© 2002 Oxford University Press
Nucleic Acids Research, 2002, Vol. 30, No. 3
803–809
In vitro transcription of a torsionally constrained
template
Thomas Bentin and Peter E. Nielsen*
Center for Biomolecular Recognition, IMBG, Laboratory B, The Panum Institute, University of Copenhagen,
Blegdamsvej 3c, 2200 Copenhagen N, Denmark
Received August 28, 2001; Revised and Accepted December 4, 2001
ABSTRACT
RNA polymerase (RNAP) and the DNA template must
rotate relative to each other during transcription
elongation. In the cell, however, the components of
the transcription apparatus may be subject to rotary
constraints. For instance, the DNA is divided into
topological domains that are delineated by rotary
locked boundaries. Furthermore, RNAPs may be
located in factories or attached to matrix sites limiting
or prohibiting rotation. Indeed, the nascent RNA alone
has been implicated in rotary constraining RNAP. Here
we have investigated the consequences of rotary
constraints during transcription of torsionally
constrained DNA by free RNAP. We asked whether or
not a newly synthesized RNA chain would limit
transcription elongation. For this purpose we
developed a method to immobilize covalently closed
circular DNA to streptavidin-coated beads via a
peptide nucleic acid (PNA)–biotin conjugate in
principle mimicking a SAR/MAR attachment. We
used this construct as a torsionally constrained
template for transcription of the beta-lactamase gene
by Escherichia coli RNAP and found that RNA
synthesis displays similar characteristics in terms of
rate of elongation whether or not the template is
torsionally constrained. We conclude that transcription of a natural bacterial gene may proceed
with high efficiency despite the fact that newly
synthesized RNA is entangled around the template in
the narrow confines of torsionally constrained supercoiled DNA.
INTRODUCTION
The helical structure of DNA necessitates relative rotary
movement between RNA polymerase (RNAP) and the DNA
template during transcription elongation. This phenomenon
has been recognized for years (1) and indirectly addressed (2),
but only recently has this right-handed rotary process been
observed directly (3). While the mechanistic details remain to
be elucidated, our current understanding of rotary motion
during transcription elongation may be simplistically pictured
with a nut (RNAP) on a bolt (DNA).
Four major topological modes for rotary movements during
transcript extension exist (Fig. 1) (reviewed in 4). In the
simplest case both RNAP and DNA are unrestrained and may
rotate freely (Fig. 1A). This situation presumably applies to
simple phage and bacterial in vitro transcription systems that
employ small DNA fragments and purified enzyme. However,
such a degree of freedom is unlikely to exist in cells. Second,
RNAP may be immobilized and the template may rotate freely
(Fig. 1B). This case has been studied in vitro using phage
RNAP attached to microbeads via an engineered immobilization
domain (5). Transcript extension occurred as efficiently as when
using enzyme liberated from the support prior to elongation.
Another approach involves Escherichia coli RNAP adsorbed
to glass slides (6–8). Single molecule measurements showed
elongation rates identical to that found using free enzyme (3).
Third, the DNA may be torsionally constrained while RNAP
rotates (Fig. 1C). In this scenario, newly synthesized RNA is
entangled around the DNA template. This model provides at
least two topological problems: If the newly synthesized RNA
produces hydrodynamic friction that inhibits free rotation of
RNAP (9,10), then the rate of transcription elongation could
decrease. Efficient translation could also require a faithful
untangling mechanism. Fourth, both RNAP and DNA may be
rotationally constrained (Fig. 1D). The DNA template is then
expected to experience torsional tension and become progressively
more positively and negatively supercoiled ahead of and
behind RNAP, respectively (1).
While recent evidence suggests that RNAPs can be immobilized
in multi-enzyme factories [the data being especially compelling
for RNAP I transcription (4)] it is difficult to distinguish firmly
between the depicted modes. It could well be that the issue of
rotation depends on the particular transcription system in
question and the state of that system.
RNAPs share notable sequence and structural similarities.
For example, prokaryotic, chloroplast, archaebacterial and
eukaryotic RNAPs have homologous largest and second
largest subunits (11 and references therein) and sequence
alignment of bacteriophage T3, T7 and K11 RNAPs shows
66% identity, SP6 RNAP being more distantly related. In contrast,
RNAPs exhibit remarkable variety in subunit composition and
co-factor requirements. For instance, the single subunit phage
RNAPs can perform all steps of the transcription cycle alone
(12–14), whereas bacterial RNAP core enzyme containing four
subunits needs sigma factor for initiation (15). At the other
extreme RNAP II core enzyme contains 12 subunits but needs
approximately eight additional auxiliary factors for transcription.
*To whom correspondence should be addressed. Tel: +45 3532 7762; Fax: +45 3539 6042; Email: [email protected]
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Nucleic Acids Research, 2002, Vol. 30, No. 3
MATERIALS AND METHODS
Plasmids
Figure 1. Topology of transcription. Modes of the relative rotary movements
between RNAP and DNA template during transcription elongation [see also
(4)]. (A) Transcription without rotary constraints. The newly synthesized RNA
is omitted because its fate is uncertain; it may or may not be entangled around
the template. (B) Transcription with rotary constrained RNAP and unrestrained
DNA. The nascent RNA remains free of the DNA. (C) Transcription of rotary
(torsionally) constrained DNA with rotary unrestrained RNAP. The nascent
RNA will become entangled around the template (the scenario studied in this
communication). (D) Transcription with rotary constrained RNAP and DNA.
The nascent RNA remains free of the DNA template. In this case significant
DNA supercoiling is expected to accumulate as indicated by (+) and (–). Black
and gray arrows show the direction of RNAP and DNA movement, respectively.
Black bars indicate rotary boundaries.
Moreover, each of these co-factors can contain several
subunits (16).
The organization of DNA into topological domains (17) has
been demonstrated in various organisms including E.coli (18),
Drosophila (19) and also in mamalian cells (20). A topological
domain is delimited by boundaries that prevent or limit DNA
rotation. Such boundaries could arise from attachment of DNA
to the nuclear matrix via specialized sites such as MARs and
SARs (21,22); yeast telomeric sequences (23) and the yeast
REP partition system (24) are well-established examples.
Transcription of a gene may also provide a domain boundary,
e.g. in case of coupled transcription and translation (25).
Moreover, in mammalian cells the size of a topological domain
can change in response to transcription, underlining the
dynamic character of some of these boundaries (20). Finally,
efficient transcription elongation by RNAP I in Saccharomyces
cerevisiae (26,27) and in HeLa cells (28) requires DNA
relaxing activities.
Currently, it is unclear to what extent different components
of the transcription apparatus limit RNAP and DNA rotation
during transcription elongation. In vitro data suggest that
nascent RNA chains can be sufficient to impede RNAP
rotation (9,10). In contrast, in vivo results suggest that
inhibition of RNAP rotation may involve additional restrictions
such as those occurring during transcription of genes encoding
membrane-associated proteins (25,29–32). In order to study
issues concerning RNAP rotation during transcription in a
well-defined system, we have examined transcription elongation
rates using a rotary impaired (torsionally constrained) DNA
template (the scenario depicted in Fig. 1C). For this, we have
developed a method for the immobilization of covalently
closed circular DNA (cccDNA) and used the resulting torsionally
constrained DNA as a template for transcription elongation
using free E.coli RNAP.
Plasmid DNA was prepared by standard techniques (33).
pT3T7: the peptide nucleic acid (PNA) target was cloned by
insertion of the oligonucleotides 5′-dGAAGAAGAAAACTGCA-3′/5′-dGTTTTCTTCTTCTGCA-3′ into the PstI site of
pUC19 (the target sequence is underlined). Oligonucleotides
containing the phage T3 (5′-dAATTAATTAACCCTCACTAAAGGGT-3′/5′-dAATTACCCTTTAGTGAGGGTTAATT-3′) and phage T7 (5′-dAGCTGTAATACGACTCACTATAGGG-3′/5′-dAGCTCCCTATAGTGAGTCGTATTAC3′) RNAP promoter sequences were then inserted into the
EcoRI and HindIII sites, respectively. Consequently, pT3T7 is
∆EcoRI and ∆HindIII. pT3T7(–): the control construct
pT3T7(–) is identical to pT3T7 except that it does not carry a
cognate PNA target. pTEC: the control DNA pTEC was generated
by inverse PCR using pT3T7 as a template and the oligonucleotides 5′-dCGGAATTCAATTAACCCTCACTAAAGGGTAATGATTAGGTAGGAAGAGATCTGGCCGTCGTTTTAC3′/5′-dCGGAATTCGAGCTCGGTACC-3′. pTEC is thus similar
to pT3T7 except that it carries a functional EcoRI site (and a
C-free cassette extending 22 bp from the T3 promoter +1,
which is not relevant to this study). DNA was propagated in
E.coli strain XL1-Blue under conditions found previously to
give a superhelical density (σ) approximately –0.05 (34).
PNA
PNA1021 [biotin-(eg1)3-TTJTTJTTTT-Lys-aha-Lys-aha-LysTTTTCTTCTT-Lys-NH2] and PNA1767 [biotin-eg1-Pcl(eg1)2-TTJTTJTTTT-Lys-aha-Lys-aha-Lys-TTTTCTTCTT-LysNH2] were synthesized using solid phase Boc chemistry as
described previously (35,36). The symbols used are as follows: eg1
(8-amino-2,6-dioxaoctanoic acid), Lys (lysine), aha (6-aminohexanoic acid), J [pseudoisocytosine (36)], Pcl [ortho-nitrobenzyl
type photo-cleavable linker (37)]. The PNAs were dissolved in H2O
to a concentration of 100 µM using the molar extinction coefficients
for the corresponding nucleobases [ε260 = 8.000 M–1 cm–1 (T),
7.300 M–1 cm–1 (C) and 3.000 M–1 cm–1 (J)] and stored in
aliquots at –20.
Triplex invasion
Triplex invasion was done by incubating 1 µg DNA/10 µl total
volume and 1 µM PNA in 10 mM Tris pH 7.5, 10 mM NaCl,
0.1 mM EDTA (TEN) for 1–1.5 h at 37°C. This reaction was
scaled from 1 to 40 µg of DNA in 10–400 µl volumes
depending on the purpose. Samples aimed for restriction
digestion were buffer adjusted for digestion with EcoRI/PvuII
or HaeIII. Streptavidin supershift analysis was done by adding
10 µg of streptavidin per µg of restriction digested triplex
invasion complex and incubation was continued for 30 min at
room temperature. Photo-induced cleavage of the nitrobenzyl
type linker in PNA1767 was done by irradiation of streptavidinbound triplex invasion complex using a 365 nm lamp. Triplex
invasion, streptavidin binding and photo-induced cleavage
were analyzed by 6–10% native PAGE (30:1 in acrylamide to
bisacrylamide) and ZybrGreen or ethidium bromide staining.
Formation of torsionally constrained DNA
To remove non-specifically formed triplex invasion
complexes, the samples were incubated at 65°C for 10 min in
Nucleic Acids Research, 2002, Vol. 30, No. 3
the presence of a 10-fold surplus of an oligonucleotide complementary to the PNAs used. Triplex invasion complex was then
purified by agarose gel electrophoresis. The band representing
supercoiled DNA was identified by ethidium bromide staining
(0.5 µg/ml) of a gel slice. The triplex invasion complex was
electrophoresed on to dialysis tubing and eluted into TEN. The
quality and yield of purified triplex invasion complex was
examined by OD260 measurements and gel electrophoresis.
Streptavidin-coated paramagnetic beads (streptavidin beads)
(Dynal, Roche) were prepared for binding by a single wash in
1 vol TEN. The supernatant was removed and triplex invasion
complex added to restore the original volume in TEN supplemented with 0.5 M NaCl for 1–2 h at room temperature. The
beads were kept in suspension by rotation. To remove unbound
DNA, the beads were washed extensively with high (0.6 M
NaCl, 0.12 M Tris pH 7 and 4 mM EDTA) and then low (TEN)
salt buffers.
The amount of triplex invasion complex bound to the
streptavidin beads was quantified in two ways. First, the
concentration of purified triplex invasion complex (input
material) was determined by OD260 measurement. The eluates
from the binding and washing steps were pooled and the
amount of non-bound DNA was determined. The amount of
DNA bound to the streptavidin beads was then calculated by
subtracting the non-bound material from the input material.
According to this procedure a loading of 20–30 ng/µl streptavidin
beads was obtained, corresponding to more than half of the
input material. Second, the amount of DNA bound was quantified
by stripping an aliquot of the streptavidin beads after binding
of the triplex invasion complex. This was done by incubation
at 85°C in the presence of 1% SDS for 10 min (SDS stripping).
Comparison of the two procedures showed that at least 80%
was released by SDS stripping.
Biotin competition experiments were performed by preincubating the streptavidin beads with biotin dissolved in
DMSO (1 µl 1 mM biotin per 20 µl streptavidin beads at a final
DMSO concentration of 5%) for at least 30 min prior to the
addition of triplex invasion complex.
In vitro transcription
Similar amounts of pT3T7 (0.5 µg) in the form of torsionally
constrained DNA or as triplex invasion complex were incubated in
the presence of 1 U (∼1 µg) E.coli RNAP (Boehringer Mannheim,
Epicentre Technologies), 10 µM RNA primer (5′-ACCCUG),
5 µM ATP and ∼0.4 MBq [α-32P]UTP in 100 mM KCl, 40 mM
Tris pH 8, 8 mM MgCl2, 10 mM DTT, 0.1 µg/µl BSA in a final
volume of 47 µl for 5 min at 37°C. The reactions were equilibrated
to room temperature for 1 min and elongation initiated by addition
of 3 µl ribonucleoside triphosphate (NTP)/rifampicin mixture
to make a final concentration of 0.5 mM each of ATP, CTP,
GTP and UTP and 0.25 µg/µl rifampicin. At the relevant
length of incubation, 5 µl aliquots were removed and combined
with 1 µl 0.1 M EDTA to quench elongation. The streptavidin
beads and supernatant fractions were separated using a
magnetic separator and analyzed using 8% sequencing gels
(20:1 acrylamide to bisacrylamide–7 M urea) and autoradiography.
RNA markers were made as run-off transcripts using linear
templates of known size and phage T7 or T3 RNAP.
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Figure 2. Structure of PNA and PNA–nucleic acid triplexes. (A) cccDNA, for
simplicity shown as relaxed circular DNA, bound to streptavidin beads via a
triplex invasion complex. Note that only one DNA strand has to be fixed to prevent rotation of the other. The arrow and the T indicate promoter and terminator
elements of a gene, respectively. (B) Comparison of DNA and PNA chemical
structures (B = nucleobase). (C) Hydrogen bonds formed in the T·A–T and J·G–C
PNA·DNA–PNA base triplets (J = pseudoisocytosine). (D) PNA·DNA–PNA/DNA
triplex invasion complex containing a bis-PNA equipped with a biotin moiety.
In (A) and (D) the PNA is drawn in bold.
RESULTS
Preparation of torsionally constrained DNA
Our approach to generation of torsionally constrained DNA is
outlined in Figure 2. The high specificity and stability of
triplex invasion complexes formed between homopyrimidine
PNA and DNA was exploited (Fig. 2B–D). Triplex invasion
complexes contain an internal PNA–DNA·PNA triplex in which
two PNA strands hybridize to the complementary DNA strand by
combined Watson–Crick and Hoogsteen base pairing. Consequently, the non-complementary DNA strand is displaced as a
loop (38–40). Supercoiled DNA, or in principle any DNA
molecule containing a suitable target, can be attached to
streptavidin beads using a biotin conjugated PNA as a tether.
Because the streptavidin bead is too large to rotate within the
DNA circle, the rotation of one DNA strand about the DNA
helical axis is prevented at the point of tether. The triplex invasion complex therefore delineates DNA rotary boundaries. The
bound plasmid circle is torsionally constrained and continues
to be so as long as the continuity of the strands is preserved and
the DNA remains bound to the streptavidin bead.
PNA1021 containing a biotin moiety at the N-terminus (see
Materials and Methods) was used for triplex invasion with
supercoiled plasmid DNA in which the complementary target
has been cloned. Resulting triplex invasion complexes were
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Nucleic Acids Research, 2002, Vol. 30, No. 3
gel-purified and bound to streptavidin beads via the biotin
residue. Non-bound nucleic acids were then removed by
extensive washing.
To investigate the specificity of the various interactions in
the streptavidin bead–triplex invasion complex we employed
electrophoretic mobility shift analysis (EMSA). PNA1021 was
incubated with a mixture of two plasmids containing (pTEC)
or not containing [pT3T7(–)] a cognate PNA target. The DNA
was subsequently restriction digested and analyzed (Fig. 3A).
The DNA fragment harboring the PNA target displayed
reduced mobility after incubation with PNA. The DNA
fragments that lack a target were entirely unaffected even at the
highest PNA concentration. This suggested that complete and
sequence-specific triplex invasion could be obtained.
Addition of streptavidin to pre-formed triplex invasion
complexes supershifted the band representing the PNA-bound
DNA fragment (Fig. 3A), showing that the biotin–PNA conjugate
was intact and accessible for streptavidin binding.
This analysis, however, examines only a fraction of the
DNA. Because a triplex invasion complex situated on the
template strand obstructs elongation, it was important to
analyze PNA binding along the entire length of the DNA molecule.
This was accomplished by digesting the DNA with HaeIII
thereby generating restriction fragments of suitable sizes for
EMSA (Fig. 3B).
As anticipated, binding occurred to the fragment containing
the cloned PNA target. Unexpectedly, two additional DNA
fragments changed mobility, indicating that non-target binding
also occurred. Upon heating at 65°C, the mismatched
complexes gradually dissociated. In contrast, the fully matched
triplex invasion complex was relatively unaffected by the
treatment as expected from the Tm value of >90°C for the
corresponding PNA·DNA–PNA triplex (data not shown). The
entire plasmid except for small fragments encompassing <5%
of the total template could be analyzed in this manner.
Moreover, inspection of the sequence revealed that in the
fraction of DNA that could not be analyzed by this procedure,
possible imperfect PNA targets contained three or more
mismatches. Triplex invasion at such sites should be very
inefficient (41). Thus, by using a combination of triplex invasion,
thermal dissociation of non-specific PNA·DNA–PNA/DNA
complexes and gel purification, it was possible to obtain intact
plasmids containing a single site-specifically formed triplex
invasion complex.
To ascertain that binding of the triplex invasion complex to
the streptavidin beads occurred via biotin–streptavidin interactions, a biotin competition experiment was carried out
(Fig. 3C). Incubation of the streptavidin beads with excess of
biotin inhibited the subsequent binding of the triplex invasion
complex. Thus free biotin competed efficiently for binding to
the streptavidin beads. When the biotin concentration was
reduced, binding was restored (data not shown). This supports
that the triplex invasion complex bound to the streptavidin
beads via biotin–streptavidin interactions.
One way to generate a control template for transcription
without rotational constraints, would be to first generate the
streptavidin bead–triplex invasion complex and then cleave off
half of the template for separate analysis. This approach was
taken using a PNA1021 analog, PNA1767, containing a
nitrobenzyl type photolabile linker that is cleaved upon
irradiation at 365 nm (Fig. 4) (37). To test the cleavage
Figure 3. Specificity of triplex invasion and function of the PNA–biotin conjugate.
(A) EMSA was used to monitor triplex invasion and to examine the accessibility
of the biotin–PNA conjugate for streptavidin binding. One microgram of DNA
[0.5 µg pTEC (containing a PNA target) and 0.5 µg pT3T7(–) (w/o a PNA target)]
was incubated with PNA1021 as described (Materials and Methods). Following
triplex invasion, the DNA was restriction digested with EcoRI and PvuII. Each
sample was split into two and streptavidin was added as indicated. The samples
were analyzed by 6% native PAGE and ethidium bromide staining. The PNA
concentrations used are indicated. The different species are indicated with
arrows: TIC (triplex invasion complex), SA-TIC (streptavidin–triplex invasion
complex). (B) Specificity of triplex invasion and heat denaturation of mismatched
complexes as examined by EMSA. Triplex invasion was conducted using 10 µg of
pT3T7 and PNA1021. The triplex invasion complex involving supercoiled
DNA was gel purified and buffer adjusted for restriction digestion. An aliquot
(∼1 µg) was subjected to 65°C for the indicated period of time, restriction
digested with HaeIII and analyzed as above. The fragment carrying the fully
matched target and those containing single or double mismatch binding sites
are indicated. (C) Specificity of the streptavidin bead–triplex invasion
complex interaction. Agarose gel showing the result of a biotin competition
experiment. Twenty microliters of streptavidin beads were washed and then
incubated with excess biotin in DMSO as indicated. Gel-purified triplex
invasion complex was added and the NaCl concentration adjusted to 0.5 M.
The streptavidin beads were further processed as described (Materials and
Methods).
Nucleic Acids Research, 2002, Vol. 30, No. 3
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Figure 4. Photocleavage of the o-nitrobenzyl type PNA linker. EMSA showing
photo-induced cleavage of PNA1767. Streptavidin-bound triplex invasion
complex (0.4 µg DNA/sample) containing PNA1767 was irradiated for the
indicated period of time and analyzed by PAGE as described (Materials and
Methods). The various species are indicated. The nomenclature is as in Figure 3.
[The discrete high molecular weight band appearing upon addition of streptavidin
could be due to binding of streptavidin to non-specific triplex invasion complexes
since the reactions were not heated to eliminate mismatch binding (see Fig. 3)
as was otherwise done for samples intended for transcription (Fig. 5)].
reaction, a triplex invasion complex was formed (lane 1) and
incubated further with streptavidin to establish the fully supershifted streptavidin–triplex invasion species (lane 2). This
complex was irradiated at 365 nm for various periods of time
and linker cleavage was scored by the gradual loss of the
band representing streptavidin supershift and gain of the band
representing triplex invasion complex. Half cleavage was
observed at ∼20 min. When bound to streptavidin beads,
however, much higher light intensities were required and thus
irradiation using a xenon lamp was attempted. Unfortunately,
such treatment led to DNA strand breakage thus compromising
the topology of the plasmid. DNA strand cleavage was
observed even when using an acetone filter with a cut-off of
∼315 nm. Therefore we decided to use triplex invasion
complex that had not been bound to streptavidin beads as a
control template.
Transcription using torsionally constrained DNA as a
template
Transcription elongation using a torsionally constrained
template was investigated using immobilized DNA and E.coli
RNAP. The reaction was initiated using a hexamer RNA primer
complementary to the β-lactamase promoter –5 to + 1 region (42)
and a subset of NTPs. Elongation was initiated by adding a
mixture containing the full complement of NTPs and
rifampicin to inhibit reinitiation. Aliquots were withdrawn at
the times indicated and quenched with EDTA. After separation
of the bead and supernatant fractions, the generated RNA was
analysed using sequencing gels (Fig. 5).
Most notably, the elongation rate, estimated at ∼20 nt/s
(cannot be determined exactly due to heterogeneity in
promoter escape and pausing times), was consistently as high
when using torsionally constrained DNA as a template as
compared with rotary free DNA (Fig. 5A). This indicates that the
rotary movement of RNAP was not impeded by hydrodynamic
friction on the nascent RNA or other factors, provided that the
Figure 5. Transcription of torsionally constrained DNA directed from the
β-lactamase promoter. (A) Autoradiograph showing the RNAs resulting upon
single round transcription by E.coli RNAP of the beta-lactamase gene in
torsionally constrained and rotary unrestrained conformations. The incubation
times were as follows: 0 s (lanes 1, 8 and 15), 10 s (lanes 2, 9 and 16), 30 s
(lanes 3, 10 and 17), 1 min (lanes 4, 11 and 18), 3 min (lanes 5, 12 and 19),
10 min (lanes 6, 13 and 20), 30 min (lanes 7, 14 and 21). Control experiments
in which rifampicin was added to the RNAP mix prior to the addition of DNA
template verified that only a single initiation round took place (data not
shown). The size of the RNA molecular markers is indicated. (B) Analysis of
the template before and after transcription. Aliquots (100 ng) of the indicated
template were withdrawn after incubation for 10 min in the relevant buffer and
subjected to SDS stripping (Materials and Methods) as indicated. The DNA
was analyzed on 1% agarose TAE gels, stained with ZybrGreen and further
processed using a Pharmacia Imagemaster documentation system and ImageQuant
software. Prolonging transcription to 30 min did not produce results significantly
different from those obtained after 10 min of incubation (data not shown).
Nomenclature: T (total sample), B (bead fraction), S (supernatant fraction),
Strip (SDS stripping), TB (transcription buffer), RNAP (RNA polymerase), Nc
(nicked circular), Sc (supercoiled). Other nomenclature is as in Figure 3.
supercoiled template remains torsionally constrained throughout
the experiment. To examine template nicking during the
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Nucleic Acids Research, 2002, Vol. 30, No. 3
experiment, aliquots of the reactions were removed and
analyzed by agarose gel electrophoresis (Fig. 5B). Quantification
showed that significantly <10% of the DNA bound to streptavidin
beads was nicked in any sample whether transcribed or not (cf.
lanes 3, 5 and 7). Moreover, very little DNA was released
during the transcription reaction (Fig. 5B, cf. lanes 7 and 8).
Finally, because identical amounts of torsionally constrained
and triplex invasion complex were employed (see Materials
and Methods), the RNAs produced during transcription of
DNA on beads must originate from a torsionally constrained
template.
Further examination of the autoradiograph reveals a number
of distinct RNA species. Some of these RNAs are genuine
pause products since they were converted into longer species
on further incubation. Moreover, some are termination products
because they predominantely appear in the supernatant fraction
after transcription of the torsionally constrained template.
The experiment also provides a possibility for examination
of ‘torsion-specific’ pausing sites identifiable as RNA products
that depend on template rotary constraints. While we find only
small variations between the torsionally constrained and
triplex invasion complex templates, some RNAs are more
pronounced in the former (Fig. 5A and data not shown). Further
investigation will be necessary to elucidate the significance of these
‘torsion-specific’ RNA products and to establish their dependence
on reaction conditions. We conclude that the transcription
elongation rate, at least in this system, depends very little, if at
all, on template constraints.
We also wanted to test transcription over longer distances
and for this purpose we used T3 RNAP because it is more
straightforward to construct a template without a terminator for
phage RNAPs. Our preliminary results show that RNA species
several times the length of the template could be generated
albeit at much reduced efficiency compared with uncomplexed
DNA (results not shown). However, several obstacles
hampered a detailed analysis. A relatively low signal made it
impossible to monitor elongation directly. Moreover, the PNA
interfered with elongation in a non-trivial way although
situated on the non-template strand, which should normally
allow efficient passage of phage RNAPs across the triplex
invasion complex (43).
especially when additional factors are bound to the transcript,
it should be energetically favored to rotate DNA within RNAP.
If template rotation (and translocation) is precluded, RNAP is
forced to perform all motion during elongation. For transcription of beta-lactamase (0.9 kb) under comparable ionic
strength conditions, the 5′-RNA must be carried a distance of
∼8000 Å. Furthermore, the RNA chain is probably entangled
around the DNA. In spite of this, the E.coli RNAP maximal
elongation rate was unaffected by torsional constraints. This
conclusion is certainly true for RNA sizes up to ∼0.9 kb (cf.
Fig. 5) and our preliminary data using phage RNAPs suggest
that this could also be the case for even longer RNAs (data not
shown). This conclusion is in agreement with the original
considerations of Liu and Wang (1), who calculated that nascent
transcripts would not provide sufficient hydrodynamic drag to
rotary immobilize RNAP unless bound by additional components. Before broader generalizations can be made concerning
transcription of torsionally constrained templates, however, it
needs to be addressed whether parameters such as gene and
plasmid size, and position of promoter and immobilization
elements have any effect on the elongation process.
In addition to these issues, our system opens up the possibility
of investigating the propensity of RNAP to perform elongation
using rotary immobilized enzyme (e.g. by employing His-tagged
RNAP) and template (this work) and thus to identify at which
point a topoisomerase swivel becomes a necessity for
continued elongation. It could also be possible to identify
factors that rotary constrain bacterial and eukaryotic RNAPs
using torsionally constrained templates and transcription
proficient cell lysates. In this respect such templates can be
regarded as mimics of MAR/SAR sites. Finally, this system
could be used to assemble and study well-defined transcription
complexes and auxiliary factors on supercoiled templates.
ACKNOWLEDGEMENTS
We thank Mrs Annette W. Jørgensen for Tm measurements and
Dr Ivo Krab for comments on the manuscript. The Danish
Biotechnology Programme and the Lundbeck Foundation
supported this work.
REFERENCES
DISCUSSION
Several scenarios have been proposed to explain the mutual
RNAP/DNA rotary requirements during transcription (Fig. 1).
In the present case, transcription was studied in the mode
where rotary unrestrained RNAP transcribes a torsionally
constrained DNA. Previous in vitro data suggest that nascent
RNA alone (9,10) may inhibit rotation of RNAP around a
template thus forcing the DNA to rotate instead. The argument
for this is as follows. During elongation, RNAP traverses a
distance of ∼34 Å from its initial position per 10.5 bp transcribed.
This translational motion is accompanied by a rotational path
of ∼132 Å, using an effective DNA diameter of 42 Å as determined
by Rybenkov et al. (44) for DNA at 0.1 M NaCl, 5 mM MgCl2
(versus 0.1 M KCl, 8 mM MgCl2 and 2 mM total NTP used in
our experiment). Because the RNA is extended by only ∼68 Å
per DNA turn [obtained using an RNA contour length of 6.5 Å/nt
(45)], RNAP must drag along the trailing RNA. At some point,
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