From the Département de biochimie, Faculté de médecine, Université de Sherbrooke, Québec, J1H 5N4, Canada
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ABSTRACT |
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The specificity of ribozyme
cleavage was investigated using a trans-acting antigenomic
ribozyme. Under single turnover conditions, the wild
type ribozyme cleaved the 11-mer ribonucleotide substrate with a rate
constant of 0.34 min
1, an apparent Km
of 17.9 nM and an apparent second-order rate constant of
1.89 × 107 min
1
M
1. The substrate specificity of the
ribozyme was thoroughly investigated using a collection
of substrates that varied in either the length or the nucleotide
sequence of their P1 stems. We observed that not only is the base
pairing of the substrate and the ribozyme important to cleavage
activity, but also both the identity and the combination of the
nucleotide sequence in the substrates are essential for cleavage
activity. We show that the nucleotides in the middle of the P1 stem are
essential for substrate binding and subsequent steps in the cleavage
pathway. The introduction of any mismatches at these positions resulted
in a complete lack of cleavage by the wild type ribozyme. Our findings
suggest that factors more complex than simple base pairing
interactions, such as tertiary structure interactions, could play an
important role in the substrate specificity of
ribozyme
cleavage.
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INTRODUCTION |
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ribozymes derived from the genome of hepatitis
virus (HDV)1
are metalloenzymes. Like other catalytically active ribozymes, namely
hammerhead and hairpin ribozymes, the
ribozymes cleave a
phosphodiester bond of their RNA substrates and give rise to reaction
products containing a 5'-hydroxyl and a 2',3'-cyclic phosphate termini.
Two forms of
ribozymes, namely genomic and antigenomic,
were derived and referred to by the polarity of the HDV genome from
which the ribozyme was generated. Both
ribozyme forms
exhibit self-cleavage activity, and it has been suggested that they are
involved in the process of viral replication (1). This type of activity
has been described as cis-acting
ribozymes (2).
Like other ribozymes, ribozymes have a potential
application in gene therapy in which an engineered ribozyme is directed to inhibit gene expression by targeting a specific mRNA molecule. It has been demonstrated that a very low concentration (<0.1
mM) of Ca2+ and Mg2+ is required
for
ribozyme cleavage (3).
ribozymes have a unique
characteristic in their substrate binding, namely that only the
3'-portion of the substrate is required for binding to the ribozyme. A
short stretch of nucleotides (7 nt) located on the substrate is
required for cleavage. Although one might suspect the specificity of
ribozyme cleavages due to their short recognition site,
we view this characteristic of the
ribozyme as an
advantage for the future development of a therapeutic means of
controlling, for example, a viral infection.
Since little is known about the kinetic properties of ribozymes, study of the trans-acting system will enable us
to answer some basic questions on both the structure required and the
kinetic properties, including the substrate specificity, of
ribozymes. Depending on the predicted secondary
structures used, various trans-acting
ribozyme systems were generated by separating the RNA molecule into
ribozyme and substrate molecules at various positions (4-6). Here, we
generate a trans-acting
ribozyme, based on
the pseudo knot-like structure proposed by Perrotta and Been (2), by
separating the single-stranded region located at the junction between
the P1 and P2 stems (Fig. 1). Although, several investigations have
been performed to address the questions related to the substrate
specificity of
ribozymes in both the cis- and
trans-acting forms (2, 5-12), most, if not all, experiments were carried out by randomly changing the base pairing combinations or
by introducing mismatches which interfere with the Watson-Crick base
pairing between the substrate and the ribozyme in the P1 stem (Fig. 1).
It was demonstrated that cleavage activity was not destroyed by the
interchanging of one to four nucleotide pairs between the substrate and
the
ribozyme (2, 8, 11, 12). One or two nucleotide
mismatches at any position of the P1 stem, except positions 5 and 11 (numbering according to Fig. 1), completely destroyed the activity (2,
5-12). Although these are composite results from various versions of
ribozymes, these findings could be interpreted as
indicating that the positions located at both extremities of the base
paired stem formed by the substrate and the ribozyme were more likely
to tolerate a mismatch, resulting in distortion of the P1 stem, than
the internal positions. There is no information on how each nucleotide
of the substrate affects the cleavage activity and its kinetics since
most investigations were carried out at only one or two positions at a
time, and the findings generally reported in a plus/minus manner
(e.g. cut or uncut). Therefore, the substrate specificity of
ribozyme could not be deduced from previous reports. To
determine how substrate sequences affect
ribozyme
cleavage activity, we performed kinetic studies using a collection of
short oligonucleotide substrates (11 nt) with a trans-acting
ribozyme. In this report, we demonstrate that each
nucleotide of the P1 stem contributes differently to the cleavage
activity. We compare the observed cleavage rate constants for cleavable
substrates and the equilibrium dissociation constants for the
uncleavable substrates with those of the wild type substrate. We
present evidence that strongly suggests that the nucleotides located in
the center of the P1 stem formed between substrate and ribozyme (Fig.
1, positions 7 and 8) are important not only for substrate recognition
but probably also for subsequent steps, for example a conformation
change yielding a transition complex.
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MATERIALS AND METHODS |
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Plasmids Carrying Ribozymes
The antigenomic ribozyme sequence of the hepatitis
virus described by Makino et al. (13) was
used to generate a trans-acting
ribozyme with
some modifications as shown in Fig. 1. Briefly, the construction was
performed as follows. Two pairs of complementary and overlapping
oligonucleotides, representing the entire length of the ribozyme (57 nt), were synthesized and subjected to an annealing process prior to
cloning into pUC19. The annealed oligonucleotides were ligated to
HindIII and SmaI co-digested pUC19 to give rise to a plasmid harboring the
ribozyme (referred to as
p
RzP1.1). A mutant ribozyme (
RzP1.2) was then constructed by
modifying the substrate recognition site of p
RzP1.1 by ligation of
an oligonucleotide containing the altered sequence flanked by
restriction endonuclease sites to RsrII/SphI
predigested p
RzP1.1. The sequences of engineered ribozymes were
confirmed by DNA sequencing. Plasmids containing wild type and mutant
ribozymes were then prepared using Qiagen tip-100 (Qiagen Inc.),
digested with SmaI, purified by phenol and chloroform
extraction, and precipitated for further use as templates for in
vitro transcription reactions.
RNA Synthesis
Ribozyme-- In vitro transcription reactions contained 5 µg of linearized recombinant plasmid DNA as template, 27 units RNAGuard® RNase inhibitor (Amersham Pharmacia Biotech), 4 mM of each ribonucleotide (Amersham Pharmacia Biotech), 80 mM HEPES-KOH, pH 7.5, 24 mM MgCl2, 2 mM spermidine, 40 mM dithiothreitol, 0.01 unit of pyrophosphatase (Boehringer Mannheim) and 25 µg of purified T7 RNA polymerase in a final volume of 50 µl, and were incubated at 37 °C for 4 h.
Substrates-- Deoxyoligonucleotides (500 pmol) containing the substrate and T7 promoter sequence were denatured by heating at 95 °C for 5 min in a 20-µl mixture containing 10 mM Tris-HCl, pH 7.5, 10 mM MgCl2, 50 mM KCl2, and allowed to cool slowly to 37 °C. The in vitro transcription reactions were carried out using the resulting partial duplex formed as template under the same conditions as described for the production of the ribozyme.
After incubation, the reaction mixtures were fractionated by denaturing 20% polyacrylamide gel electrophoresis (19:1 ratio of acrylamide to bisacrylamide) containing 45 mM Tris borate, pH 7.5, 7 M urea, and 1 mM EDTA. The reaction products were visualized by UV shadowing. The bands corresponding to the correct sizes of either ribozymes or substrates were cut out, and the transcripts eluted overnight at 4 °C in a solution containing 0.1% SDS and 0.5 M ammonium acetate. The transcripts were then precipitated by the addition of 0.1 volume of 3 M sodium acetate, pH 5.2, and 2.2 volumes of ethanol. Transcript yield was determined by spectrophotometry.End-labeling of RNA with [-32P]ATP
Purified transcripts (10 pmol) were dephosphorylated in a
20-µl reaction mixture containing 200 mM Tris-HCl, pH
8.0, 10 units of RNAGuard®, and 0.2 units of calf
intestine alkaline phosphatase (Amersham Pharmacia Biotech). The
mixture was incubated at 37 °C for 30 min and then extracted twice
with a same volume of phenol:chloroform (1:1). Dephosphorylated
transcripts (1 pmol) were end-labeled in a mixture containing 1.6 pmol
[-32P]ATP, 10 mM, Tris-HCl, pH 7.5, 10 mM MgCl2, 50 mM KCl, and 3 units of
T4 polynucleotide kinase (Amersham Pharmacia Biotech) at 37 °C for
30 min. Excess [
-32P]ATP was removed by applying the
reaction mixture onto a spin column packed with a G-50 Sephadex gel
matrix (Amersham Pharmacia Biotech). The concentration of labeled
transcripts was adjusted to 0.01 pmol/ml by the addition of water.
Cleavage Reactions
To initiate a cleavage reaction, we tested different procedures and chose the method that yielded the highest cleavage rate constant and the maximum cleavage product as described by Fauzi et al. (14). Various concentrations of ribozymes were mixed with trace amounts of substrate (final concentration <1 nM) in a 18-µl reaction mixture containing 50 mM Tris-HCl, pH 7.5, and subjected to denaturation by heating at 95 °C for 2 min. The mixtures were quickly placed on ice for 2 min and equilibrated to 37 °C for 5 min prior to the initiation of the reaction. Unless stated otherwise, cleavage was initiated by the addition of MgCl2 to 10 mM final concentration. The cleavage reactions were incubated at 37 °C, and followed for 3.5 h or until the end point of cleavage was reached. The reaction mixtures were periodically sampled (2-3 µl), and these samples were quenched by the addition of 5 µl of stop solution containing 95% formamide, 10 mM EDTA, 0.05% bromphenol blue, and 0.05% xylene cyanol. The resulting samples were analyzed by a 20% polyacrylamide gel electrophoresis as described above. Both the substrate (11 nt) and the reaction product (4 nt) bands were detected using a Molecular Dynamics radioanalytic scanner after exposition of the gels to a phosphorimaging screen.
Kinetic Analysis
Measurement of Pseudo First-order Rate Constant
(kcat, Km and
kcat/Km)--
Kinetic analyses were performed
under single turnover conditions as described by Hertel et
al. (15) with some modifications. Briefly, trace amounts of
end-labeled substrate (<1 nM) were cleaved by various
ribozyme concentrations (5-500 nM). The fraction cleaved was determined, and the rate of cleavage (kobs)
obtained from fitting the data to the equation At = A(1
e
kt) where
At is the percentage of cleavage at time
t, A
is the maximum percent
cleavage (or the end point of cleavage), and k is the rate
constant (kobs). Each rate constant was
calculated from at least two measurements. The values of
kobs obtained were then plotted as a function of
ribozyme concentrations for determination of the other kinetic
parameters: kcat, Km and
kcat/Km. Values obtained from
independent experiments varied less than 15%. The requirement for
Mg2+ by both ribozymes was studied by incubating the
reaction mixtures with various concentrations of MgCl2
(1-500 mM) in the presence of an excess of ribozyme (500 nM) over substrate (<1 nM). The concentrations
of Mg2+ at the half-maximal velocity were determined for
both ribozymes.
Determination of Equilibrium Dissociation Constants (Kd)-- For mismatched substrates that could not be cleaved by the ribozyme, the equilibrium dissociation constants were determined using a slight modification of the method described by Fedor and Uhlenbeck (16). Eleven different ribozyme concentrations, ranging from 5 to 600 nM, were individually mixed with trace amounts of end-labeled substrates (<1 nM) in a 9-µl solution containing 50 mM Tris-HCl, pH 7.5, heated at 95 °C for 2 min and cooled to 37 °C for 5 min prior to the addition of MgCl2 to a final concentration of 10 mM, in a manner similar to that of a regular cleavage reaction. The samples were incubated at 37 °C for 1.5 h, at which time 2 µl of sample loading solution (50% glycerol, 0.025% of each bromphenol blue and xylene cyanol) was added, and the resulting mixtures were electrophoresed through a nondenaturing polyacrylamide gel (20% acrylamide with a 19:1 ratio of acrylamide to bisacrylamide, 45 mM Tris borate buffer, pH 7.5 and 10 mM MgCl2). Polyacrylamide gels were prerun at 20 W for 1 h prior to sample loading, and the migration was carried out at 15 W for 4.5 h at room temperature. Quantification of bound and free substrates was performed following an exposure of the gels to a phosphorimaging screen as described earlier.
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RESULTS |
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The trans-acting ribozymes used in this
report were derived from the antigenomic
ribozyme of HDV
(13). Some features of the antigenomic
ribozyme were
modified to improve its structural stability and to aid in transcript
production. Based on a pseudo knot-like structure described by Perrotta
and Been (2), Fig. 1 shows the structure
of the
ribozymes used with some modifications: (i) the
single-stranded region between substrate and ribozyme (region J1/2) was
eliminated to separate the substrate molecule from the ribozyme; (ii)
the substrate contains only 11 nt and produces 7- and 4-nt cleavage
products, and the GGG at the 5'-end was added to increase the yield
during in vitro transcription (17); (iii) three G-C base
pairs were introduced in the P2 region to improve both the structural
stability and transcript production; and (iv) the P4 stem was shortened
to the minimum length reported to result in an active ribozyme (18).
Prior to performing a cleavage reaction, native gel electrophoresis was
used to test for the possible presence of aggregates or multimer forms
of the transcripts. Various concentrations of ribozyme, ranging from 5 nM to 2 µM, were mixed with trace amounts of
end-labeled ribozyme (less than 0.5 nM) and fractionated
under nondenaturing conditions as described under "Materials and
Methods." We detected the presence of a slow migrating species of
ribozyme in the mixture containing 2 µM ribozyme (data
not shown). The quantification of the slow migrating band showed that
the band amounted to approximately 2% of the total radioactive
material. However, a single band was detected at the concentrations
used for kinetic analysis and under single turnover conditions (5-600
nM). Similar experiments were performed for each substrate.
There was no substrate multimer detected at the concentrations used
(data not shown). The equimolar mixture of end-labeled substrate and
ribozyme was also fractionated under nondenaturing conditions, and it
resulted a single band of ribozyme and substrate complex similar to
those observed for the Kd measurement shown in Fig.
4.
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Cleavage Kinetics of Constructed Antigenomic Ribozymes
Two forms of trans-acting ribozymes
(
RzP1.1 and
RzP1.2) were used with their corresponding substrates
(11 nt) for the kinetic studies.
RzP1.2 differs from
RzP1.1 in
that
RzP1.2 has two nucleotides, at positions 22 and 24 of
RzP1.1, interchanged (Fig. 1,
5'-CCCAGCU-3'). Time course experiments for
cleavage reactions catalyzed by both
RzP1.1 and
RzP1.2 were
monitored by the appearance of the 4 nt cleavage product. An example of a time course experiment for a cleavage reaction catalyzed by
RzP1.1
is shown in Fig. 2, panel A.
In this particular experiment, 100 nM of
RzP1.1 was
incubated with 1 nM end-labeled substrate, SP1.1. The newly
formed product and the remaining substrate bands at each time point
were quantified, and the percentage of cleavage was plotted as a
function of time (Fig. 2, panel B).
RzP1.1 cleaved approximately 60% of the substrate within 10 min. The data were fitted
to a single exponential equation as described under "Materials and
Methods" so as to obtain the observed rate constant
(kobs = 0.21 min
1). We attempted
to fit the data as biphasic reactions as described for the hairpin (19)
and the hammerhead (20) ribozymes. We observed that the standard
deviation (
2) of data fitted to a double-exponential
equation was higher (
2 = 0.01203) than that fitted to a
single exponential equation (
2 = 0.000203). Although we
could not exclude or dismiss completely the possibility that more than
one conformation of the active ribozyme could be formed, the data were
treated as if the reactions were monophasic in their kinetics for
comparison purposes.
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Similar experiments were performed using trace amounts of substrate
(<1 nM) and various ribozyme concentrations to measure kobs at each ribozyme concentration. The values
of kobs of both RzP1.1 and
RzP1.2
increased with an increase in ribozyme concentration up to
approximately 200 nM (Fig. 3,
panel A). The concentration of ribozyme at which the
reaction velocity reached half-maximal (apparent Km,
Km') is 17.9 ± 5.6 nM for
RzP1.1 and 16.7 ± 6.4 nM for
RzP1.2. Under the
reaction conditions used, in which the increase in ribozyme
concentration has no significant effect on the rate of cleavage, the
cleavage rate (kobs) is therefore represented by
the catalytic rate constant (kcat). The cleavage rate constants are 0.34 min
1 for
RzP1.1 and 0.13 min
1 for
RzP1.2. Apparent second-order rate constants
(kcat/Km') were calculated to
be 1.89 × 107 min
1
M
1 for
RzP1.1 and 0.81 × 107 min
1 M
1 for
RzP1.2 (Table I).
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Since we observed that the kcat of RzP1.2 is
about 3 times less than that of
RzP1.1, whereas the
Km' is similar, we investigated whether an increased
amount of Mg2+ in the cleavage reaction would affect the
kcat of
RzP1.2. Under single turnover
conditions, in which the ribozyme and substrate concentrations were
kept at 500 and 1 nm, respectively, we found that both ribozymes
cleave their complementary substrates at Mg2+
concentrations as low as 1 mM, which is the estimated
physiological concentration of Mg2+ (21). At this
concentration, the kobs obtained were 0.11 ± 0.01 and 0.04 ± 0.01 min
1, for
RzP1.1 and
RzP1.2, respectively (Fig. 3, panel B). A maximum kobs for
RzP1.2 was observed when the
concentration of Mg2+ was 10 mM. Higher
concentrations of Mg2+ did not increase either the
kobs or the extent of cleavage for both
ribozymes. We did not observe a decrease in the cleavage rate when
higher concentrations of Mg2+ were used (e.g.
500 mM). The requirement for magnesium at half-maximal velocity (KMg) was 2 mM for both
RzP1.1 and
RzP1.2.
Substrate Specificity
To compare the specificity of the ribozyme with
various substrates,
RzP1.1 was used under single turnover conditions
as described above. The cleavage reactions were performed with a trace
amount of each substrate (<1 nM) and 500 nM
RzP1.1. Under these conditions, the observed rates reflect the rates
of cleavage without interference from either product dissociation or
inhibition. For each substrate both the observed cleavage rate
constants (kobs) and the extent of cleavage were
calculated and compared with those of the wild type substrate, as shown
in Table II.
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Shorter Substrates--
Three shorter substrates containing 10, 9, and 8 nt were tested individually and compared with the 11-nt substrate
(SP1.1) in which 7 nt base paired with RzP1.1. The 10-, 9-, and 8-nt substrates contain 6, 5, and 4 nt regions complementary to
RzP1.1, respectively. We observed that the 10-nt substrate was cleaved with a
kobs of 0.02 ± 0.01 min
1 and
a maximal cleavage of 28.8% (Table II). We could not detect the
cleavage product formed when the 9- and 8-nt substrates were used, even
after a 3.5-h incubation time. The cleavage reactions were also carried
out in the presence of 100 mM Mg2+ instead of
the 10 mM concentration used in a regular cleavage reaction. We observed no improvement in the values of the
kobs and the extent of cleavage for the 10-nt
substrate and still detected no cleavage for both the 9- and 8-nt
substrates.
Mismatched Substrates--
We have generated a collection of
substrates in which single mismatches were individually introduced into
the P1 region of the substrate and then used in the cleavage reactions
(Table II). Mutation at position 5 resulted in at least a 9-fold
decrease in kobs as compared with that of SP1.1
(0.34 min1). However, for SG5A, in which A was
substituted for G at position 5 of SP1.1, the extent of cleavage was
only reduced by half. When this nucleotide was changed to cytosine, the
cleavage was reduced almost to nil (ca. 1.7%).
RzP1.1
cleaved approximately 4% of the SG6A and SG6U substrates, in which A
or U were substituted for G at position 6. The alteration of either
position 7 or 8, located in the middle of the P1 stem, yielded
uncleavable substrates (SG7A, SG7U, SU8C, SU8G). The
kobs was also drastically decreased when the C
at position 9 was altered to A or U. The extent of cleavage was reduced
to approximately 50%, when SC9U was used. The SG10U substrate, in
which U was substituted for G at position 10, gave a similar result to
SC9A. Finally,
RzP1.1 cleaved the substrate SG11U almost as well as
SP1.1, although the kobs was considerably slower
(0.01 min
1). The relative activity of each single
mismatched substrate was calculated to obtain an apparent free energy
of transition-state stabilization,
G
(22, 23). We found that the values of
G
range between
0.96 to
2.25 kcal
1 mol
1.
This apparent difference in activation energy was also observed when
substrates of leadzyme were altered and used in a cleavage assay
(22).
Equilibrium Dissociation Constant (Kd)
The four substrates containing a single mismatch either at
position 7 or 8, which were not cleaved by RzP1.1, were used to determine an equilibrium dissociation constant (Kd). Trace amounts of end-labeled substrates (SG7A, SG7U, SU8C, or SU8G)
were individually incubated with various concentrations of
RzP1.1
for the gel shift analysis as described under "Materials and
Methods." To ensure that the dissociation equilibrium was reached, we
incubated the reaction mixtures at various intervals. We found that the
equilibrium was reached within 5 min, and that a longer incubation of
28 h did not affect the measurement of Kd.
Since SP1.1 can be cleaved under native gel electrophoresis conditions,
we therefore used its analog which has a deoxyribose at position 4 (SdC4) to obtain the estimated Kd of the wild type
substrate. This analog could not be cleaved by
RzP1.1 under the
conditions used (2), and has been shown to be a competitive inhibitor
of
RzP1.1 cleavage.2 An
example of a gel shift analysis carried out for the analog is shown in
Fig. 4. In this particular analysis,
trace amounts of SdC4 (<1 nM) were incubated with 11 concentrations of
RzP1.1 ranging from 5 to 600 nM. An
autoradiogram of the resulting gel obtained by a Molecular Dynamics
radioanalytic scanner is shown in Fig. 4, panel A. The bands
of the bound SdC4 and the free SdC4 at each
RzP1.1 concentration
were quantified, and the percentage of the bound SdC4 was plotted as
shown in Fig. 4, panel B. The experimental data were fitted
to a simple binding equation as described under "Materials and
Methods" to obtain Kd = 31.9 ± 2.7 nM. Similar experiments were performed for SG7A, SG7U,
SU8C, and SU8G. The substrates in which A or U were substituted for G
at position 7 were observed to have a lower affinity for
RzP1.1 than
those of the substrates in which the U at position 8 is altered. The
higher Kd values obtained for SG7A (320 ± 20 nM) and SG7U (220 ± 60 nM), as compared
with those of the analog (31.9 ± 2.7 nM), SU8C
(36.1 ± 2.5 nM), and SU8G (71.5 ± 3.2 nM) are summarized in Table
III. We observed that the single mismatch
introduced at position 7 disturbed the equilibrium of substrate-ribozyme complex formation to a greater extent than the
mutation at position 8. The values of Kd were used in the determination of the free energy of substrate binding (Gibbs energy change,
GE·S). The mismatch at
position 7 interfered with the stabilization of the substrate-ribozyme
complex, resulting in
GE·S between
0.43 and
1.4 kcal
1 mol
1.
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DISCUSSION |
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ribozymes derived from the genome of HDV are of interest in
the development of a gene regulation system in which the designed ribozymes would down-regulate the expression of a target gene. The
facts that
ribozymes are derived from HDV and that this pathogen naturally replicates in animal systems, suggest that this
catalytic RNA could be used to control gene expression in human cells.
Like other ribozymes, the designed ribozyme should specifically cleave
its target substrates while leaving other cellular RNA molecules
intact. We designed a trans-acting
ribozyme harboring a recognition sequence similar to the HDV antigenomic
self-cleaving motif so as to have a minimal system for
the study of the specificity of the base pairing interaction between
the
ribozyme and its substrate.
Although a number of trans-acting ribozymes
have been generated, they appear to have variable cleavage rate
constants. The discrepancy of cis-acting
ribozyme activities has been reviewed, and it was suggested that the
variation of the cleavage activity, at least for cis-acting
forms, may result from the nonribozyme flanking sequences used by each
investigator (24). Our trans-acting
ribozyme,
RzP1.1, exhibited an activity with a cleavage rate of 0.34 min
1, or a t1/2 of 2 min, under
pseudo first-order conditions. These data are in good agreement with
the observed rate constant (0.35 min
1) of a
cis-acting
ribozyme derived from antigenomic
HDV RNA (7). We found that the extent of cleavage is approximately 60%, regardless of the concentration of ribozyme used, suggesting possibilities that (i) a fraction of the substrate was bound to an
inactive form of the
ribozyme; (ii) substrate was bound
to Cs of the 3' of the ribozyme, instead of to the P1 region of the ribozyme, causing a misfold or a nonactive substrate-ribozyme complex;
or (iii) a portion of the ribozyme might adopt another conformation
following substrate binding. Based on the latter hypothesis, the
alternative form of ribozyme-substrate complex could undergo cleavage
at a very low rate. We first investigated whether or not the presence
of the alternative form could be a result of an infidelity of the T7
RNA polymerase transcription. Two batches of purified T7 RNA polymerase
were tested using various amounts of enzyme and incubation times (data
not shown). We found that the transcripts produced by both batches of
purified T7 RNA polymerase at the different incubating times exhibited
a similar cleavage pattern and extent, suggesting that it is the nature of ribozyme transcripts to adopt an alternative form in the reaction mixtures, as previously reported for the hairpin ribozyme (19). A
possible occurrence of misplaced or misfold substrate-substrate complex
was dismissed since there is no evidence of other formed complexes
detected under nondenaturing gel electrophoresis and also by RNase
mapping.3 Finally, the
possible occurrence of a slow cleaving form of
ribozyme
was assessed following cleavage reactions. We attempted to fit the
experimental data using a multiphasic kinetic equation. Since we could
not clearly describe the kinetics of our trans-acting
ribozyme as biphasic or multiphasic reactions, we
measured initial rates of cleavage for comparative purposes.
To summarize the cleavage reactions catalyzed by RzP1.1 and
RzP1.2, free energy diagrams of the reaction coordinates were constructed (Fig. 5). The diagrams relate
the two states in the cleavage reactions using kinetic parameters
obtained under single turnover conditions.
RzP1.1 and
RzP1.2
differ in that they have two base pairs in the middle of the P1 stem
interchanged. As expected, the free energies of substrate binding are
virtually identical (
11 kcal
1 mol
1). The
base pair interchange in
RzP1.2 increased the value of
G
by approximately 0.5 kcal
1
mol
1. It is interesting to note that the free energy of
the transition state was affected by the changes in the base pairing of
the P1 stem. Although several kinetic parameters were greatly different from those reported here, similar findings were previously reported when two nucleotides (positions 7 and 8) of the substrate were interchanged and complemented by the
ribozyme (12).
Since the kinetics of
cleavage reactions appear to be
affected by the particular combination of base pairs, it is very likely
that in addition to P1 base pairing a tertiary interaction might also participate in substrate recognition. In this scenario the
substrate-ribozyme complex would undergo a conformational transition,
following formation of P1 stem, which involves tertiary interaction(s).
These interactions might result in the positioning of the scissile bond
in the catalytic center, a key step in the reaction pathway.
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The substrate specificity of the ribozyme was studied
using
RzP1.1. First, we found that the
ribozyme can
cleave a substrate having a minimum of 6 nucleotides adjacent to the
cleavage site. This result is in an agreement with those previously
reported for both the cis-acting form (5) and the
trans-systems (11) that a minimum of 6 base pairing is
required for cleavage. The kobs of the 10-nt
substrate is at least 10 times slower than that of SP1.1 (Table II). We
also used shorter substrates generated by alkali hydrolysis as
described by Perrotta and Been (11) to verify the cleavage reactions
catalyzed by the two similar trans-acting
ribozymes. Due to the slow cleavage rate, the detection of the
disappearance of shorter substrates in the mixtures could not
accurately be measured (data not shown).
Second, to estimate the contribution of base pairing interaction of the
P1 stem to the cleavage reaction, a collection of single mismatched
substrates was generated by introducing point mutations into the
substrate sequence. Although there are a number of reports on the base
pairing requirement of the P1 region (8, 10-12, 14), no extensive
investigation has been performed on each individual nucleotide of
either cis- or trans-acting ribozymes. The determination of ribozyme specificity against various
substrates was first attempted by comparing the apparent second-order
rate constant (kcat/Km') of
each substrate to that of wild type substrate. We found that the
ribozyme cleaved single mismatched substrates very slowly and gave a
low percent cleavage (maximum of 2-20%) within the reaction time
studied (3.5 h). As a consequence, the measurement of the apparent
second-order rate constants as a function of ribozyme concentration
yielded values with a high margin of error. We thus reported the
cleavage activity of the ribozyme against various single mismatched
substrates in terms of extent of cleavage and
kobs, which at a high ribozyme concentration reflects the kcat of the cleavage reaction. In
all cases, we observed the decrease in cleavage extent, which we
suspected to be due mainly to the poor binding between the substrate
and the ribozyme. The wobble base pair (G-U) at the cleavage site is
required to maintain a high level of cleavage (10, 11). Mismatches at this position, which create either an A-U or a C-U pairing, decreased the cleavage activity in a manner analogous to that reported in another
version of trans-acting
ribozymes (10). It is
interesting to note that the extent of cleavage decreases
proportionally to the mismatches introduced into the 3' and 5'
positions of the middle of the P1 stem. The simultaneous alteration of
two nucleotides in the middle of the P1 stem was reported to give rise
to an uncleavable substrate in both the cis- and
trans-acting systems (11). However, in both cases the
activity could be restored by the generation of a complementary
ribozyme or a substrate.
The calculated free energy of transition-state stabilization
(G
) for each substrate listed in Table
II varies between
0.9 and
2.25 kcal
1
mol
1. Each position of base pairing between the substrate
and the ribozyme appears to affect the reaction pathway differently, at least with regard to transition state complex formation. If we assume
that mismatched substrates yield the same level of
GE·S, various end points of cleavage for
mismatched substrates could be resolved depending upon the height of
the energy barrier level to be overcome in the transition state. To
address these questions precisely, more experiments on the equilibrium
binding constant and the internal equilibrium of the reactions are
required. We have determined the calculated Kd of P1
duplex formation using the equation described by Serra and Turner (25)
to be 28.5 nM. By using an analog, we have shown that the
Kd of the wild type substrate to its ribozyme is
31.9 nM. It is very interesting to note that the mismatch
introduced at position U8 of the substrate has little effect on
substrate binding affinity. However, the change completely eliminated
cleavage activity. The mismatch introduced at position G7 of the
substrate affected both the binding and chemical steps since it not
only lowered the binding affinity of the substrate for the ribozyme,
but also destroyed the cleavage activity. These findings suggest that
some base pairs of the P1 stem have dual roles, participating in the
substrate binding and subsequent steps leading a chemical cleavage, as
was observed for the base pair interactions between the hammerhead ribozyme and its substrates (26). To address these findings more
precisely some preliminary experiments have been carried out using the
metal-ion induced cleavage method to study the tertiary structure of
ribozyme 3. The data obtained to date suggests that
positions U8 and G7 are likely involved in the formation of an
essential metal-ion binding site. The mismatches introduced at either
of the two positions destroyed the formation of this metal-ion binding
site, a process which has been found to be highly associated with
cleavage activity.
We present here evidence that aside from the base pairing between the
substrate and the ribozyme, tertiary interactions, especially ones
involving the P1 stem, appear to dictate the reaction pathway of
ribozyme. To fully comprehend how the cleavage reactions are governed, the elucidation of these tertiary interactions is essential.
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ACKNOWLEDGEMENTS |
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We thank Dr. Stephane Mercure for the
pRzP1.1 construct.
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FOOTNOTES |
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* This work was supported in part by a grant from the Medical Research Council (MRC) of Canada (to J. P. P.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Recipient of a postdoctoral fellowship from Natural Sciences and
Engineering Research Council (NSERC) of Canada.
§ Medical Research Council scholar. To whom correspondence should be addressed. Tel.: 819-564-5310; Fax: 819-564-5340; E-mail: jp.perre{at}courrier.usherb.ca.
1
The abbreviations used are: HDV, hepatitis
virus; nt, nucleotide(s).
2 S. Mercure and J. P. Perreault, unpublished data.
3 D. Lafontaine and J. P. Perreault, unpublished data.
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REFERENCES |
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