activate the ATPase activity of factor 2 (14) and the observation that factor 2 is able to bind DNA affinity column2 suggest that factor 2 may bind DNA and associate with elongation complexes through the interaction with DNA template. To understand the molecular mechanism of factor 2 function, in this study we further characterized the effects of various nucleic acids on the ATPase activity of factor 2. In addition, we examined the interaction of factor 2 with DNA and its significance for the ATPase and transcript release activity of factor 2. Furthermore, we explored the potential helicase activity of factor 2.
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EXPERIMENTAL PROCEDURES |
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ATPase Assay--
The ATPase activity of Drosophila
factor 2 was assayed by measuring the release of inorganic phosphate
from [32P]ATP as described (14). Immobilized dsDNA was
used in the ATPase assay to examine the effect of ionic strength on the
interaction of factor 2 with dsDNA. The immobilized dsDNA was generated
by coupling biotinylated DNA with streptavidin-conjugated paramagnetic Dynabeads as described earlier (7). Factor 2 was first incubated with
immobilized dsDNA in the presence of 20 mM HEPES (pH 7.6), 5 mM MgCl2, 0.2 mg/ml bovine serum albumin, and
55 mM KCl for 5 min at 25 °C. The mixture was then
aliquoted, concentrated magnetically, and washed three times with HMBK
buffer that contained 20 mM HEPES, 5 mM
MgCl2, 0.2 mg/ml of BSA, and KCl of indicated
concentration. The resulting complexes were washed another two times
with 55 mM HMBK buffer, resuspended in 55 mM
HMBK buffer, aliquoted to individual reaction tubes (6 µl/each tube),
and then examined for their ability to hydrolyze ATP. The ATPase assay
was initiated by the addition of 2 µl of 4 × label mixture (14)
and incubated at 25 °C for 20 min. The ATPase reactions were
terminated by the addition of 1 µl of 0.5 M EDTA. The
supernatant fractions containing the released
32Pi and the unhydrolyzed
[
32P]ATP were isolated by magnetic concentration and
analyzed by thin layer chromatography.
Gel Mobility Shift Assay--
The dsDNA substrate for gel
mobility shift assay was a 238-bp polymerase chain reaction product
amplified from PET-21a plasmid (Novagen, Inc.). The single-stranded DNA
(ssDNA) substrate was a synthetic oligo (dT)60 (kindly
provided by Dr. M. Wold, University of Iowa). Both the 238-bp duplex
DNA and oligo (dT)60 were 5 end-labeled and
electrophoretically purified. The binding of factor 2 with the labeled
DNA was carried out in 20 mM HEPES (pH 7.6), 5 mM MgCl2, 0.2 mg/ml bovine serum albumin, 1 mM dithiothreitol, and 50 mM KCl in the
presence or absence of 0.6 mM ATP at 25 °C for 20 min.
The reaction products were analyzed by electrophoresis on a 4.5%
polyacrylamide gel in 0.5 × TAE buffer (40 mM
Tris-acetate, pH 8.5, 2 mM EDTA) at 100-200 V.
Helicase Assay--
Two kinds of helicase substrates were
generated to examine the helicase activity of factor 2. One was a
partial duplex DNA as described (22). 5.2 ng of 17-mer universal primer
was annealed to 2 µg of ssM13 mp18 DNA by incubation in 40 mM Tris-HCl (pH 7.5), 10 mM MgCl2,
1 mM dithiothreitol and 50 mM NaCl2
at 95 °C for 5 min followed by 20 min at 65 °C and 20 min at
23 °C. The primer was extended by DNA polymerase I-Klenow fragment
(10 units) in the presence of 10 µCi of [32P]dCTP,
50 µM dATP and dGTP, 40 mM Tris-HCl (pH 7.5),
10 mM MgCl2, 1 mM dithiothreitol,
and 50 mM NaCl2 at 23 °C for 20 min, then the reaction mixture was supplemented with dCTP with a final
concentration of 50 µM and incubated for another 20 min.
The reaction was stopped by EDTA and the free nucleotides were removed
by gel filtration on a NICKTM column (Pharmacia Biotech
Inc.). The second type of substrate was a 40-bp duplex DNA made by
annealing two complementary oligonucleotides, one of which was labeled
by [
32P]ATP at the 5
end.
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RESULTS |
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The ATPase Activity of Factor 2 Requires dsDNA and Is Inhibited by
ssDNA--
Factor 2 exhibited an ATPase activity that was dramatically
stimulated by DNA (14). To gain further understanding of the nucleic
acid requirement, a variety of DNA and RNA molecules were examined for
their effect on the ATPase activity of factor 2. A standard ATPase
assay that measures the release of free phosphate from
[32P]ATP was performed in the presence of a constant
amount of different nucleic acids. The percent hydrolysis for each
reaction was normalized to the low level of activity exhibited by
factor 2 in the absence of any nucleic acid and is presented in Table
I. Up to 100-fold stimulation was seen
for dsDNA molecules of natural sequences. Supercoiled DNAs stimulated
slightly more than linear molecules. When the ability of supercoiled
and linear forms of either Bluescript or A2 DNA to stimulate the ATPase
activity were compared, we found that supercoiled DNA increased the
ATPase from 20 to 30% compared with the linear form (data not shown).
Synthetic homopolymeric dsDNA, such as poly(dA)·(dT),
poly(dG)·(dC), and poly(dA-dG)·(dT-dC), also stimulated the ATPase
activity of factor 2, although with somewhat lower efficiency than
dsDNA with natural sequences. ssDNA or dsRNA did not activate the
ATPase. Partial duplex DNA poly(dC), poly(dG)12-18, or a
DNA-RNA hybrid poly(I)·poly(rC) both failed to activate the ATPase.
Thus, the ATPase activity of factor 2 was efficiently stimulated only
by dsDNA.
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Factor 2 Forms a Stable Complex with dsDNA That Is Unstable in the
Presence of ATP or ssDNA--
Since dsDNA was required for the ATPase
activity of factor 2, we examined the interaction between factor 2 and
dsDNA with a gel mobility shift assay (Fig.
2). A 5-end labeled 238-base pair dsDNA
was used as the DNA substrate. Factor 2 formed stable complexes with
the dsDNA in the absence of ATP (Fig. 2). Two complexes were observed,
a major complex and a lower mobility minor complex. Results with DNAs
of different length suggest that the two complexes may be different
because of the number of factor 2 molecules bound (data not shown). The
binding of factor 2 to the labeled DNA was competed by a 10-fold excess
of unlabeled DNA (Fig. 2, last three lanes). Binding
reactions were also carried out with the addition of a nucleoside
triphosphate. The interaction of factor 2 with dsDNA was destabilized
in the presence of ATP, but was not affected by CTP, GTP, or ATP
S.
These results argued that the ATP hydrolysis by factor 2 destabilized
the dsDNA-factor 2 complexes.
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DNA Binding Properties of Factor 2 Correlate with Transcript Release Activity-- Since the preceding results showed that ssDNA was able to compete for dsDNA binding by factor 2 and suppress the dsDNA-dependent ATPase activity of factor 2, we tested the effect of ssDNA on the transcript release activity of factor 2. If transcript release mediated by factor 2 is coupled to the dsDNA-activated ATPase, ssDNA should effectively inhibit the transcript release activity of factor 2. High salt-washed early elongation complexes were generated on an immobilized DNA template, and the transcript release reaction was carried out as diagrammed above Fig. 4. Released transcripts were separated from transcripts in early elongation complexes by magnetic concentration of the template. The high salt-washed early elongation complexes lack factor 2 (13), thus almost all of the transcripts remained associated with early elongation complexes when only ATP was added to the reaction. As was found before, transcripts were released in the presence of both factor 2 and ATP (Fig. 4A, lane 2). However, when factor 2 was first preincubated with 15 µg/ml of ssDNA for 5 min and then incubated with the early elongation complexes in the presence of ATP, more than 70% of the transcripts remained bound to the early elongation complexes (Fig. 4A). Almost no transcript release by factor 2 was observed when factor 2 was preincubated with 45 µg/ml of ssDNA (Fig. 4A). These results demonstrate that ssDNA was able to suppress the transcript release activity and suggest that the ATPase activity of factor 2 is essential for the transcript release activity of factor 2.
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Factor 2 Has No Apparent DNA Helicase Activity-- Since factor 2 was able to bind dsDNA and hydrolyze ATP, we investigated whether the factor was able to utilize the energy from ATP hydrolysis to unwind duplex DNA. We first used a conventional helicase assay that utilized a labeled 24-nucleotide oligonucleotide annealed to M13 DNA. In the presence of ATP, the 24-mer was readily released by a bona fide DNA helicase, SV40 large T antigen, but not by factor 2 (Fig. 6A). To eliminate the possibility that the potential helicase activity was inhibited by ssDNA, we also examined the ability of factor 2 to first bind and then unwind a 40-bp DNA duplex. A gel mobility shift assay indicated that factor 2 was able to bind the 40-bp duplex (Fig. 6B). As was found previously, the binding was destabilized by ATP, suggesting that the short DNA was able to activate the ATPase activity of factor 2 (Fig. 6B). Indeed, the ATPase assay demonstrated this directly (data not shown). Even though factor 2 bound to the DNA fragment and exhibited ATPase activity, it failed to unwind the dsDNA either at 25 °C (Fig. 6C) or 37 °C (data not shown). These results suggest that factor 2 does not have a DNA helicase activity.
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DISCUSSION |
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Factor 2 was identified as a component of N-TEF that was able to block transcription elongation by promoting the release of nascent transcripts in an ATP-dependent manner (13). Further study revealed that factor 2 had a DNA-dependent ATPase activity (14). In this study, we characterized the ATPase activity with respect to nucleic acid requirement and explored the possibility of associated helicase activities. In addition, a correlation was made between the ATPase and the transcript release activity of the factor. The results indicate that factor 2 possesses a unique collection of properties.
Characterization of the effect of nucleic acids on the ATPase activity of factor 2 distinguished it from almost all other proteins. Only dsDNA effectively supported the ATPase activity, with supercoiled DNA having a slightly higher efficiency compared with linearized DNA. ssDNA not only failed to activate but actually suppressed the dsDNA-dependent ATPase activity of factor 2. The strict requirement of dsDNA for ATPase activity has only been observed for RuvB protein from eubacteria Thermus thermophilus and Thermotoga martima (25). Studies of T. thermophilus RuvB suggests that is involved in recombinational DNA repair and the homologous genetic recombination process (25). The involvement of factor 2 in these processes has not been determined. The inhibitory effect of ssDNA on the dsDNA-dependent ATPase activity of factor 2 has not been observed for other proteins; however, uncovering the significance of this property to factor 2 function will require further investigation.
The strict requirement of dsDNA for the ATPase activity of factor 2 suggested that double helical structure of DNA is an essential element in activating the ATPase. ssDNA was able to bind factor 2 but failed to activate the ATPase. dsRNA or DNA-RNA hybrid that adopts the A-form helical structure also failed to activate the ATPase. Since dsDNA is usually in the B-form helical structure, it is tempting to speculate that only the typical B-form helical structure of dsDNA is competent to activate the ATPase. Indeed, this may provide a possible explanation for why a double-stranded homopolymer had a lower efficiency in activating the ATPase activity of factor 2 compared with dsDNA with natural sequence (Table I). Poly(dA)·(dT) and poly(dG)·(dC) has been shown to adopt a helical conformation that is quite different from the typical B-form DNA (26, 27). Ethidium bromide, which intercalates within dsDNA and untwists the DNA helix, was also found to suppress the dsDNA-dependent ATPase activity of factor 2 (Fig. 1A). All of these findings support the hypothesis that B-form helical structure of DNA is important for activating the ATPase activity of factor 2. DNA structure, especially that altered by protein factors, may play an important role in controlling the action of factor 2.
The binding of factor 2 to dsDNA seems to be a primary requirement for the transcript release activity of factor 2, and we propose that factor 2 initially associates with the elongation complex through interaction with the template. When the interaction between factor 2 and dsDNA was blocked by ssDNA, the transcript release was also inhibited (Fig. 4). ssRNA, however, did not stimulate or suppress the ATPase activity (Fig. 1B), suggesting that factor 2 does not bind to RNA and may not associate with the elongation complex through nascent RNA. This assumption was also reached from our previous finding that factor 2 was able to release very short transcripts which are sequestered within the ternary complexes and not accessible to factor 2. Recent characterization of factor 2 function using a dC-tailed template suggested that the dsDNA upstream of the elongation complexes was required for the transcript release activity of factor 2 (14). It is possible that not only the binding of factor 2 with dsDNA but also its appropriate orientation to elongation complexes is necessary to effectively couple the dsDNA-dependent ATPase activity of factor 2 with its transcript release activity.
The effect of ssDNA on the properties of factor 2 can be used in understanding some aspects of factor 2 function. Direct binding of factor 2 to dsDNA in the absence of ATP as measured with a gel mobility shift assay was efficiently competed only with a 100-fold excess of ssDNA. However the ATPase activity of factor 2 was about 100 times more sensitive to ssDNA. Since ATP weakens the association of factor 2 with dsDNA but not with ssDNA, it is likely that ssDNA traps factor 2 when both dsDNA and ATP are present. The effect of ssDNA on transcript release also supported this notion. ssDNA had a greater inhibitory effect on transcript release when it was preincubated with factor 2, presumably because the factor 2 complexed with ssDNA was unable to bind to the template. Although ssDNA was able to inhibit transcript release when the factor 2 was prebound to the template, it was required in about a 10-fold excess over the dsDNA. This is a significantly higher single strand/double strand ratio than was required in the ATPase assay even though both assays contained ATP. There are several explanations for the relative resistance to inhibition by ssDNA of transcript release compared with the ATPase activity. One possibility for the lower efficiency of ssDNA inhibition in transcript release is that factor 2 may interact not only with dsDNA but also with other protein factors in the elongation complexes. Therefore, even in the presence of ATP, more ssDNA may be required to dissociate factor 2 from elongation complexes and suppress its transcript release activity. Another interesting possibility is that factor 2 may exhibit an ATP-dependent translocation along dsDNA. In this way, ATP could cause factor 2 to release dsDNA by causing it to run off the end of the fragment. Translocation along the template would be impeded by proteins including RNA polymerase II ternary complexes. Perhaps the mechanism of termination involves the translocation of factor 2 into the trailing edge of the polymerase. If the translocation model is correct, ssDNA would be less effective at inhibiting transcript release because factor 2 is not released from the template until after it has caused transcript release.
An alternative model for the effect of ATP is that it may cause direct release of factor 2 from the template. Since factor 2 associated tightly with dsDNA in the absence of ATP, destabilization of the dsDNA-factor 2 complexes by ATP may be necessary to reduce the nonspecific retention of factor 2 on dsDNA and facilitate its search for stalled RNA polymerase. However, this model does not explain the relative resistance of transcript release to ssDNA. In addition, factor 2 would be quite inefficient in localizing non-processive elongating polymerase and mediating transcript release. Perhaps more likely is an intermediate model in which factor 2 scans the template for a period of time and then dissociates from the template. Reassociation and resuming of scanning would be the normal pathway for transcript release and this process would be inhibited by ssDNA. The intermediate inhibition by ssDNA of transcript release could mirror the balance between processive scanning and transient interaction. More detailed kinetic experiments will be required to verify or differentiate between the models.
The effect of elevated salt on the transcript release activity of factor 2 correlated with the effect on the ATPase activity but did not correlate with release of factor 2 from dsDNA. At 200 mM KCl, factor 2 remained associated with the template but was not active in transcript release, presumably because the ATPase activity was inhibited. Factor 2 may be responsible for our previous unpublished results which indicated that early elongation complexes would lengthen transcripts in the presence of 250 mM KCl but would stop if the salt was lowered. Evidently, at intermediate salt concentrations, factor 2 remains bound to the template but does not function. This easily accessible state, in which factor 2 is bound but not active, suggests that a mechanism for regulation of factor 2 transcript release activity may exist. Perhaps other protein factors can interact with factor 2 and cause it to adopt the inactive bound state. Such interactions may be important in regulating the selectivity of factor 2 so that the action of factor 2 is restricted to non-productive elongation complexes.
Biochemical characterization of factor 2 shows that it shares multiple properties with the TRCF. Like E. coli TRCF protein Mfd, (20, 21) and two potential eucaryotic TRCF, human ERCC6 (28) and its yeast homologue Rad26 (29), factor 2 has an associated ATPase and ATP-dependent transcript release activity, binds to dsDNA, and lacks an apparent helicase activity. However, ERCC 6 lacks one of the major features of E. coli TRCF, the ability to dissociate the stalled RNA polymerase II complexes. It has been argued that RNA polymerase II stalled by DNA lesion may not necessarily be dissociated by eucaryotic TRCF but may be removed from damaged sites through the action of SII or other unknown factors (28, 30). Though the multiple biochemical properties shared between factor 2 and E. coli TRCF suggests that factor 2 may be a homologue of eucaryotic TRCF, more in vitro and in vivo functional study of factor 2 will be required to justify its cellular function.
In summary, characterization of the enzymatic activity of factor 2 and its interaction with DNA provided further insights into the mechanism of factor 2 action. Compared with other known transcription termination factors, factor 2 may utilize a novel mechanism for transcript release. Unlike E. coli termination factor rho (15-17) or RNA polymerase III termination factor La (18, 19) or termination of vaccinia virus polymerase mediated by VTF (virus-encoded termination factor) (31, 32), no specific signal in nascent RNA or binding to the nascent RNA is required for the transcript release mediated by factor 2. Instead, our results indicate that factor 2 is likely recruited to the elongation complex through interactions with the template DNA. Although a strong correlation between the ATPase activity and transcript release activity of factor has been made, the mechanism of transcript release remains to be determined.
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ACKNOWLEDGEMENT |
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We thank Marc Wold (University of Iowa) for kindly providing SV40 large T antigen and the 40-bp dsDNA used for one of the helicase assays.
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FOOTNOTES |
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* This work was supported by National Institutes of Health Grant RO1-GM35500.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.
Present address: Howard Hughes Medical Institute, Vanderbilt
University, Nashville, TN, 37232.
§ To whom correspondence should be addressed. Tel.: 319-335-7910; Fax: 319-335-9570; E-mail: david-price{at}uiowa.edu.
1
The abbreviations used are: N-TEF, negative
transcription elongation factors; dsDNA, double-stranded DNA; ssDNA,
single-strand DNA; P-TEF, positive transcription elongation factors;
HIV, human immunodeficiency virus; TRCF, transcription repair coupling
factor; bp, base pairs(s); ATPS, adenosine
5
-O-(thiotriphosphate).
2 Xie, Z., and Price, D.H., unpublished data.
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REFERENCES |
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