Molecular Cell, 2005, in press
The RNA Tether from the Poly(A) Signal to the Polymerase Mediates
Coupling of Transcription to Cleavage and Polyadenylation
Frank Rigo, Amir Kazerouninia, Anita Nag and Harold G. Martinson
Department of Chemistry and Biochemistry and the Molecular Biology Institute
University of California at Los Angeles
Los Angeles, California 90095-1569
Summary
We have investigated the mechanism by which transcription accelerates cleavage and
polyadenylation in vitro. Using a coupled transcription-processing system we show that rapid
and efficient 3′-end processing occurs in the absence of crowding agents like polyvinyl alcohol.
The continuity of the RNA from the poly(A) signal down to the polymerase is critical to this
processing. If this tether is cut during transcription using DNA oligonucleotides and RNase H,
the efficiency of processing is drastically reduced. The polymerase is known to be an integral
part of the cleavage and polyadenylation apparatus. RNA polymerase II pull-down and
immobilized template experiments suggest that the role of the tether is to hold the poly(A) signal
close to the polymerase during the early stages of processing complex assembly until the
complex is sufficiently mature to remain stably associated with the polymerase on its own.
Running Title: A tether couples 3′-end processing to transcription
Introduction
The production of mRNA in the nuclei of eukaryotes is a complex multistep process that begins
with the initiation of transcription and culminates in the export of the mature message. All of
these steps, including all stages of transcription, processing and export, are functionally
interconnected through a web of mutually synergistic interactions (Maniatis and Reed, 2002).
Here we focus on the role of transcription in facilitating the final step of processing, cleavage
and polyadenylation.
Although the ability of the poly(A) signal to modulate transcription (by causing termination)
has been known now for almost two decades (Whitelaw and Proudfoot, 1986), the idea that
transcription, in turn, can affect 3′-end processing is more recent (Dantonel et al., 1997;
McCracken et al., 1997). A widely accepted proposal is that RNA polymerase II, through the
carboxyl-terminal repeat domain of its large subunit (CTD), gathers processing factors and
delivers them to the emerging transcript during transcription (Proudfoot, 2004; Bentley, 2005).
However, this postulated recruitment function is difficult to distinguish experimentally from the
known function of the CTD as a required participant in the poly(A) site cleavage reaction
(Hirose and Manley, 1998; Ryan et al., 2002). This CTD requirement for cleavage is manifested
in the absence of transcription and is thus distinct from any transcription-related recruitment
function. Interestingly, there is another connection between transcription and processing that has
received almost no experimental attention. This is the nascent RNA that links the processing
apparatus to the polymerase. This tether is known to be functionally important in prokaryotes
(Nodwell and Greenblatt, 1991) but its role in eukaryotes has not been examined.
A typical core poly(A) signal in mammals consists of two recognition elements (an
AAUAAA hexamer upstream and a U or GU-rich element downstream) flanking the poly(A)
cleavage site (Zhao et al., 1999). Although the only chemistry required for cleavage at the
poly(A) site is hydrolysis of a single phosphodiester bond in the RNA, the apparatus that must be
assembled to do this is enormously complex (Calvo and Manley, 2003; Proudfoot, 2004).
Presumably this reflects regulatory functions consistent with its connection to such far-flung
activities as transcription, capping, splicing and transport (Flaherty et al., 1997; Hammell et al.,
2002; Proudfoot et al., 2002; Calvo and Manley, 2003). The ultimate consequence is an
apparatus so large that if bound to the CTD it would dwarf the polymerase.
It is not known how the cleavage apparatus is assembled on the poly(A) signal, but various
data suggest that it is a stepwise process (Chao et al., 1999; Takagaki and Manley, 2000). This is
consistent with its complexity and with the lag that reportedly precedes cleavage in vitro
(Ruegsegger et al., 1998). It is also unclear what special contribution transcription might make
to assembly. For example, chromatin immunoprecipitation data from yeast suggest that the CTD
may play only a limited role in factor recruitment prior to the appearance of the poly(A) signal
(Kim et al., 2004). To better understand the role of transcription in facilitating 3′-end processing
we have initiated experiments to investigate this problem in vitro. We first sought conditions in
which transcription would yield RNA that is rapidly and efficiently processed under conditions
of ongoing transcription, and then we asked questions about the mechanism. Interestingly, our
results highlight, not the role of the CTD, but of the RNA itself in the assembly of the cleavage
complex. We favor a model in which the polymerase on its own cannot bind the assembling
cleavage complex with sufficient stability to support maturation, but relies on the RNA to retain
the nascent apparatus in close proximity until assembly on the polymerase is complete.
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Results
3′-end processing coupled to transcription. Fig. 1A shows 3′-end processing that is fast and
efficient when coupled to transcription in vitro in a nuclear extract. Following pre-initiation
complex formation, transcription was initiated with a pulse of [α-32P] CTP, and then chased for
various times with a high concentration of unlabeled CTP (Fig. 1A). The gel in Fig. 1A shows
transcript length steadily increasing so that by 4 min the majority of polymerases have passed the
poly(A) site. Then, after a short lag, cleaved and polyadenylated transcripts appear and rapidly
accumulate. The % polyadenylation is given below the gel and is plotted as a function of time in
Fig. 1E (closed squares). Note that the burst of polyadenylation is rapidly finished. Although
the intensity of the poly(A) RNA band increases significantly between 20 and 60 min (Fig. 1A,
lanes 10 and 11), most of this increase probably reflects the overall boost to intensity in the upper
part of lane 11 that can be attributed to laggard polymerases crossing the poly(A) site after 20
min. This is consistent with the corresponding decrease in intensity below the position of the
poly(A) site in lane 11 relative to lane 10. These results suggest that most polymerases remain
transcriptionally engaged for the duration of the experiment, or at least until processing occurs.
Note that little or no cleaved but non-polyadenylated RNA appears on the gel, consistent with the
known tight coupling of cleavage and polyadenylation (Manley et al., 1982; Moore and Sharp,
1985). The polyadenylated RNA band is broad because of heterogeneity in poly(A) tail length
(Wahle, 1995).
To confirm that the RNA was polyadenylated and accurately cleaved we oligo(dT) selected
the RNA and characterized it by RNase protection (Fig. 1B). The oligo(dT) selected RNA (lane
4) gave a protected fragment, identical to that from RNA transcribed in vivo in transfected cells
(lane 1), that was not there when the poly(A) signal was inactivated by mutation (lanes 3 and 5).
Fig. 1A used the early promoter and late poly(A) signal of SV40. The cytomegalovirus
(CMV) promoter and bovine growth hormone (BGH) poly(A) signal gave similar results (Fig.
1C and Fig. 1E, open squares). We confirmed that processing in Fig. 1C was correct by blocking
polyadenylation and measuring the size of the cleaved but non-polyadenylated RNA produced,
an approach more convenient than RNase protection. To block polyadenylation we added 3′dATP to the coupled reaction before processing began but after a large fraction of the
polymerases had crossed the poly(A) site (4 min). Fig. 1D shows that, indeed, the broad band of
correctly processed RNA in lane 1 gave way, after 3′-dATP treatment, to a sharper band of
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cleaved but non-polyadenylated RNA running faster in the gel (lane 2). This band was authentic
poly(A) site cleaved RNA—a mutant BGH poly(A) signal yielded no such band (lane 3),
whereas mutant transcripts cut at the poly(A) site using RNase H did (lane 4).
Fig. 1E compares transcription-polyadenylation as a function of time for three different
nuclear extract preparations and two different promoters and poly(A) signals. All display similar
3′-end processing kinetics and efficiencies.
We wondered whether the rapidity of 3′-end processing seen in Fig. 1 requires on-going
transcription. To answer this we took advantage of the lag before processing begins (see Fig.
1A, lane 3). We pulsed, chased for 3.5 min and then, before processing began, added α-amanitin
to stop transcription. We also added 3′-dATP, as above, to block poly(A) tail growth and
highlight the poly(A) site cleavage event per se (Niwa et al., 1990; Cooke et al., 1999). Fig. 2A,
lane 1 confirms that shortly after α-amanitin and 3′-dATP were added the majority of transcripts
remained uncleaved. However, efficient poly(A) site cleavage rapidly ensued (Fig. 2A, lanes 24) and displayed similar kinetics to a parallel reaction in which both transcription and
polyadenylation were allowed to proceed (lanes 5 and 6). The data are plotted as squares and
triangles in Fig. 2B. The line in Fig. 2B, however, is a direct reproduction of the dashed line in
Fig. 1E—a previous experiment using the same extract under continuous transcription
conditions. It can be seen that the rate of 3′-end processing is similar both in the absence and
presence of on-going transcription. Therefore, once the polymerase has crossed the poly(A)
signal ongoing transcription is no longer required for rapid and efficient 3′-end processing.
Moreover, poly(A) tail growth does not contribute to the speed or efficiency of poly(A) site
cleavage in our coupled system.
Is there any role for transcription in this system, beyond merely producing the RNA that is to
be processed? For example, perhaps processing in these extracts is efficient simply because the
extracts are unusually effective at processing per se, or because they produce RNA having some
special property that facilitates processing. To address this we made 32P labeled RNA under
coupling conditions, used gel extraction to purify RNA of sufficient length to contain the
poly(A) site, and then added this back to a coupled reaction in which transcription had been
initiated with a cold pulse (Fig. 2C). We also added 3′-dATP to facilitate detection of any
cleaved RNA (i.e. so a sharp band low in the gel would be produced rather than a broad
polyadenylated band overlapping the unprocessed precursor). The results do not show any
poly(A) site cleavage of the pre-made RNA over at least 30 min (Fig. 2C). In contrast, when the
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larger amount of RNA synthesized in situ in an identical parallel sample was labeled, it could be
seen to undergo fast and efficient cleavage at the poly(A) site (Fig. 2D). Thus coupled, but not
uncoupled, processing under these conditions is fast and efficient.
To show that the RNA in Fig. 2C had not been damaged by the purification procedure, we
subjected some of the same RNA to standard uncoupled processing conditions in the presence of
polyvinyl alcohol (PVA). Fig. 2E shows that this gel-purified RNA is fully capable of poly(A)
site cleavage when conditions that drive uncoupled processing are used. Although we have
included PVA in some of our coupled processing reactions (e.g. Fig. 1), we now realize that,
while it does improve efficiencies, it is not a required ingredient for coupling. Thus, coupled
processing proceeds quickly in the absence of PVA (Fig. 2D) whereas uncoupled processing
requires PVA (compare Figs. 2C and 2E). Taken together these results show that some property
of the ternary transcription complex itself or of the associated RNA (as opposed to on-going
transcription) allows rapid processing to occur even in the absence of crowding agents such as
PVA.
The RNA tether from the poly(A) signal to the polymerase mediates coupling. To evaluate
the role of the transcription complex in coupling we decided to focus on its defining feature—the
nascent RNA. Specifically, we severed the RNA tether between the poly(A) signal and the
polymerase to see if this would impair coupling. To sever the tether we added to the
transcription mixture short DNA oligonucleotides complementary to sequences downstream of
the poly(A) signal (Fig. 3A). Hybrid formation by these oligos with their RNA targets then led
to cutting by the RNase H endogenous to the extract.
Fig. 3B shows the outline and the results of such an experiment. Five different oligos were
used. Three targeted cutting to positions 77, 158 and 397 nt downstream of the SV40 late
poly(A) site (see maps beside the gels in Fig. 3B). The other two oligos (one for each panel of
Fig. 3B) were controls, not complementary to any part of the RNA. All oligos were added with
the chase (30 s into the reaction) before any polymerases had reached the poly(A) signal (see Fig.
1A, lane 1).
Thus RNase H cutting began as soon as the oligo target was extruded from the
polymerase. After 3.5 min of chase, during the lag before cleavage begins (Fig. 1A, lane 3), αamanitin and 3′-dATP were added to facilitate a quantitative evaluation of the results (by
preventing new transcripts from entering the processing pool during the time course, and by
blocking polyadenylation so that cleaved RNA appears as a distinct band not overlapping the
RNase H-cut RNA higher in the gel).
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Lanes 4-6 and 13-15 in Fig. 3B show that poly(A) site cleavage remained fast and efficient in
the presence of the control oligos. In contrast the 77 oligo almost completely eliminated poly(A)
site cleavage at the 10 min time point (lane 2), and even after an hour (lane 3) only very little
poly(A) site cleavage was observed. Targeting the oligos farther and farther downstream
allowed a progressive rescue of poly(A) site cleavage (lanes 7-12).
We wanted to verify that the RNase H-cut RNA was not intrinsically incapable of being
cleaved at the poly(A) site (unlikely because RNase H cutting occurred in sequences from the
cloning vector). Therefore, we gel extracted the RNase H-cut RNA from bands like those in
lanes 1 and 10 of Fig. 3B and subjected it to standard uncoupled processing in the presence of
PVA (Fig. 3B had no PVA). Fig. 3C shows that both 77-cut and 397-cut RNA were processed
efficiently in PVA. Moreover, the efficiency of this uncoupled processing was comparable for
both RNAs, showing that cutting farther downstream rescues coupled processing (Fig. 3B) for
some reason other than the length increase per se of the cut RNA.
The SV40 late poly(A) signal is strong (Carswell and Alwine, 1989), having enhancer
elements both upstream and downstream of the hexamer and G/U-rich core elements (Lutz and
Alwine, 1994; Bagga et al., 1995). To assess the tether requirement for a weak poly(A) signal,
composed of core elements only, we carried out the experiment of Fig. 3D. Lanes 1 and 2 show
that the coupled in vitro system can support processing of this weak poly(A) signal. Lane 5
shows that when the tether was cut 129 nt downstream of the poly(A) site, little poly(A) site
cleaved RNA was produced.
The results of Fig. 3 suggest a model in which a tether is required to hold the poly(A) signal
close to the CTD during the early stages of cleavage complex assembly until this complex is
sufficiently mature to remain stably associated with the CTD on its own (Fig. 3A). We can also
envision a “structural” model that invokes a 3′-end processing complex that is large when
coupled to transcription, and that includes hundreds of nucleotides of downstream RNA.
According to this model, the complex needs to be large to support functions related to coupling,
and cutting interferes with its assembly. Coupled processing in the structure model would be
rescued upon cutting farther downstream (Fig. 3B) because assembling this large structure (on
the polymerase, in the simplest version of the model) would get easier as the 3′ extension on the
RNA gets longer (not true, recall, for uncoupled processing of these RNAs, Fig. 3C). This
contrasts with the tether model for which any RNase H-cut RNA, long or short, gets lost and can
never be processed efficiently if it is cut before becoming stably associated with the polymerase.
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For the tether model, targeting the RNase H farther downstream rescues processing because a
larger number of transcription complexes are able to assemble a mature cleavage complex before
the RNase H target is extruded.
We can distinguish between these two models by determining how efficiently the RNase Hcut RNA gets processed. In Fig. 3B the amount of RNase H-cut RNA decreases slowly with
time once cutting is complete (e.g. lanes 2-3, 8-9 and 11-12). This decrease could reflect nonspecific degradation, uncoupled processing, and/or coupled processing.
Non-specific
degradation can be accounted for by reference to RNase H-cut RNA having a mutant poly(A)
signal. The structure and the tether models can then be distinguished by comparison of RNAs
cut by RNase H at the 397 and the 77 oligo positions. Both of these RNAs are similarly
accessible to uncoupled processing (Fig. 3C), but the structure model predicts that the longer
397-cut RNA will more easily assemble the large apparatus required for coupled processing and
will, therefore, get processed and decrease in amount more rapidly than the 77-cut RNA. An
analysis of the data in Fig. 3B (after normalizing to parallel data for mutant RNAs) indicates,
however, that this is not the case, disfavoring the structure model.
To test the structure model explicitly, we carried out additional experiments (Fig. 4). In Fig.
3B the oligos had been added early, before the transcribing polymerases reached the poly(A) site,
for maximum effect. However, for Fig. 4, to restrict our attention to structural issues, the oligos
were not added until transcription was stopped with α-amanitin, just before the start of
processing. We then allowed 5 min for the RNase H to cut, and finally took time points to ask
whether 397-cut RNA is processed more rapidly than 77-cut RNA as required by the structure
model.
Fig. 4A shows the results from such an experiment. Lanes 1-3, 7-9 and 13-15 are exactly the
same as Fig. 3B except, importantly, as indicated by the time line. Fig. 4A confirms both the
inhibition of processing by RNase H cutting (lanes 1-3 and 7-9 vs. 13-15) and rescue from this
inhibition by cutting farther downstream (lanes 1-3 vs. 7-9). However, these effects are muted
relative to Fig. 3B because the oligos were added later, the overall time window was only half as
big, and the first time point was taken at a later time, after a significant proportion of whatever
processing would occur had already taken place. The effects of oligo cutting on processing are
summarized in the upper panel of Fig. 4B where we plot the amount of poly(A) cleaved RNA
produced after completion of RNase H cutting (i.e. after 5 min of oligo) over both short and long
time intervals. The data show that, like coupled processing itself, the effects of cutting the tether
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are not dependent upon on-going transcription at the time of cutting.
Recall, the purpose of the experiments in Fig. 4 was to test the prediction of the structure
model that the 397-cut RNA will get processed more efficiently than the 77-cut RNA. The
differences in RNase H-cut band intensities in Fig. 4A are not sufficient to see by eye, but can be
revealed by quantitation. The heavy bars in Fig. 4B (lower panel) represent the range of values
obtained for the decrease in these RNase H-cut RNAs over the two different time intervals (after
normalizing to the mutant RNAs in lanes 4-6 and 10-12). It is clear from the overlapping data
sets that there is no significant difference in the rates at which these RNAs disappear. In
particular, there is no support for the requirement in the structure model that the 397-cut RNA
should disappear faster than the 77-cut RNA.
Since the “rescue” that occurs upon moving the cutting downstream to the 397 position (Fig.
4B, upper panel) cannot be accounted for by superior processing of the 397-cut RNA, this
processing must arise from the ternary complexes stalled by α-amanitin at positions preceding
the 397 location on the template. Indeed, material can be seen to be disappearing from this
region of the gel in lanes 10-12 of Fig. 3B (these differences are difficult to see in Fig. 4A for the
reasons noted above). Thus, rescue presumably arises from ternary complexes in which the 397
target has not yet been exposed and in which the poly(A) signal remains tethered to the
polymerase.
RNA binds more tightly to the ternary complex after assembling a cleavage complex. The
tether model is based on the idea that the poly(A) signal does not become firmly associated with
the polymerase until late in the processing complex assembly pathway. To evaluate this idea we
examined the relative tendencies of the various RNA species in Fig. 3B to remain associated
with the ternary complex in pull-down experiments. The bands in Fig. 3B are of two
types—those arising from cleavage at the poly(A) site, and those resulting from cutting by
RNase H. The poly(A) site cleaved RNA, of course, is representative of RNA that was
successful in assembling a cleavage apparatus on its poly(A) signal. In contrast, the longer
RNase H-cut RNAs, which got cut before cleavage could occur at the poly(A) site, are
representative of RNAs that were not successful in assembling a functional cleavage apparatus.
We asked to what extent these two classes of RNA are pulled down with the ternary complex.
We began by using immobilized templates (Fig. 5B). To facilitate quantitation we included
an internal standard in our reactions in the form of an oligonucleotide targeted to a region 5′ of
the poly(A) signal (see Fig. 5A). The concentration of this oligo was chosen so that only a small
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proportion of the transcripts would be cut by RNase H at this up-stream location. The
experiments then consisted of determining how the two classes of transcript described above [i.e.
cleaved at the poly(A) site or cut within the tether by RNase H] partition between pellet and
supernatant relative to partitioning of the 5′ fragment. Transcription on the immobilized
templates was initiated with a pulse of [α-32P] CTP in the usual way (Fig. 5B) except that the
pulse was lengthened to give more counts in the transcripts and the oligos were added during the
pulse. Then α-amanitin and 3′-dATP were added during the processing phase of the reaction, as
before, to facilitate quantitation. Finally, magnetic selection was used to separate the templates
and their associated ternary complexes from any RNA that was released.
An inconvenience of these experiments is the presence in HeLa nuclear extracts of TTF2, an
ATP-dependent, poly(A)-independent transcript release factor (Jiang et al., 2004). We have
confirmed, using activation by ATP but not by AMPPNP (Xie and Price, 1997) that our extracts
contain such an activity (data not shown). The properties of this protein have been likened to
those of E. Coli Mfd (Hara et al., 1999; Jiang et al., 2004) which preferentially attacks stalled
polymerases (Park et al., 2002). Unfortunately, our need to block poly(A) tail growth [so as to
resolve poly(A) cleaved transcripts from those cut by RNase H] requires the use of 3′dATP—which stalls transcription (regardless of whether α-amanitin is also present). Although
we have shown that this stalling does not interfere with coupling (Figs. 2A and B), many of the
stalled transcription complexes get released from the immobilized templates before they can be
isolated for analysis. Moreover, the problem is exacerbated by the fact that magnetic beads
reduce processing efficiency (Yonaha and Proudfoot, 2000) which necessitates longer incubation
times. Consequently in Fig. 5B most transcripts are actually released (see lanes 1 and 3 of the
gel in Fig. 5B) so that the supernatant reflects primarily the overall composition of the sample
rather than the composition of a preferentially released fraction. Fortunately, however, the
significant observations in this experiment come from the pellets, which still contain sufficient
material to quantitate.
The gel in Fig. 5B shows that the pellet of a transcription-processing reaction is substantially
enriched for poly(A) site cleaved RNA (compare lane 1 with lane 2a). The pellet has twice as
much poly(A) site cleaved as 5′-cut RNA (line graph 2) whereas the supernatant has only one
third as much (line graph 1), for an enrichment of 6 fold (1.91/0.315) in this experiment (4 fold
on average). Note that these transcripts are cleaved but, because of the 3′-dATP, they are not
polyadenylated. Therefore, their preferential association with the immobilized templates is not
9
the result of poly(A) tail binding proteins.
If the preferential association of poly(A) site cleaved RNA with ternary complexes is
mediated by the cleavage apparatus, then RNAs that are unable to assemble this apparatus should
not preferentially associate with the ternary complex, even if they contain a poly(A) signal. To
test this we ran a parallel sample to the above in which we replaced the non-complementary
oligo with the 77 oligo. Importantly, Fig. 5B, lanes 1 and 3 show that cutting the tether using the
77 oligo uncouples transcription and processing for immobilized templates just as for circular
plasmid DNA (compare with Fig. 3B, lanes 1-6). As predicted by the tether model, the 77-cut
RNA, unlike the poly(A) site cleaved RNA, is distributed similarly between supernatant (0.93
relative to 5′, line graph 3) and pellet (0.81 relative to 5′, line graph 4), and is therefore not
preferentially retained on the ternary complex. Moreover, these same results show that whether
RNA contains a poly(A) signal or not, if it is not processed it behaves pretty much the same,
since the ratio of RNA cut at the 77 oligo position to that cut at the 5′ position is not much
different between supernatant and pellet (0.93 and 0.81). On average, RNA cleaved by
processing is enriched 5 fold (4.4/0.86) in the pellet compared to the similar, poly(A) signalcontaining RNA cut 77 nt downstream. Thus, it is apparent that cutting the tether with RNase H
interferes with a process that causes the poly(A) signal to become associated with the
polymerase.
To explore this further using a different method we carried out a polymerase pull-down
experiment. Transcription was carried out in the presence of the 397 oligo as shown in the time
line of Fig. 5C (essentially as for lane 11 of Fig. 3B). Then RNA polymerase II was pulled down
using an antibody to the N-terminus of its large subunit. Recall that the 397 oligo allows some
rescued processing to occur but that the 397-cut RNA itself is not efficiently processed (Fig. 4).
Lanes 1 and 2 of Fig. 5C show that most of the 397-cut RNA was left in the supernatant but
about half of the poly(A) cleaved RNA appeared in the pellet. Almost no RNA appeared in the
pellet if an irrelevant antibody or naked beads were used (Fig. 5C, lanes 4 and 6). In each of five
independent repeats of this experiment the poly(A) site cleaved RNA was enriched in the pellet
after pulling down the polymerase—on average 2.6 fold.
It is interesting that processed transcripts remain associated with the polymerase. This could
reflect the persistence of the entire processing apparatus on the polymerase even after cleavage,
or it could reflect the action of a sub-set of factors that bind the cleaved 5′ fragment. To evaluate
these alternatives we focused on CstF which binds the GU-rich element of the poly(A) signal
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downstream of the cleavage site. Since CstF would not be expected to remain associated with
the processed transcript unless much of the apparatus remains intact we pulled down its RNAbinding subunit (CstF 64) to see if the cleaved transcripts come along. A 5′ oligo was also
included for normalization as in Fig. 5B. Fig. 5D, lanes 2 and 4 show that, indeed, a substantial
amount of poly(A) site cleaved transcript, polyadenylated or not, was pulled down by CstF. In
contrast, transcripts cut by RNase H at the 5′ oligo position were not significantly pulled down
(lanes 2 and 4), and an irrelevant antibody pulled down nothing at all (lane 6). Apparently, the
cleavage apparatus remains at least partially intact even after half of the RNA sequence
responsible for its initial assembly has been removed.
Discussion
We have described an in vitro system in which fast, efficient and accurate cleavage and
polyadenylation is coupled to transcription. In addition we have shown, for two different
poly(A) signals (Figs. 3B and 3D), that an intact tether of nascent RNA from the poly(A) signal
to the polymerase (Fig. 3A) is required for this coupled processing. We also observed this tether
requirement for 3′-end processing when we used a transcription unit that exhibits active splicing
(data not shown). We suggest that the immature cleavage apparatus is unable to cling securely to
the polymerase on its own and requires a tether to hold the poly(A) signal close to the
polymerase until a mature, and stable, processing complex has formed (Figs. 4 and 5). A similar
idea has been suggested on the basis of experiments performed in vivo in which a ribozyme
rather than RNase H was used to cut the tether (David Bentley, personal communication). The
simplest model for cleavage apparatus assembly thus appears to be that the poly(A) signal is
extruded from the polymerase and then collaborates with the CTD to recruit factors (Kim et al.,
2004) and to assemble a complex that does not bind strongly to either the RNA or the CTD
alone. Of course, some factors may be recruited to the polymerase in advance of the appearance
of the poly(A) signal (Calvo and Manley, 2003).
But why have the decisive stages of assembly been designed to be so fragile? An attractive
possibility is that this is a manifestation of the previously proposed check-point activity of the
poly(A) signal (Orozco et al., 2002). Perhaps a tenuous assembly scheme allows the nascent
processing apparatus to sample multiple inputs before committing to a final course of action.
Though the assembly process is initially tentative, once mature, the association of the
apparatus with the polymerase apparently survives even cleavage at the poly(A) site (Fig. 5B and
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C). Adamson et al. (2005) have made a similar observation. Both Adamson et al. and we looked
at cleaved transcripts for which polyadenylation had been blocked.
However, once
polyadenylation occurs, the transcripts appear to be released (Yonaha and Proudfoot, 2000),
consistent with earlier results on uncoupled processing (Zarkower and Wickens, 1987).
Interestingly, these presumptively released polyadenylated transcripts appear to retain their
association with CstF (Fig. 5D, lane 4) consistent with a function for CstF beyond the cleavage
reaction itself (Moreira et al., 1998).
We must emphasize that the tether is only part of the coupling story. There are several
methods for preparing nuclear extract and a variety of conditions that can be used for
transcription. But although the tether exists in all of these, many of the conditions fail to give
rapid and efficient 3′-end processing, concurrent with transcription. Thus, the tether is required
for coupling, but it is not sufficient.
We began this study by optimizing for rate and efficiency of processing concurrent with
transcription. Mindful that under the right conditions processing can be fast and efficient in vitro
even when not coupled (Zarkower and Wickens, 1987) we sought functional connections beyond
a mere precursor-product relationship between the processing and the transcription (functional
coupling). Moreover, we wanted a criterion that points uniquely to the coupled state. For
example, both the cap and the CTD are required even for efficient uncoupled processing
(Flaherty et al., 1997; Hirose and Manley, 1998; Ryan et al., 2002) so a requirement for these
cannot be used as an indicator of coupling in vitro. The tether, however, is a unique signature of
the coupled state, so we directed our initial attention to the tether, and cut it using RNase H to see
if this would disrupt coupling.
Disrupting a functional connection to investigate functional coupling requires some caution
so that the only difference is the disrupted functional connection itself. Thus, the failure of RNA
to be processed efficiently after removal from the ternary complex by RNase H (Figs. 3B and 4)
provides strong evidence of coupling because it is not likely that the state of the RNA has been
altered beyond its removal from the ternary complex. Yonaha and Proudfoot (2000) have made
a comparable observation using immobilized templates. In contrast, reduced efficiency of
processing after removal of the RNA from the ternary complex by phenol
extraction—occasionally applied as a criterion for functional coupling (Adamson et al.,
2005)—is not a sufficient criterion because free RNA is unlikely to fold and associate with
proteins in the same way as RNA extruded from a polymerase. Very likely, newly extruded
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RNA is packaged in a way that makes it a preferred substrate for the next step of mRNA
production, but this is really coupling in the broader precursor-product sense. For this reason
studies directed at functional coupling often compare the processing of RNA produced by RNA
polymerase II with the processing of RNA produced by T7 RNA polymerase under the same
conditions (Mifflin and Kellems, 1991; Ahuja et al., 2001). However, even here caution is
required because, as discussed earlier, mere involvement of the polymerase as a participant in the
processing reaction is not synonymous with functionally coupling processing to transcription.
We have been careful to show that processing proceeds concurrently with transcription in our
system (Fig. 1A). Although for technical clarity we sometimes stopped transcription prior to
processing (e.g. Fig. 3B), the overall conditions were not otherwise changed, and care was taken
to demonstrate that this did not have a significant effect on processing (Fig. 2A and B). In one
recent study on functional coupling, transcription and processing were carried out under
markedly different conditions (Adamson et al., 2005). Apparently it was necessary to stop
transcription with EDTA and then add PVA in order to obtain robust poly(A) site cleavage.
However, at least for the SV40 late poly(A) site, we have found that PVA promotes rapid and
efficient processing in reactions that have been uncoupled by RNase H cutting (data not shown).
This is expected given the known ability of PVA to drive uncoupled poly(A) site cleavage
(Zarkower and Wickens, 1987; McLauchlan et al., 1988). Indeed, a hallmark of the coupled
processing we report here is that its rate and efficiency are not dramatically different in the
presence or absence of PVA (e.g. compare Fig. 3B, lanes 14-15 with Fig. 1E, diamonds). In
summary, after conditions have been changed, caution must be exercised in concluding that what
happens next is functionally coupled to what happened before.
Although the present study has a number of features in common with previous studies on the
coupling of 3′-end processing to transcription, it is difficult to make direct comparisons. For
example, like us, Mifflin and Kellems (1991) observed processing that was fast and efficient
when coupled to transcription, and Yonaha and Proudfoot (1999; 2000) and Ahuja et al. (2001)
observed functional interactions between processing and transcription. Nevertheless, it is not
possible to say if a tether requirement would have been evident in those studies because they all
employed crowding agents like PVA to enhance the processing. In some cases (but not always,
e.g. Fig. 3D), crowding agents can mask the effects of uncoupling by accelerating the uncoupled
reaction to levels that can even exceed those of coupled processing. Therefore, the rate and
efficiency of processing per se are not necessarily reliable indicators of coupling, at least when
13
PVA is involved. Instead, evidence of functional interactions must be sought. We think it likely
that PVA mimics an activity in nuclear extract that accelerates processing—but that this activity
only acts on processing that is coupled to transcription (i.e. with the tether still intact). In
contrast, PVA appears to promote rapid processing indiscriminately.
Experimental Procedures
Plasmids. In pSV40E/P the transcription unit and several kb of surrounding sequence are
identical to pAP〈117cat〉 (Tran et al., 2001). In pSV40E/L a SmaI-BamHI fragment containing L
(Tran et al., 2001) replaces the HpaI-BamHI fragment in pSV40E/P that contains P. In the
mutated P and L poly(A) signals, the AATAAA hexamers have been changed to AgTAct and
AAgtAc respectively.
For pCMV/BGH a 1778 bp PCR fragment was made from pcDNA3 (Invitrogen) using oligos
#1 and #2 as primers. Two cloning steps were then carried out, essentially to replace the
EcoO109I-SalI segment of pAP〈117cat〉 containing the SV40 early promoter with the 5′ portion
of this fragment (up to the XhoI site) containing the CMV promoter, and to replace the SalIBamHI segment of pAP〈117cat〉 containing the P poly(A) signal with the remainder of the PCR
fragment containing the BGH poly(A) signal. For the mutant BGH poly(A) signal the AATAAA
hexamer was changed to AgTAct.
DNA oligomers (5′→3′).
1) CGGATCGGGAGATCTCCCGATCCCCTATGG
2) GGACTTTCCACACCCTAACTGACACACATTCC
3) CTCAGACAATGCGATG
4) 77 oligo: GTAGGGAGTATTGGG
5) 158 oligo: TGGGAGTGGAATGAG
6) 397 oligo: CGGAATTCCGGATGAGCATTCATCAGGCGGGC
7) CTCATTCCACTCCCACCCGGGCAAGCTTTTCAGGAGCTAAGG
8) CAACTAGAATGCAGTG
9) -147 oligo: CGAGGTCGACAGTGGTACTCGTGGGCCAGC
10) -181 oligo: CCATCTTCTGCCAGG
11) AAACAAATAGGGGTTCCGCGCACATTTCCC
12) GGTATCGATAAGCTGATCTCATGCACCATTCG
Coupled processing assay. HeLa nuclear extract was prepared as described (Tran et al., 2001).
14
This is a very crude extract. For the success of our previously reported elongation assay
("signaling", Tran et al., 2001) we found it essential to conduct the final centrifugation at a much
lower speed than called for in the earlier protocol that we followed (Flaherty et al., 1997). We
continued that practice for these studies although we have not determined whether it is as
important for coupled processing as it is for signaling. We did find, however, that for efficient
coupled processing it was particularly important to achieve complete cell lysis and to remove all
material above the nuclear pellet after centrifugation.
A typical pulse-chase assay began with 3 µl of nuclear extract that was mixed with antiRNase (Ambion), DTT, MgCl2, sodium citrate, DNA and water up to 5.9 µl. Amounts of
magnesium, citrate and nuclear extract were individually optimized for each extract preparation.
The mixture was pre-incubated at 30°C for 30 min and then pulsed with 3 µl containing 20 µCi
of [α-32P] CTP, nucleotide triphosphates and creatine phosphate. Then 3.6 µl of chase mix was
added containing a high concentration of non-radiolabeled CTP. Final concentrations in a
standard pulse-chase assay (unless otherwise noted) were as follows: 4.8% glycerol, 4.8 mM
HEPES (pH 7.9), 24 mM KCl, 48 µM EDTA, 2.1 mM DTT, 24 µM PMSF, 10 U anti-RNase, 4
mM MgCl2, 3 mM sodium citrate (pH 6.7), 0.3 µg DNA, 200 µM each of ATP, UTP and GTP,
20 mM creatine phosphate and 2 mM CTP. PVA or DNA oligonucleotides, when used, were
usually added with the chase. When α-amanitin and 3′-dATP were used, they were added in a 1
µl volume. The final concentrations of these additions, if used, were: 2.1% PVA, 8 ng/µl DNA
oligo, 37 ng/µl α-amanitin and 400 µM 3′-dATP.
In vitro transcription was terminated by the addition of a “stop solution”: 65 µl of 10 mM
TrisHCl, 10 mM EDTA, 0.5% SDS, and 100 µg proteinase K (Ambion). After 30 min at 30°C
the RNA was extracted with 350 µl TRIzol (Invitrogen), 70 µl chloroform, then precipitated with
4 µl of 5 mg/ml glycogen (Ambion) and 350 µl isopropanol (30 min, room temperature), and
finally run on a 5% polyacrylamide gel. Following electrophoresis, results were recorded and
analyzed using a PhosphorImager with ImageQuant software (Molecular Dynamics). No
Photoshop was used.
Immobilized template experiments. PCR was carried out using oligos #11 and #12 as primers
on pSV40E/L DNA following the manufacturer’s protocol for ThermalACE (Invitrogen) except
that 2 U each of Taq and Pfu polymerase were used. Primer #11 was biotinylated. The PCR
products were purified using agarose gels and a Qiagen Gel Extraction kit. The eluted DNA was
then bound to Dynabeads M-280 Streptavidin (Dynal) using the manufacturer’s protocol. The
15
beads were then washed 2 times with 400 µl of Dynal Washing Buffer, followed by washing
once, rotating 10 h and washing again 4 times with 400 µl of 10 mM Tris, 1 mM EDTA, pH 8.
Attachment was confirmed by digesting the bead-bound DNA with restriction enzymes.
Approximately 0.5 pmol of DNA was attached per mg of bead. Magnetic selection, following
transcription, was allowed to proceed for 2 min. The bead fraction (pellet) was then washed
twice with vortexing in 50 µl of buffer D (nuclear extract dialysis buffer), and these washes were
combined with the original supernatant to which 65 µl of the stop solution was added. The beads
were also incubated in stop solution at 30ºC for 30 min to liberate the template-associated RNA,
and then the beads were removed prior to isolation of the RNA.
Antibody pull-down experiments. For the polymerase pull-down, 20 µl of protein A or G
magnetic beads (Dynal) were incubated overnight at 4°C with 20 µl of anti-RNAP II (N-20,
Santa Cruz Biotechnology), anti-E1B hybridoma supernatant (2A6, Dass et al., 2001), or buffer
alone. The beads were washed twice with 100 µl PBS (phosphate-buffered saline), mixed with
13.5 µl of transcription mixture and then incubated for 20 min at room temperature. Then the
bead-bound fraction (pellet) was magnetically selected, washed gently with 100 µl of PBS, and
the RNA in the pellet and supernatant isolated as above. The CstF pull-down was the same
except that 40 µl of anti-CstF 64 (a mixture of 3A7 and 6A9, Wallace et al., 1999) or anti-E1B
hybridoma supernatant were used in a 12 min pull-down, and the wash was combined with the
supernatant for analysis.
Acknowledgments
We thank Clint MacDonald for the 3A7, 6A9 and 2A6 antibodies, David Tsao for plasmids, and
David Bentley for communicating unpublished results. This work was supported by NIH grant
GM50863.
Figure Legends
Figure 1. Transcription-coupled 3′-end processing is fast, efficient, accurate and reproducible.
(A) Circular plasmid DNA was transcribed for increasing lengths of time and the RNA was
displayed on a gel. The % poly(A) refers to the ratio of polyadenylated RNA to all RNA
extending past the poly(A) site. This assay contained PVA and the [MgCl2] was 5 mM.
(B) Wt and mt refer to a poly(A) signal with intact or mutated hexamer respectively. For lanes
2-7 RNA was isolated from a 15 min, 5 fold transcription-processing reaction as for (A) that had
2 µM of cold CTP in place of [α-32P] CTP. One third of the RNA was set aside as the Input and
16
the remainder was oligo(dT) selected using a Poly(A) Purist MAG kit (Ambion). The Free
RNA was taken from a first round of selection and the Bound RNA was from the second. The
Input, Free, and Bound fractions were then digested with DNase I (Roche), hybridized at 65°C,
and subjected to RNase protection using RNase T1 (Chao et al., 1999). The mole % of
processing in lane 2 is 30% of the total. The DNA used here, pSV40E/L′, had the same promoter
and poly(A) signal as in (A) but set in a different plasmid background that provided an intron to
allow for expression in vivo. Lane 1 shows a control using cytoplasmic RNA, isolated from
transfected cells as previously described (Park et al., 2004). The probe used was a run-off
transcript from a derivative of pSV40E/L′ containing an inserted T7 RNA polymerase promoter.
Since only RNase T1 (specific for G residues) was used in the RNase protection a single probe
sufficed for both wt and mt RNAs. The intensities of lanes 6 and 7 were reduced in ImageQuant
for purposes of comparison.
(C) This assay was like that in (A) but using DNA with a different promoter and poly(A) signal,
and at [citrate] and [MgCl2] of 5 and 6 mM respectively.
(D) Cleavage at the BGH poly(A) signal is accurate. Reactions were performed as in (C) with
pCMV/BGH having either a wildtype or mutant poly(A) signal. For samples receiving 3′-dATP
to block polyadenylation, α-amanitin was also added, simply because this had become part of a
standard procedure (see Fig. 2). Lane 4 was as for lane 3 except that oligo #3 was added with
the chase to direct RNase H cutting to the BGH poly(A) site.
(E) Transcription-polyadenylation as a function of time for different extracts, promoters and
poly(A) signals. The % polyadenylation is quantitated as for Fig. 1A. The data for extract 1 are
from an experiment that differed from the standard assay in having a 15 min pre-incubation in a
volume of 6.9 µl, a chase of 2.6 µl containing PVA, and a final [ATP] of 500 µM. PVA was also
used in the experiment for extract 2. The data for extract 3 are from the gels in Figs. 1A and 1C
and include some time points not shown in Fig. 1A.
Figure 2. Coupling requires a ternary complex but not ongoing transcription.
(A) Ongoing transcription and poly(A) tail growth are not required for coupling. The assay was
done using pSV40E/L, extract 1 and PVA.
(B) Quantitative comparison of processing efficiency with and without transcription. The data
points are from Fig. 2A and the line is from Fig. 1E.
(C) Exogenous RNA added to a coupled reaction does not get processed. Pre-made 32P-labeled
RNA was isolated from a 10 fold coupled processing reaction of pSV40E/L in extract 3. RNA
17
running slower than poly(A) cleaved RNA was purified from a 5% polyacrylamide gel and
15,000 CPM of the RNA was added to a standard (i.e. containing no PVA) coupled processing
reaction in extract 3 (along with α-amanitin to 34 ng/µl and 3′-dATP to 372 µM) that was pulsed
with cold rather than hot CTP. The resulting gel was exposed 3 days to the phosphor screen.
(D) Exogenous RNA is not inhibitory. A reaction was carried out in parallel with the above
which differed only in that the coupled reaction to which the gel-purified RNA was added was
pulsed with 32P as usual. This gel was exposed only for 8 h to the phosphor screen and the newly
made RNA accounts for over 97% of the signal.
(E) To demonstrate that the gel-purified RNA is capable of under-going processing under special
conditions it was incubated at 37°C with PVA for 2 h under standard uncoupled processing
conditions (e.g. Wahle and Keller, 1994). Final amounts or concentrations in 30 µl were 2.5 µg
tRNA, 0.67 mM 3′-dATP, 17 mM creatine phosphate, 1.9 mM DTT, 10 U anti-RNase (Ambion),
42 µM PMSF, 2.1% PVA, 8.3% glycerol, 8.3 mM Hepes (pH 7.9), 42 mM KCl, 83 µM EDTA,
12.5 µl extract 3 and 6.25 µl PBS.
Figure 3. Severing the RNA tether from the poly(A) signal to the polymerase disrupts coupling.
(A) Cartoon of a ternary elongation complex in the process of assembling a cleavage and
polyadenylation apparatus. A DNA oligonucleotide is shown hybridized to a target in the RNA
thereby directing RNase H to cut the tether.
(B) Severing the tether prevents coupled 3′-end processing. The oligonucleotide names refer to
the distance from the principal poly(A) cleavage site to the predominant RNase H cutting site
(Wu et al., 1999). The control oligos for lanes 4-6 and 13-15 were the 77 oligo-complement and
oligo #7 respectively.
(C) RNase H-cut RNA is not intrinsically resistant to processing. RNase H-cut RNA was
generated as in (B) using a 10 fold coupled processing reaction. The bands were gel purified and
incubated under uncoupled processing conditions for 2 h with PVA. The % processing given is
mole % of the total.
(D) Tether requirement for a weak poly(A) signal. This assay used PVA, with [citrate] and
[MgCl2] of 4 and 5 mM respectively. Lane 4 was as for lane 3 except that oligo #8 was added
with the chase to direct RNase H cutting to the poly(A) site. The 129 oligo here is the same as
the 158 oligo in Fig. 3B, but cloning has placed the identical cut site closer to the poly(A) site in
this construct.
Figure 4. The RNA tether mediates coupling.
18
Wt and mt refer to a poly(A) signal with intact or mutated hexamer respectively. Three
independent experiments like that shown in part A were carried out (except that lanes 13-15 were
lacking in one). The increases in poly(A) cleaved RNA [as a % of all RNA extending past the
poly(A) site] between 5 min and either 10 or 30 min in the presence of the 77 and 397 oligos and
the control oligo (77-oligo complement) are plotted as the standard deviations in the upper panel
of part B. The decreases in RNase H-cut RNA are plotted similarly in the lower panel. In each
experiment the % of decrease in the RNase H-cut RNA was adjusted by subtracting the amount
of decrease observed for the corresponding poly(A) signal mutant RNA.
Figure 5. Processed RNA remains associated with the polymerase and the processing apparatus.
(A) Cartoon of an elongation complex. The template for all parts of this figure was pSV40E/L.
(B) Processed RNA is preferentially retained with the template. Transcription was initiated in
extract 2. At 30 s 0.005 µg of the -181 (5′) oligo and 0.05 µg of either the control oligo (oligo
#7, lanes 1 and 2) or the 77 oligo (lanes 3 and 4) were added. PVA causes all cut RNA, short
and long, to remain template-associated and was therefore not used. Bound RNA (pellet, lanes 2
and 4) was separated from released RNA (supernatant, lanes 1 and 3) by magnetic selection. The
averages given are the mean ± the difference from the mean for two independent experiments
using two different 5′ oligos (-181 or -147).
(C) Processed RNA is preferentially retained by the polymerase (extract 2). The error shown is
the standard deviation.
(D) Processed RNA remains preferentially associated with CstF (extract 4). This experiment
was carried out using 5 mM MgCl2, 4 mM citrate and 0.013 µg of the -181 (5′) oligo.
References
Adamson, T. E., Shutt, D. C., and Price, D. H. (2005). Functional coupling of cleavage and
polyadenylation with transcription of mRNA. J Biol Chem 280, 32262-32271.
Ahuja, D., Karow, D. S., Kilpatrick, J. E., and Imperiale, M. J. (2001). RNA Polymerase IIdependent Positional Effects on mRNA 3' End Processing in the Adenovirus Major Late
Transcription Unit. J Biol Chem 276, 41825-41831.
Bagga, P. S., Ford, L. P., Chen, F., and Wilusz, J. (1995). The G-rich auxiliary downstream
element has distinct sequence and position requirements and mediates efficient 3' end premRNA processing through a trans-acting factor. Nucleic Acids Res 23, 1625-1631.
Bentley, D. L. (2005). Rules of engagement: co-transcriptional recruitment of pre-mRNA
processing factors. Curr Opin Cell Biol 17, 251-256.
19
Calvo, O., and Manley, J. L. (2003). Strange bedfellows: polyadenylation factors at the
promoter. Genes & Dev 17, 1321-1327.
Carswell, S., and Alwine, J. C. (1989). Efficiency of utilization of the simian virus 40 late
polyadenylation site: effects of upstream sequences. Mol Cell Biol 9, 4248-4258.
Chao, L. C., Jamil, A., Kim, S. J., Huang, L., and Martinson, H. G. (1999). Assembly of the
cleavage and polyadenylation apparatus requires about 10 seconds in vivo and is faster for
strong than for weak poly(A) sites. Mol Cell Biol 19, 5588-5600.
Cooke, C., Hans, H., and Alwine, J. C. (1999). Utilization of splicing elements and
polyadenylation signal elements in the coupling of polyadenylation and last-intron removal.
Mol Cell Biol 19, 4971-4979.
Dantonel, J. C., Murthy, K. G., Manley, J. L., and Tora, L. (1997). Transcription factor TFIID
recruits factor CPSF for formation of 3' end of mRNA. Nature 389, 399-402.
Dass, B., Attaya, E. N., Michelle Wallace, A., and MacDonald, C. C. (2001). Overexpression of
the cstf-64 and cpsf-160 polyadenylation protein messenger rnas in mouse male germ cells.
Biol Reprod 64, 1722-1729.
Flaherty, S. M., Fortes, P., Izaurralde, E., Mattaj, I. W., and Gilmartin, G. M. (1997).
Participation of the nuclear cap binding complex in pre-mRNA 3' processing. Proc Natl Acad
Sci USA 94, 11893-11898.
Hammell, C. M., Gross, S., Zenklusen, D., Heath, C. V., Stutz, F., Moore, C., and Cole, C. N.
(2002). Coupling of termination, 3' processing, and mRNA export. Mol Cell Biol 22, 64416457.
Hara, R., Selby, C. P., Liu, M., Price, D. H., and Sancar, A. (1999). Human transcription release
factor 2 dissociates RNA polymerases I and II stalled at a cyclobutane thymine dimer. J Biol
Chem 274, 24779-24786.
Hirose, Y., and Manley, J. L. (1998). RNA polymerase II is an essential mRNA polyadenylation
factor. Nature 395, 93-96.
Jiang, Y., Liu, M., Spencer, C. A., and Price, D. H. (2004). Involvement of transcription
termination factor 2 in mitotic repression of transcription elongation. Mol Cell 14, 375-385.
Kim, M., Ahn, S. H., Krogan, N. J., Greenblatt, J. F., and Buratowski, S. (2004). Transitions in
RNA polymerase II elongation complexes at the 3' ends of genes. EMBO J 23, 354-364.
Lutz, C. S., and Alwine, J. C. (1994). Direct interaction of the U1 snRNP-A protein with the
upstream efficiency element of the SV40 late polyadenylation signal. Genes & Dev 8, 576-586.
20
Maniatis, T., and Reed, R. (2002). An extensive network of coupling among gene expression
machines. Nature 416, 499-506.
Manley, J. L., Sharp, P. A., and Gefter, M. L. (1982). RNA synthesis in isolated nuclei:
Processing of adenovirus serotype 2 late messenger rna precursors. J Mol Biol 159, 581-599.
McCracken, S., Fong, N., Yankulov, K., Ballantyne, S., Pan, G., Greenblatt, J., Patterson, S. D.,
Wickens, M., and Bentley, D. L. (1997). The C-terminal domain of RNA polymerase II
couples mRNA processing to transcription. Nature 385, 357-361.
McLauchlan, J., Moore, C. L., Simpson, S., and Clements, J. B. (1988). Components required for
in vitro cleavage and polyadenylation of eukaryotic mRNA. Nucleic Acids Res 16, 5323-5344.
Mifflin, R. C., and Kellems, R. E. (1991). Coupled transcription-polyadenylation in a cell-free
system. J Biol Chem 266, 19593-19598.
Moore, C. L., and Sharp, P. A. (1985). Accurate cleavage and polyadenylation of exogenous
RNA substrate. Cell 41, 845-855.
Moreira, A., Takagaki, Y., Brackenridge, S., Wollerton, M., Manley, J. L., and Proudfoot, N. J.
(1998). The upstream sequence element of the C2 complement poly(A) signal activates mRNA
3' end formation by two distinct mechanisms. Genes & Dev 12, 2522-2534.
Niwa, M., Rose, S. D., and Berget, S. M. (1990). In vitro polyadenylation is stimulated by the
presence of an upstream intron. Genes & Dev 4, 1552-1559.
Nodwell, J. R., and Greenblatt, J. (1991). The nut site of bacteriophage lambda is made of RNA
and is bound by transcription antitermination factors on the surface of RNA polymerase. Genes
& Dev 5, 2141-2151.
Orozco, I. J., Kim, S. J., and Martinson, H. G. (2002). The poly(A) signal, without the assistance
of any downstream element, directs RNA polymerase II to pause in vivo and then to release
stochastically from the template. J Biol Chem 277, 42899-42911.
Park, J. S., Marr, M. T., and Roberts, J. W. (2002). E. coli Transcription repair coupling factor
(Mfd protein) rescues arrested complexes by promoting forward translocation. Cell 109, 757767.
Park, N. J., Tsao, D. C., and Martinson, H. G. (2004). The Two Steps of Poly(A)-Dependent
Termination, Pausing and Release, Can Be Uncoupled by Truncation of the RNA Polymerase
II CTD. Mol Cell Biol 24, 4092-4103.
Proudfoot, N. (2004). New perspectives on connecting messenger RNA 3' end formation to
transcription. Curr Opin Cell Biol 16, 272-278.
21
Proudfoot, N. J., Furger, A., and Dye, M. J. (2002). Integrating mRNA processing with
transcription. Cell 108, 501-512.
Ruegsegger, U., Blank, D., and Keller, W. (1998). Human pre-mRNA cleavage factor Im is
related to spliceosomal SR proteins and can be reconstituted in vitro from recombinant
subunits. Mol Cell 1, 243-253.
Ryan, K., Murthy, K. G., Kaneko, S., and Manley, J. L. (2002). Requirements of the RNA
polymerase II C-terminal domain for reconstituting pre-mRNA 3' cleavage. Mol Cell Biol 22,
1684-1692.
Takagaki, Y., and Manley, J. L. (2000). Complex protein interactions within the human
polyadenylation machinery identify a novel component. Mol Cell Biol 20, 1515-1525.
Tran, D. P., Kim, S. J., Park, N. J., Jew, T. M., and Martinson, H. G. (2001). Mechanism of
poly(A) signal transduction to RNA polymerase II in vitro. Mol Cell Biol 21, 7495-7508.
Wahle, E. (1995). Poly(A) tail length control is caused by termination of processive synthesis. J
Biol Chem 270, 2800-2808.
Wahle, E., and Keller, W. (1994). 3' end-processing of mRNA. In RNA processing: A practical
approach, S. J. Higgins, and B. D. Hames, eds. (New York, Oxford University Press), pp. 1-34.
Wallace, A. M., Dass, B., Ravnik, S. E., Tonk, V., Jenkins, N. A., Gilbert, D. J., Copeland, N.
G., and MacDonald, C. C. (1999). Two distinct forms of the 64,000 Mr protein of the cleavage
stimulation factor are expressed in mouse male germ cells. Proc Natl Acad Sci USA 96, 67636768.
Whitelaw, E., and Proudfoot, N. (1986). α-thalassaemia caused by a poly(A) site mutation
reveals that transcriptional termination is linked to 3' end processing in the human α2 globin
gene. EMBO J 5, 2915-2922.
Wu, H., Lima, W. F., and Crooke, S. T. (1999). Properties of cloned and expressed human
RNase H1. J Biol Chem 274, 28270-28278.
Xie, Z., and Price, D. (1997). Drosophila factor 2, an RNA polymerase II transcript release
factor, has DNA-dependent ATPase activity. J Biol Chem 272, 31902-31907.
Yonaha, M., and Proudfoot, N. J. (1999). Specific transcriptional pausing activates
polyadenylation in a coupled in vitro system. Mol Cell 3, 593-600.
Yonaha, M., and Proudfoot, N. J. (2000). Transcriptional termination and coupled
polyadenylation in vitro. EMBO J 19, 3770-3777.
Zarkower, D., and Wickens, M. (1987). Specific pre-cleavage and post-cleavage complexes
22
involved in the formation of SV40 late mRNA 3' termini in vitro. EMBO J 6, 4185-4192.
Zhao, J., Hyman, L., and Moore, C. (1999). Formation of mRNA 3' ends in eukaryotes:
mechanism, regulation, and interrelationships with other steps in mRNA synthesis. Microbiol
Mol Biol Rev 63, 405-445.
23
E) Figure
Figure 1
A
SV40 early
promoter
B
SV40 late
poly(A) site
pSV40E/L
15 min transcription (extract 3)
Oligo(dT) selection
846 bp
32P
pulse
Extract 3
Time (min)
1
2
0
4
Chase
6
RNase protection
Time
30 s
8 10 12 14 16 20 60
Poly(A) site
Oligo(dT) s election
Input
Bound
Free
wt wt mt wt mt wt mt
0
Lane #
1 2 3 4 5 6 7 8 9 10 11
1
9 16 21 26 29 36 43
BGH
poly(A) site
CMV
promoter
pCMV/BGH
934 bp
32P
pulse
Extract 3
0
Time (min)
Chase
Time
30 s
1 2 3 4 5 6 7 8 9
D
32P
pulse Chase (-/+) 3'-dATP
0
pCMV/BGH
Extract 3
5 10 15 20 30 60
30 s
wt
20 min
wt
mt
1 2
3
~1.2 kb
Poly(A) RNA
Poly(A) RNA
Poly(A) site
% Poly(A) 0
4 min
Marker
0
+ 3'-dATP
0
No 3'-dATP
C
% Poly(A)
Probe alone
In vitro
In vivo
~1.1 kb
Poly(A) RNA
Probe + RNase
Probe (350 nt)
Uncleaved (300 nt)
Cleaved (242 nt)
21 31 40 47 54
Poly(A) site
cleaved
E
Promoter
Poly(A)
signal
4
E) Figure
Figure 2
A
32P
pulse Chase
0
30 s
(+/ -)
α-ama
3'-dATP
No
additions
With
α-ama, 3'-dATP
Total (min)
time
B
Total
time
4 min
4.5
9
14
24
14
24
1
2
3
4
5
6
Poly(A) RNA
Poly(A) site
cleaved
C
D
32P
pulse Chase
E
32P
pulse Chase
Pre-made 32P RNA
Cold
pulse
-4
α-ama (min)
3'-dATP
0
-2
0.5
α-ama
3'-dATP
Chase
5
10
30
Pre-made RNA
32 P
Time (min)
α-ama
3'-dATP
pulse Chase
-4
0
-2
α-ama (min)
3'-dATP
0.5
5
10
30
Time (min)
2 hr
Poly(A) site
cleaved
Poly(A) site
cleaved
Long exposure
Pre-made 32P RNA
Short exposure
Nascent 32P RNA
Control
E) Figure
Figure 3
A
C
32P
Chase +
pulse DNA oligo
0s
Tether
30 s
4 min
Gel purify the
RNase H-cut
RNA bands.
RN a se H

DNA oligo
CPSF
Uncoupled
processing
Cs t F
RNase H-cut RNA
2 hr
2 hr
Input
Input
77 oligo 397 oligo
Po l y(A) si g n a l
397 oligo RNase H-cut
77 oligo RNase H-cut
Poly(A) site cleaved
B
SV40 early
promoter
SV40 late
poly(A) site
% Poly(A) cleaved
46
44
pSV40E/L
32P
Chase +
pulse DNA oligo
Extract 2
-4
-3.5
α-ama
3'-dATP
Time (min)
0
D
Oligos for RNase H cutting
77 oligo
α-ama
(min)
3'-dATP
0.5 10
Control oligo
60 0.5 10
60
Weak
poly(A) site
SV40 early
promoter
pSV40E/P
3'
756 bp
(-/+)
α-ama
3'-dATP
32P
pulse Chase
Extract 3
0
30 s
4 min
20 min
0
1
5
0
14
27
Lane #
1
2
3
4
5
6
5'
wt
wt mt
Marker
% Poly(A)
129 oligo
Poly(A) site
+ 3'-dATP
Poly(A) site
cleaved
No 3'-dATP
Oligo to direct RNase H cutting
77 nt past poly(A) site.
wt
3'
Oligos for RNase H cutting
158 oligo
α-ama
(min)
3'-dATP
0.5 10
397 oligo
60 0.5 10
Control oligo
60 0.5 10
60
Poly(A) RNA
(~1 kb)
3'
129 oligo
Poly(A) site
cleaved
397
158
Poly(A) site
cleaved
% Poly(A)
Oligos for
RNase H
cutting
Poly(A) site
2
Lane # 7
10 16
8
2
14
21
2
19
28
9 10 11 12 13 14 15
5'
1 2 3 4 5
5'
E) Figure
Figure 4
5
10 30
5
10 30
wt
5
10
30 5
10 30
wt
5
10 30
397
RNase Hcut RNA
77
Poly(A) site
cleaved RNA
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15
8%
6%
4%
2%
0
- 2%
- 4%
- 6%
- 8%
397
Control oligo
mt
397
α-ama (min)
3'-dATP
mt
77
397 oligo
77 oligo
wt
10%
77
Oligos for RNase H cutting
Between
5 & 30 min
control oligo
12%
30 min
control oligo
0
397
Chase
-4
Time interval
Between
5 & 10 min
77
-5
B
30
77
pulse
10
α-ama
3'-dATP
DNA oligo
Increase in
poly(A) cleaved RNA
32P
5
397
Time points
Extract 2
Decrease in
RNase H- cut RNA
(wt relative to mt)
A
E) Figure
Figure 5
A
C
RNAPII pull-dow n
32P
pulse
5' oligo

77 oligo

0
Chase +
397 oligo
4 min
30 s
Tether
S
α-ama
3'-dATP Pull-down
α-RNAPII
15 min
P
No antibody
α-E1B
S
P
S
P
S
P
1
2
3
4
5
6
3'
CPSF
Cs t F
397 oligo

Po l y(A) si g n a l
397 oligo
Poly(A) site
cleaved
B
Immobilized template
SV40 early SV40 late
promoter poly(A) site
-299
0
5'
846 bp
32P
pulse
0
10 min
38 min
P
DNA oligos
at 30 s
3'
397 oligo
S
Magnetic
selection
α-ama
3' -dATP
Chase
2 min
Magnetic
bead
3480
Poly(A) site
cleaved
Pellet
Supernatant
Poly(A) site
397 oligo
1
S
0.29
2
P
0.78
This
expt. Average
2.7
Control oligo
Control
oligo
77 oligo
S
S
P
P
2.6 ± 0.8
77 oligo
P
P
D
77 oligo
Poly(A) site
cleaved
CstF pull-dow n
S
32P
α-ama
Chase +
pulse DNA oligo (+/-) 3' -dATP Pull-down
Longer exposure
of lanes 2 & 4
0
2 min
5 min
15 min
P
5' Oligo
α-CstF 64
5'
1 2
3 4
2a 4a
α-E1B
No 3'-dATP + 3'-dATP
S
P
S
P
S
P
1
2
3
4
5
6
3'
Poly(A) site cleaved (control oligo)
5' oligo
+ 3'-dATP
Poly(A) site
5' oligo
Poly(A) site
Pellet
Supernatant
This Average
expt.
Poly(A) RNA
1
S
0.315
6.1
P
2
RNase H-cut (77 oligo)
1.91
77 oligo
5' oligo
77 oligo
5' oligo
Poly(A) site
cleaved
4.4 ± 1.6
Poly(A) site
77 oligo
5' oligo
5.1 ± 1.9
3
S
5'
0.93
0.87 0.86 ± 0.01
4
P
0.81