Departments of Genetics and Surgery, Center
for Genetic and Cellular Therapies, Duke University Medical Center,
Durham, North Carolina 27710
Received 14 April 1999/Returned for modification 16 May
1999/Accepted 30 June 1999
 |
INTRODUCTION |
The observation that certain RNA
enzymes can efficiently perform cleavage or splicing reactions upon RNA
substrates in the test tube (7) has engendered much
speculation about the potential utility of ribozymes for inhibiting
gene expression (5, 26, 30, 37) or for repairing mutant RNAs
in human cells (28, 36). Such therapeutic development has
been hindered by the fact that it has been difficult to directly
evaluate RNA catalysis in clinically relevant settings. Experimental
approaches in which trans-splicing group I ribozymes may be
evaluated for substrate specificity and RNA repair efficiency in
mammalian cells have recently been described (15, 16). While
a significant fraction of substrate RNAs were converted to products by
trans-splicing in this experimental system, one significant
limitation of these studies is that they did not allow one to determine
what fraction of the ribozymes was folded into a catalytically
competent conformation after expression from a cellular polymerase.
Herein we set out to begin to directly evaluate this important reaction
parameter, which we call catalytic efficiency, for group I ribozymes
inside human cells.
The group I intron from Tetrahymena thermophila has served
as a model system for studing both RNA catalysis and RNA folding (8). This intron catalyzes its own excision from precursor 23S rRNAs without the aid of proteins by a well-characterized two-step
pathway (Fig. 1) (6). In the
context of the pre-rRNA transcript, the intron folds into a complex
tertiary structure, forming an active site for phosphodiester transfer.
This structure is achieved by base-pairing interactions throughout the
intron sequence forming RNA helices, which then fold into a specific conformation to create a catalytic core within the intron. The 5'
splice site is determined by the P1 helix, which is formed by base
pairing between the internal guide sequence of the intron and the last
6 nucleotides (nt) of the 5' exon. In the first step of splicing,
P1 docks into the catalytic core, and the 5' splice site is
cleaved by an exogenous guanosine nucleophile. In the second step of
splicing, the 5' and 3' exons are ligated together, and the intron is
released (6). Since generation of functional ribosomes in
T. thermophila depends on this rRNA processing event, it is
not surprising that intron-containing precursor transcripts rapidly
self-splice to completion in their native cellular environment (4).
View larger version (11K):
[in this window]
[in a new window]
|
FIG. 1.
Self-splicing reaction of the Tetrahymena
group I intron. The exons are shown as dashed lines with capital
letters, and the intron is depicted as a plain line with lowercase
letters. The 5' exon recognition site (CUCUCU) and the
internal guide sequence (ggaggg) are indicated. The
exogenous guanosine (G) nucleophile, the terminal guanosine (g) of the
intron, and the 5' ( ) and 3' ( ) splice sites are also shown. The
junction in the ligated exon sequences is noted (CUCUCU/UAAG).
|
|
The group I intron can also self-splice from full-length and truncated
versions of Tetrahymena rRNAs in vitro. However, the rate of
self-splicing even under optimal in vitro conditions does not reach the
apparent rate achieved in Tetrahymena (1, 4). A
similar rate enhancement has been observed in Escherichia
coli when the Tetrahymena intron was expressed in the
homologous position in the E. coli rRNA gene
(40). Because group I introns are not naturally found in
E. coli, this result strongly supports the hypothesis that
species-specific cofactors are not required for efficient group I
ribozyme activity in vivo (40). Other studies with E. coli indicate that the Tetrahymena intron can
efficiently self-splice from non-rRNA transcripts expressed from the
lacZ and malE genes (23, 25, 33).
These E. coli studies suggest that the
Tetrahymena ribozyme may be an attractive molecule for therapeutic development since this catalytic RNA can function efficiently in an unnatural cellular setting and form an active catalytic center even when the ribozyme is present in a variety of
sequence contexts.
For the Tetrahymena ribozyme to become a useful therapeutic
agent, however, it must be able to function efficiently in human cells.
Two recent reports demonstrating that trans-splicing
versions of the Tetrahymena ribozyme can be employed to
repair clinically relevant mutant mRNAs in two different human cell
types are encouraging in this regard (19, 22). However, in
both of these studies the level of ribozyme-mediated splicing was
apparently quite low, with repaired products being detected only after
PCR amplification. One potential explanation for this limited activity
in human cells is that the ribozyme may not readily adopt a
catalytically competent conformation in this setting.
To begin to evaluate the potential effects of the cellular environment
as well as the importance of RNA sequence context on group I catalytic
efficiency in human cells, we chose to assess the self-splicing
activity of the Tetrahymena intron when it is embedded in
cellular transcripts. The self-splicing version of the ribozyme was
used because, in contrast to the trans-splicing version,
substrate binding would not be expected to limit ribozyme activity
(16). Thus, self-splicing represents a more direct measure
of the ribozyme's ability to form a properly folded catalytic center.
Our results confirm that self-splicing is supported from nuclear
expressed genes in human cells and that intron excision proceeds with
high fidelity. However, we also find that sequences flanking the group
I intron can significantly affect the catalytic efficiency of the
ribozyme in vivo. Moreover, we find that the intron's ability to
efficiently splice from a given transcript in vivo correlates well with
the intron's propensity to fold into an active conformation and splice
from the same transcript in vitro.
| |
MATERIALS AND METHODS |
Plasmid construction.
pNI was constructed by inserting a DNA
fragment containing the Tetrahymena group I intron from
p
GST7 (2) into the retroviral vector N2A (13).
A fragment of pGST7 containing the Tetrahymena self-splicing intron, including the T7 promoter and 274 nt of 5' exon
sequences and 29 nt of the 3' exon, was amplified by PCR and PCR
primers (upstream primer, 5'-CGGGGATCCTAATACGACTCACTATA; downstream primer,
5'-GGGACGCGTGACCCCTTTCCCGCAATTTGACAAGCTTGTGACGAGGCA). The
PCR product was digested with BamHI and MluI and
inserted between the BglII and MluI sites in the
polylinker of N2A. A catalytically inactive version of the intron was
made by deleting a BglII to NheI fragment from
pGST7 to remove 94 bp from the intron core (29). A
retroviral construct carrying the inactive intron was constructed in
parallel to provide a negative control for splicing activity (pNId).
pMI was derived from the trans-splicing ribozyme construct
pT7 L-21 (39). First, the T7 promoter was replaced with a
Moloney murine leukemia virus (MMLV) promoter, and lacZ
sequences in the 3' exon were replaced with the open reading frame for
green fluorescent protein (GFP) by standard cloning techniques. A
fragment containing the first 43 nt of the self-splicing intron and
~750 nt of 5' exon sequences was amplified via PCR from pNI (upstream
primer, 5' - CCCGAGC TCAAAAAAAGAGCCCACAACCCCTCAC TCGGGGC TCGAGCTGGATACTTCC; downstream primer, 5'-TGACCCCTTTCCCGCAATTTGAG) and
inserted between the SacI site of the MMLV promoter and the
SphI site of the intron. Again, a catalytically inactive
version of the ribozyme was inserted in this context (pMId) to serve as
a control.
To generate pNI+gfp, the sequences between the BglII and
MluI sites in pNI were replaced by sequences contained in a
DNA fragment from pMI digested with BglII and
NotI by using blunt-end cloning techniques. This sequence
"swap" restores the 3' half of the intron but introduces 3' exon
sequences from pMI, including the coding region for GFP, upstream of
the long-terminal-repeat (LTR) sequences present in the 3' exon of pNI.
To generate pMI+ltr, the sequences between the BglII and
AgeI sites in pMI were replaced by sequences contained in a
DNA fragment from pNI digested with BglII and
SmaI, which includes the 3' half of the intron and ~500 nt
of the 3' exon sequence.
Transcription of self-splicing control transcripts.
RNAs
generated by in vitro transcription were used as controls to ensure
that self-splicing was not occurring during RNA extraction and
analysis. pNI and pMI were digested with XbaI and
NotI, respectively, to serve as DNA templates for
transcription by T7 RNA polymerase. Transcription reactions were
carried out in a volume of 75 µl in reaction buffer (40 mM Tris, pH
7.5; 5 mM MgCl2; 10 mM dithiothreitol [DTT]; 4 mM
spermidine, 4 mM deoxynucleoside triphosphates) for 1 h at 37°C.
These conditions almost exclusively yield full-length precursor
transcripts with very little self-splicing product generated during
transcription. After DNase treatment and phenol-chloroform extraction,
the transcripts were purified by spin chromatography through a
Chroma-100 column (Clontech).
Transfection and RNA isolation.
Phoenix amphotrophic cells
(17) were grown in Dulbecco modified Eagle medium (DMEM)
supplemented with 10% fetal calf serum (FCS). Cells were typically
seeded in 60-mm-diameter plates at a density of 2.4 × 106 cells per plate and allowed to settle for 18 to 24 h. Cells were transfected with 5 to 10 µg of intron-containing
plasmid DNA, and cotransfected (where appropriate) with 0.2 to 0.5 µg
of pEGFP-N1 (Clontech) (a plasmid expressing GFP used to determine
transfection efficiencies) by using the calcium phosphate method of
transfection (17). The medium containing the DNA precipitate
was replaced with fresh medium 16 h posttransfection. Total RNA
was isolated from cells 42 to 48 h posttransfection. Transfection
efficiencies were monitored by flow cytometry to assess GFP expression
(when applicable). To isolate total RNA from transfected cells, the cells were trypsinized and collected by centrifugation. The cell pellet
was then solubilized in 1.5 ml of Tri-Reagent (Molecular Research
Center, Inc.) supplemented with 10 mM EDTA. In control samples, in
vitro-generated transcripts containing the intron were added to the
Tri-Reagent before the mock-transfected cells were solubilized. The
total RNA was extracted and precipitated according to the
manufacturer's protocol and was resuspended in 10 mM Tris (pH
7.5)-0.1 mM EDTA. Prior to RNase protection analysis, the RNA samples
were DNase treated for 5 min at 37°C in reaction buffer (10 mM Tris,
pH 7.5; 6.25 mM MgCl2; 50 U of DNase per ml) containing 10 mM L-argininamide to inhibit splicing.
Isolation and sequencing of spliced products.
Retroviral
supernatants were collected 48 h posttransfection from Phoenix
cells transfected with the retroviral vector-based constructs and used
to infect 2 × 105 NIH 3T3 cells. Infections were
performed with 1 ml of undiluted supernatant in 2 ml of DMEM plus 400 µg of Polybrene per ml at 37°C. After 4 h of incubation, 2 ml
of DMEM with 10% FCS was added. The virus-containing medium was
replaced 48 h after infection with selective medium containing 0.8 mg of Geneticin (Gibco-BRL) per ml. G418-resistant cells were isolated,
and genomic DNA was harvested by using a genomic DNA isolation kit
(Qiagen, Inc.). Genomic DNA was digested with SphI to
destroy unspliced intron sequences in the DNA sample before
amplification of splice junction sequences by PCR by using primers
specific for 5' and 3' exon sequences. The PCR products were digested
with BamHI and HindIII and subcloned into
pUC19. Inserts were sequenced by using the dideoxy method and a
downstream primer (5'-GTGACGAGGCATTTGGC) complementary to 3'
exon sequence located immediately downstream of the expected spliced
junction. Sequencing reaction mixtures were separated on a 10%
denaturing polyacrylamide gel and visualized by use of a PhosphorImager
(Molecular Dynamics).
RNase protection assay.
RNase protection probes were
transcribed from DNA templates that were generated by PCR.
Intron-containing plasmids were amplified by using a downstream primer
that contains the T7 promoter sequence, followed by 30 nt of
nonspecific sequences and 18 nt which are complementary to the 3' exon
of a given construct. Upstream primers were designed to anneal to
primer sites chosen 40 nt upstream of the 3' splice site in the intron
DNA sequence or 40 nt upstream of the splice junction in DNA containing
the spliced exons. PCR products were gel purified and isolated from the
gel by using Promega PCR Preps as instructed by the manufacturer.
Radiolabeled antisense probes were generated by transcribing DNA
templates for 1 h at 37°C with 40 mM Tris (pH 8); 6 mM
MgCl2; 10 mM DTT; 2 mM spermidine; 0.5 mM GTP, ATP, and
UTP; 12.5 µM CTP; 6.25 µM [
-32P]CTP; and 20 U of
T7 RNA polymerase (Roche Molecular Biochemicals). Transcription
reaction mixtures were DNase treated with 4 U of DNase for 20 min at
37°C, and products were separated on a 6% denaturing polyacrylamide
gel. Full-length probes were excised from the gel and eluted overnight
in probe elution buffer (Ambion). Eluted probe samples were analyzed by
using a scintillation counter in order to quantitate the incorporation
of radioisotope and calculate probe concentration.
RNase protection assays were performed by using an RPA II kit (Ambion)
as described by the manufacturer. Briefly, total RNA samples (5 to 10 µg) were DNase treated, added to 2 fmol of radiolabeled probe, and
allowed to hybridize overnight at 42 to 45°C. The RNAs were then
digested with a 1:100 dilution of RNase A and RNase T1
mixture (0.5 U of RNase A, 20 U of RNase T1) (Ambion) at
37°C for 30 min. The protected RNA fragments were precipitated and separated on a 6% denaturing polyacrylamide gel. Quantitation of
protected products corresponding to precursor and spliced RNAs was
performed on a PhosphorImager by using ImageQuant software (Molecular
Dynamics). Reported values have been adjusted to account for different
specific activities of the protected products due to different lengths
of the expected fragments.
In vitro splicing reactions.
Total cellular RNA samples were
thermally renatured by incubation of the RNA in H2O at
50°C for 20 min. The in vitro splicing reaction was initiated with an
equal volume of cis-splicing buffer (2× concentration of
300 mM NaCl, 100 mM Tris [pH 7.5], 20 mM MgCl2, and 4 mM
GTP) or with an equal volume of low magnesium cis-splicing
buffer (2× concentration of 300 mM NaCl, 100 mM Tris [pH 7.5], 4 mM
MgCl2, and 1 mM GTP). Reaction mixtures were incubated at
37°C for 20 min for single time point samples or for 0, 2, 5, 20, 60, and 150 min for splicing time courses. Reactions were quenched by the
addition of an equal volume of 20 mM EDTA. Splicing efficiency was
assessed by RNase protection analysis as described above. The observed
fractions of spliced products were adjusted to determine the fraction
of spliced product accumulated in vitro; any fraction of spliced
products generated in vivo was subtracted from the spliced products
observed at each time point, and that value was divided by the fraction
of precursor transcripts present in the initial sample. Data were fit
to two-phase exponential equations by using GraphPad PRISM software. To
compare the rates of splicing for the active intron populations, the
data were first normalized by dividing the fraction of in
vitro-generated spliced products calculated at each time point by the
fraction of spliced products calculated at the last time point taken.
| |
RESULTS |
The Tetrahymena intron self-splices from
transcripts expressed in human cells.
To determine if the
Tetrahymena ribozyme can mediate splicing in mammalian cells
when expressed within an RNA polymerase II (Pol II) transcript, we
introduced the Tetrahymena self-splicing intron into a
retroviral vector called NI. To generate NI, the intron was inserted in
the U3 region of the 3' LTR of the retroviral vector N2A (Fig.
2) (13). In addition, a
control vector called NId was generated that is identical to NI except
that it contains an inactive version of the ribozyme (29).
The plasmids containing NI (pNI) and NId (pNId) were transfected
separately into the Phoenix packaging cell line (17), which
is derived from human embryonic kidney 293T cells. Inside the packaging
cells, NI and NId express viral transcripts that encode neomycin
phosphotransferase and contain the Tetrahymena intron in the
3' untranslated region (UTR).
View larger version (25K):
[in this window]
[in a new window]
|
FIG. 2.
Self-splicing intron expression constructs. The
Tetrahymena self-splicing intron (black box) and flanking
exon sequences derived from pGST7 (wavy lines) are shown. The MMLV
LTR promoter (MLV) and coding sequences for neomycin phosphotransferase
(NeoR) and GFP are indicated. RNA Pol II transcription
start sites are denoted with arrows, and the location of the
polyadenylation signals [p(A)] are labeled. Insertion sites of GFP
and LTR sequences in NI+gfp and MI+ltr, respectively, are denoted by
the dashed lines.
|
|
To establish that the Tetrahymena intron self-splices in
human cells and to directly evaluate the fidelity of this reaction, we
took advantage of our experimental system and the retroviral life
cycle. The Phoenix packaging cells used in this study encapsidate retroviral vector-derived transcripts into virions, and this virus can
be used to transduce other mammalian cells. During infection, retroviral transcripts are reverse transcribed into double-stranded DNA, which is then stably integrated into the host cell's genome (Fig.
3). The in vivo reverse transcription
step in the viral life cycle allowed us to directly identify and
characterize the splicing reaction products by analyzing proviral DNA
sequences isolated from infected NIH 3T3 cells. Spliced junctions were
preferentially amplified from genomic DNA that had been digested with
SphI, which selectively cleaves unspliced intron sequences
present in the DNA sample. The amplified products were then cloned and
sequenced. The sequences of five individual spliced junctions were
determined to be exactly as anticipated (Fig. 3). A larger sample of
spliced junctions was analyzed after PCR amplification by restriction enzyme analysis. The amplified products corresponding in size to the
expected splice junction sequences were cleaved to completion by
AflII, as would be anticipated following precise excision of the intron (data not shown). Such analyses demonstrated that the Tetrahymena group I intron had indeed self-spliced from
retroviral transcripts before reverse transcription and that such
splicing had proceeded with high fidelity.
View larger version (46K):
[in this window]
[in a new window]
|
FIG. 3.
Sequence of a group I intron-derived splice junction. A
scheme for retroviral gene transfer is shown on the left. Viral RNAs
containing the Tetrahymena intron are expressed from
retroviral vectors transiently transfected into a viral packaging cell
line. As described in the text, products of group I self-splicing from
transcripts in vivo can be evaluated at the DNA level by PCR
amplification of proviral DNA sequences integrated in the genomes of
infected cells. A representative example of a sequenced splice junction
is shown on the right, along with the expected RNA sequence for a
properly spliced product. The junction between the 5' and 3' exons and
the AflII restriction site generated by this junction are
labeled.
|
|
To determine the steady-state fraction of transcripts from which the
intron has excised itself in vivo, total RNA was extracted from Phoenix
cells 2 days after transfection with pNI or pNId and subjected to RNase
protection analysis. An RNA probe complementary to 3' exon derived LTR
sequences and spanning the 3' splice site (designated 3'ss) was
hybridized to samples of total RNA to simultaneously detect precursor
and spliced transcripts (Fig.
4a).
Moreover, a second probe complementary to the 3'-exon-derived LTR
sequences and to the expected spliced junction of a self-processed
transcript (designated sj) was also generated to detect fully spliced
products (Fig. 4a). With the 3'ss probe for RNase protection analysis, RNA isolated from cells transfected with the active intron construct NI
yielded two protected RNA fragments of the expected sizes that correspond to the unspliced precursor and spliced product transcripts (Fig. 4b). The identity of the protected fragment corresponding to the
spliced product was confirmed by incubating the total RNA sample under
splicing conditions in vitro prior to RNase protection analysis. As
predicted, the shorter RNA product increased in intensity relative to
the longer protected fragment (compare lanes 7 and 8). This in vitro
splicing analysis also demonstrated that the unreacted fraction of
remaining precursor transcripts isolated from the transfected cells
were competent to undergo self-splicing even though they did not
self-splice in the cells. Detection of the RNase protection fragment
corresponding to the spliced transcript was dependent on ribozyme
activity, since only one protected RNA fragment, corresponding to the
unspliced precursor, was detected when total RNA from cells transfected
with pNId was analyzed. This precursor transcript was not converted to
spliced product even when incubated under splicing conditions in vitro
(Fig. 4b).
View larger version (31K):
[in this window]
[in a new window]
|
FIG. 4.
RNase protection analysis of intron-containing
retroviral transcripts. (a) Schematic view of RNase protection probes
for NI. The two RNase protection probes, 3'ss and sj, are indicated.
The 3' splice site probe, 3'ss, spans the junction between the 3' exon
LTR sequences and the intron in the precursor transcript. The splice
junction probe, sj, is complementary to the expected spliced junction
product RNA which lacks the intron. Both probes contain 30 nt of
nonspecific sequence at their 5' ends which is not complementary to
NI-derived transcripts. RNA generated by in vitro transcription from
the indicated T7 promoter (T7 pro) served as a control transcript as
described in the text. (b) RNase protection analysis with the 3' splice
site probe. The probe lane shows the undigested probe used in the assay
(387 nt). T7 transcripts containing the intron were added to
mock-transfected cells in control lanes. The input control transcript
(ø) was added to buffer just prior to RNase protection analysis and
served as a size marker for unspliced transcripts. RNA samples were
analyzed before ( ) and after (+) they had been incubated under in
vitro splicing conditions. Protected RNAs of 357 and 315 nt detected in
control transcripts incubated under splicing conditions correspond to
unspliced and spliced transcripts, respectively, and are marked on the
right with arrowheads. Total RNA samples isolated from cells
transfected with pNI or pNId are labeled. (c) RNase protec- tion analysis with the splice junction probe. Lanes are
labeled as described in panel b. Protected RNAs of 357 and 315 nt
detected in control transcripts incubated under splicing conditions
correspond to spliced and unspliced transcripts, respectively, and are
marked on the right with arrowheads. (d) Evaluation of RNase protection
assays. Increasing amounts of total RNA isolated from cells transfected
with pNI were analyzed as described in Materials and Methods. Protected
fragments corresponding to unspliced (U) and spliced (S) transcripts
are marked with arrowheads. A linear increase in radioactive signal is
observed over the range of RNA amounts used in these studies. Moreover,
a constant fraction of spliced NI RNAs is observed from the range of
input RNA analyzed.
|
|
An important control supports the conclusion that all of the observed
group I intron self-splicing activity detected by RNase protection
analysis occurred in the transfected cells and not during RNA
extraction or during in vitro analysis. Control RNAs were generated by
in vitro transcription that contain the same intron and 3' exon
flanking sequences as NI. These transcripts were introduced into the
cell lysis reagent added to a mock-transfected plate of cells at the
time of cell lysis. The control RNA sample was then analyzed in
parallel with the RNAs isolated from transfected cells. As demonstrated
by RNase protection analysis, the group I intron-containing control
transcripts did not self-splice during the processing and analysis of
RNA samples (Fig. 4b, lane 2), even though introns in these control
transcripts are capable of self-splicing when incubated in vitro under
appropriate conditions (Fig. 4b, lane 3).
To verify that both steps of splicing have occurred accurately, rather
than hydrolysis of the 3' exon or intron mis-splicing, the RNase
protection probe that spans the expected splice junction sequence,
probe sj, was also employed to evaluate the splicing products present
in RNAs from cells transfected with pNI or pNId. RNase protection
analysis with this probe detected a fragment of the size expected for a
transcript containing this splice junction in addition to a shorter
fragment that corresponds to unspliced precursor transcripts (Fig. 4c,
lane 6). The identities of these products were confirmed by comparison
with the protected fragments generated from in vitro-transcribed RNAs,
which had been incubated under splicing conditions (Fig. 4c, lane 4).
Again, no protected fragments corresponding to spliced products were
detected when RNA from cells transfected with the inactive ribozyme
were analyzed (Fig. 4c, lane 5) or when the unspliced in
vitro-generated control transcript containing the group I intron was
added to mock-transfected cells during RNA extraction (Fig. 4c, lane 3).
Ribozyme catalytic efficiency was measured by quantitating the relative
abundance of the two protected RNA fragments generated by RNase
protection analysis. Because spliced and unspliced transcripts are
simultaneously detected in the RNA sample by using a single probe, the
fractions of spliced transcripts from independent samples can be
quantitated and compared. To verify that such RNase protection analysis
yielded an accurate measure of spliced and unspliced transcripts over
the working range of RNA, increasing amounts of cellular RNA containing
NI transcripts (2.5 to 20 µg) were analyzed. As shown in Fig. 4d, the
expected changes in radioactive signal were observed with increasing
input RNA, while a constant fraction of spliced transcripts were detected.
The steady-state fraction of spliced products generated in cells
transfected with pNI was calculated as the average fraction detected by
RNase protection analysis of total RNA samples isolated from multiple,
independent transfections. When the 3'ss probe was used, the intron had
excised itself from 33% ± 5%, (range, 24 to 49% for seven
independent experiments) of the retroviral transcripts. Similar
splicing efficiencies were observed in these RNA samples when the sj
probe, which spans the splice junction, was employed. These results
indicate that a significant fraction of group I introns can form
catalytic centers and promote self-splicing from nuclearly expressed
transcripts in human cells. The observation that the 3'ss and sj probes
detect the same level of splicing suggests that most group I splicing
events that initiate go to completion inside cells. Similar
self-splicing efficiencies were obtained from transcripts in two murine
fibroblast cell lines, a viral packaging cell line called E86 and the
corresponding parental cell line NIH 3T3, that had been stably
transfected with pNI and pNId (19a). The fraction of introns
which had self-spliced from the NIH 3T3 cells did not significantly
differ from the fraction observed in mouse E86 cells or human Phoenix
cells, which constitutively express viral genes. Therefore, the
efficiency of group I intron splicing observed in Phoenix cells from
intron-containing transcripts (Fig. 4) is not specific to the transient
expression of self-splicing constructs or the expression of constructs
in a specific cell type.
Transcript context affects ribozyme catalytic efficiency in
vivo.
A second intron-containing construct was designed to assess
self-splicing activity when the intron is placed in a different transcript context. MI is a construct which expresses a 1.5-kb transcript encoding GFP (Fig. 2). In contrast to the intron's location
in the 3' UTR of NI, the intron is located upstream of the coding
region in pMI. Approximately 800 nt of 5' exon sequence immediately
upstream of the intron is common to both NI and MI constructs.
Downstream of the intron, only 23 nt of the 3' flanking sequence is
common to both expression cassettes. To assess self-splicing from MI
transcripts, pMI was transiently transfected into Phoenix cells, and
total RNA was isolated and analyzed by using RNase protection analysis.
As shown in Fig. 5, this analysis
demonstrated that the intron does not self-splice from MI transcripts
at a detectable level inside the transfected cells. However, the
intron-containing RNAs isolated from the cells are able to self-splice
when incubated under standard in vitro reaction conditions (Fig. 5b,
compare lanes 1 and 2). These results suggest that the intron is
largely inactive in vivo within the MI transcript context but that this inactivity is not simply due to a defect in the intron sequence. Dideoxy sequencing of the group I intron within pMI confirmed that no
mutations were introduced into the intron during plasmid construction
(data not shown).
View larger version (47K):
[in this window]
[in a new window]
|
FIG. 5.
RNase protection analysis of MI transcripts. (a)
Schematic view of an RNase protection probe for MI. The RPA probe spans
the 3' splice junction between the 3' exon GFP sequences and the intron
in the precursor transcript. The probe contains 30 nt of nonspecific
sequence at the 5' end which is not complementary to MI-derived
transcripts. RNA generated by in vitro transcription from the indicated
T7 promoter (T7 pro) served as a control transcript to monitor for
potential self-splicing during RNA extraction and analysis. (b) RNase
protection analysis of transcripts from MI-transfected cells. The probe
lane shows the undigested probe used in this assay (322 nt). T7
transcripts containing the intron were added to mock-transfected cells
in control lanes. RNA samples were analyzed before () and after (+)
they had been incubated under in vitro splicing conditions. Protected
RNAs of 292 and 250 nt detected in control transcripts incubated under
splicing conditions correspond to unspliced and spliced transcripts,
respectively, and are marked on the left with arrows. Total RNA samples
from cells transfected with pMI are labeled.
|
|
We attempted to determine what differences between the active (NI) and
inactive (MI) transcripts accounted for the striking difference in the
ability of the group I intron to self-splice in the cellular
environment. One possible explanation is that the local sequence
context of the intron influences the proper secondary and tertiary
folding of the ribozyme in vivo (24, 27). Since the 3' exon
flanking sequences are significantly more divergent than the 5' exon
sequences in NI and MI transcripts (Fig. 2), we sought to determine
whether 3' exon sequences could influence self-splicing efficiency. Two
new expression cassettes were constructed from plasmids pNI and pMI to
test this hypothesis. In the first, pNI+gfp, the 3' exon GFP sequence
flanking the intron in pMI was inserted downstream of the group I
intron present in pNI. In the second, pMI+ltr, the 3' flanking LTR
sequence from pNI was inserted downstream of the intron present in pMI
(see Fig. 2). Thus, pNI and pMI+ltr were constructed to contain
identical 3' exon sequences immediately flanking the intron, which is
located either in the context of the 3' UTR (NI) or the 5' UTR (MI+ltr) of the expressed transcript. Similarly, pNI+gfp and pMI plasmids contain the identical flanking 3' exon sequences in two different contexts. The effects of these 3' exon sequences on group I intron splicing were then assessed. Each construct was transfected into Phoenix cells, and splicing efficiency was analyzed by RNase protection analysis by using the appropriate 3' splice site probes and total RNA
isolated from transfected cells. The resulting percentage of spliced
products detected for the each of these transcripts is shown in Fig.
6. The presence of GFP sequences
downstream of the intron in NI transcripts reduced in vivo
self-splicing efficiency from ~35% splicing to ~5% splicing. In
contrast, LTR sequences present downstream of the intron in MI
transcripts enhanced self-splicing from below detection (<2%) in MI
transcripts to ~15% in MI+ltr transcripts. These results indicate
that sequence context, and in particular the 3' exon sequences, may
significantly affect the efficiency with which the group I intron
splices in human cells.
View larger version (14K):
[in this window]
[in a new window]
|
FIG. 6.
Effects of 3' exon sequence on splicing efficiency in
vivo. A schematic comparison of various 3' exon sequences from the four
constructs tested is shown on the left. The constructs containing these
sequences are indicated, along with the corresponding splicing
efficiency observed in vivo.
|
|
A difference in transcript accumulation does not account for the
observed differences in catalytic efficiency of related
constructs.
It is possible that different mRNA stabilities could
indirectly affect the fraction of self-spliced transcripts detected
from the various constructs analyzed in vivo if self-splicing rates were low relative to the half-life of the intron-containing
transcripts. To address this issue, we compared the relative levels of
accumulation of two transcripts from which group I introns self-spliced
to significantly different extents. Equal molar amounts of pNI and pNI+gfp (Fig. 2) were cotransfected into Phoenix cells, and total RNA
was isolated in order to assess the relative abundance of transcripts
expressed from the two retroviral vectors by RNase protection analysis.
Since the two transcription units within pNI and pNI+gfp are the same,
with the exception of the 800-bp insertion of GFP coding sequences in
the 3' UTR of pNI+gfp, we assumed that the rates of transcription
associated with the retroviral promoters would be equivalent for the
two constructs. A single RNase protection probe, which spans the
GFP-LTR sequences in pNI+gfp, was employed to detect transcripts
expressed from both constructs (Fig. 7a).
With this probe design, both unspliced and spliced transcripts
contribute to the RNase protection signal detected for each construct.
Thus, if the low abundance of spliced transcripts detected in NI+gfp
(Fig. 6) resulted from a significant decrease in RNA transcript
stability, we would anticipate that a decrease in NI+gfp transcripts
relative to NI transcripts would also be evident. Results from RNase
protection analysis of total RNA samples from singly or cotransfected
cells with this probe strategy are shown in Fig. 7b. Quantitation of
the relative abundance of the two transcripts present in the RNA
isolated from cotransfected cells demonstrated that nearly equal
amounts of NI and NI+gfp transcripts were reproducibly found in the
cells. In contrast, RNase protection analysis with appropriate
3'-splice-site probes to assess splicing efficiencies in these RNA
samples confirmed that there is a vast difference in the levles of
accumulation of spliced products within cells cotransfected with NI and
NI+gfp (Fig. 7c). These observations suggest that a difference in RNA half-lives does not significantly contribute to the differences in
self-splicing efficiency observed between NI and NI+gfp transcripts in
vivo.
View larger version (23K):
[in this window]
[in a new window]
|
FIG. 7.
Accumulation of transcripts from related constructs. (a)
RNase protection analysis probe that distinguishes between the 3' exon
sequences following the intron in pNI and pNI+gfp and detects both
unspliced and spliced transcripts. (b) RNase protection analysis of
total RNA extracted from cells transfected with pNI and pNI+gfp. The
probe lane shows the full-length undigested probe used in the assay
(330 nt). Lanes indicate total RNA samples isolated from singly and
cotransfected cells (performed in triplicate and labeled samples 1 to
3). Protected RNA fragments of 260 and 300 nt correspond to NI and
NI+gfp transcripts, respectively, and are indicated by the arrows. The
band marked with the asterisk is present in all cells transfected with
retroviral vectors derived from N2A. The signal corresponds in size to
5' LTR sequences which may result from read-through transcription of
transfected plasmids. (c) Self-splicing efficiencies of NI and NI+gfp
transcripts in RNA samples isolated from cotransfected cells. RNase
protection analysis was performed by using appropriate 3' splice site
probes. Percent splicing of NI and NI+gfp transcripts is reported for
RNA samples isolated from cotransfected cells and analyzed for relative
accumulation of retroviral transcripts in panel b.
|
|
In vivo catalytic efficiency correlates with the ability of the
intron to fold properly and self-splice from a given sequence
context.
In vitro activity assays have frequently been used to
assess the rate and extent of proper folding of the group I ribozyme from Tetrahymena (9, 11, 12, 38). Since
intron-containing transcripts generated from the four self-splicing
constructs described in this study could readily undergo self-splicing
under in vitro conditions regardless of their in vivo efficiency, we
examined more carefully the in vitro splicing reactions mediated by
these introns from their Pol II transcript contexts. The reactions were performed with RNA samples isolated from cells transfected with intron-containing constructs, and the accumulation of spliced products
in these reactions was assayed by RNase protection analysis. As
expected, the data fit best to a two-phase exponential equation (11). These phases are interpreted as an initial population of properly folded introns which react quickly after initiation of the
reaction and as a second slow-reacting population of introns which must
undergo refolding to an active structure before catalysis can occur
(11). The extent of in vitro splicing for introns embedded
in the various cellular transcripts described is shown in Fig.
8a. In Fig. 8b, the data were normalized
in each reaction to the final fraction of spliced products in order to
compare the levels of progress of active intron populations. Similar
curves suggest that introns which fold into an active conformation
during the course of the in vitro reaction also self-splice at
essentially the same rate, regardless of which transcript context the
intron is found (Fig. 8b). However, the extent of group I intron
self-splicing varies when it is present in different Pol II transcripts
(Fig. 8a). This presumably reflects a difference in the ability of the intron sequence to fold properly within a given cellular transcript under the in vitro conditions used. Interestingly, the transcript context with the greatest self-splicing efficiency observed in vivo,
NI, was also the context from which the largest fraction of introns
self-spliced in vitro. Moreover, the extents of the reactions observed
in vitro for all four transcript contexts correlate exactly with the
trend of relative efficiencies observed for group I self-splicing from
these transcripts inside cells (Fig. 8a).
View larger version (20K):
[in this window]
[in a new window]
|
FIG. 8.
In vitro self-splicing reactions of various cellular
transcripts. Total RNA samples were preincubated and self-spliced in
vitro as described in Materials and Methods. The splicing reactions
were stopped at the time points indicated, and the reaction products
were subsequently analyzed by RNase protection analysis. The fraction
of spliced products that had accumulated in vitro at each time point
was quantitated and calculated as described in Materials and Methods.
Data were fit to a two-phase exponential equation. Closed symbols are
used for constructs containing LTR sequences downstream of the intron.
Open symbols are used for constructs containing GFP sequences
downstream of the intron. Circles correspond to retroviral (NI)-based
vectors, while squares correspond to MI-based vectors. (a) Progress of
in vitro self-splicing reactions. (b) Data normalized for each reaction
in panel a such that the fraction spliced (norm.) is equal to the
fraction spliced at each time point divided by the maximum fraction
spliced during the reaction.
|
|
| |
DISCUSSION |
Our results demonstrate that group I introns are able to
self-splice from transcripts expressed in the nuclei of mammalian cells. To establish a system which allows for isolation and
characterization of bona fide in vivo splicing products without
worrying that splicing was occurring during in vitro analysis, we
inserted the intron into a retroviral vector. By taking advantage of
the retroviral life cycle to recover spliced products that were reverse
transcribed into DNA inside cells, we were able to verify that splicing
does proceed in vivo and that the intron precisely ligates its flanking exon sequences together (Fig. 3). Moreover, by using quantitative RNase
protection analysis, we determined that up to 50% of the introns have
self-spliced with high fidelity from steady-state levels of the
retroviral transcript, NI, in vivo (Fig. 4). These results suggest that
a significant fraction of group I ribozymes, expressed within cellular
transcripts, could potentially react in human cells. In addition, we
compared the catalytic efficiencies of group I self-splicing from a
variety of transcript contexts. Our inability to detect group I
self-splicing from one of these transcripts, MI, indicates that
transcript context can have dramatic effects on the in vivo catalytic
efficiency of group I ribozymes (Fig. 5). The observation that
self-splicing efficiencies from cellular transcripts increased or
decreased when new sequences were inserted downstream of the ribozyme
indicates that group I ribozyme activity may be enhanced or inhibited
by the presence of certain flanking RNA sequences in vivo (Fig. 6). In
the case of NI and NI+gfp transcripts, which are transcribed from
closely related constructs yet differ greatly in catalytic efficiency, we show that differences in RNA stabilities do not contribute significantly to the markedly reduced level of spliced NI+gfp transcripts detected at steady state (Fig. 7). Finally, we obtained intriguing results when we compared the in vitro splicing efficiencies of the various intron-containing transcripts. A reduced extent of
splicing in vitro, which is believed to indicate that a greater fraction of the introns are misfolding, correlated with reduced catalytic efficiency in cells (Fig. 6 and 8). Thus, although we cannot
provide direct evidence for misfolding of ribozymes in vivo, our
results suggest that the ability of the ribozyme to fold into an active
conformation in vitro may significantly influence the catalytic
efficiency of the ribozyme containing transcript inside cells.
The steady-state level of in vivo spliced products depends upon the
relationship between the rate of self-splicing inside cells and the
half-life of the expressed transcripts. If intron-containing transcripts have a relatively short half-life, then the intron may not
have sufficient opportunity to self-splice before the transcript is
degraded. Moreover, a transcript from which the intron has self-excised
may become less stable than the precursor transcript and thus
under-represent the true fraction of transcribed RNAs which undergo
splicing. Alternatively, the intron's propensity to fold into an
active versus inactive conformation in vivo, within the context of the
expressed transcript, may significantly contribute to differences in
self-splicing efficiency. It has been demonstrated in vitro that
misfolded ribozymes may result from unfavorable interactions within the
intron and with flanking RNA sequences, as discussed below. In addition
to such intrinsic folding properties, RNA folding inside cells may be
affected by RNA binding proteins which may favorably or unfavorably
influence the intron's ability to adopt a catalytically competent
conformation. It has been demonstrated in vitro that RNA binding
proteins, such as ribosomal peptides, viral nucleocapsid proteins, and
hnRNPs, may act as RNA chaperones to increase the rate of ribozyme
reactions, presumably by enhancing ribozyme folding to achieve the
catalytic structure (3, 10, 14, 32). Similarly, RNA
chaperone activity may influence splicing rates inside cells. However,
it is unclear if the intracellular concentrations of such RNA binding
proteins would positively or negatively impact on the ability of
ribozymes to correctly fold within transcripts and if those influences
would be transcript context dependent. Therefore, a variety of factors
can potentially influence the efficiency with which a group I intron
self-splices from a Pol II transcript inside mammalian cells.
In our studies, we demonstrate that sequence context can significantly
influence the catalytic efficiency of the group I intron self-splicing
inside cells even when two similar intron containing transcripts, the
NI and NI+gfp RNAs, accumulate to similar levels and thus appear to
have similar half-lives inside cells. One simple interpretation of
these results is that in the NI context the intron tends to fold into
an active conformation more often than in the NI+gfp context. The
observation that a greater fraction of the ribozymes in NI transcripts
adopt active conformations in vitro compared to that in NI+gfp
transcripts strongly supports this theory.
The observation that exonic sequences can influence self-splicing
activity is well established for the natural sequence context of the
Tetrahymena intron. Inclusion of different lengths of exon sequences in precursor rRNA transcripts can affect the rate of intron
self-splicing in vitro (12, 34). In these studies, relative
splicing rates are more closely related to the predicted structure of
the exon sequences than to the length of the exons. In very short rRNA
precursor transcripts, exon sequences that sequester the 5' exon
recognition site from the P1 helix into an alternative helix, called
P(1), dramatically reduce the rate of group I cis-splicing
(35). The stabilization or destabilization of P(1) by
point mutations (35) or additional rRNA exon sequences (34) alters the equilibrium between active and inactive
intron conformations. Exon sequences have also been shown to limit the activities of catalytic introns of the group I td intron in
bacteria (27) and a model group II intron precursor
transcript in vitro (20). Local folding of sequences
flanking the Tetrahymena intron which enhance or inhibit
formation of the catalytically competent intron structure may similarly
contribute to the efficiency of self-splicing from the various
transcript contexts that we observe in mammalian cells.
Interestingly, such detrimental effects resulting from flanking exon
sequences that reduce group I ribozyme catalytic efficiency in E. coli have not been apparent in the few studies that have been
performed. The Tetrahymena intron self-splices from the
homologous position in E. coli rRNA with a rate rivaling
that of Tetrahymena rRNA processing (40). In
addition, a number of studies have demonstrated that the
Tetrahymena intron self-splices readily from sequences
unrelated to the natural rRNA context in bacteria (23, 25,
33). The intron self-splices from the lacZ transcripts efficiently enough to fully complement -galactosidase activity in
E. coli (23, 33). The efficiency of splicing from
the lacZ transcript was not directly quantitated in these
studies, but RNA blot analysis demonstrated that the free intron was
found to be much more abundant than precursor lacZ RNAs in
E. coli. In a more recent study, the intron was expressed
downstream of the malE gene (25). From this
position the intron splices extremely efficiently, as evidenced by the
fact that unspliced introns were detected in only 0.2% of the
steady-state population of malE transcripts in the bacteria.
Efficient self-splicing from these unnatural sequence contexts in
E. coli demonstrates that rRNA exon sequences are not a
prerequisite for efficient in vivo catalysis. In our mammalian cell
studies, we observed that the intron self-excised from different
transcript contexts with a range of catalytic efficiencies, including
undetectable levels of self-splicing from certain transcripts. While
these results cannot be compared directly to those of existing studies
of E. coli since it is unknown how relative transcript stabilities affect the different steady-state levels of spliced transcripts, it is somewhat surprising that transcript context has not
been shown to significantly impact upon self-splicing efficiency in the
few studies undertaken with nonribosomal RNA transcripts in the
prokaryotic cellular environment.
As established for the Tetrahymena ribozyme in vitro,
kinetically trapped inactive conformations of the folded intron is one plausible mechanism by which self-splicing activity may be limited in
human cells. A number of ribozyme derivatives containing point mutations have recently been made that modulate ribozyme folding properties in vitro (21, 31). These include mutations which stabilize the P3 helix of the intron core to increase the fraction of
active intron conformers after in vitro transcription (21). Other mutations have been identified that allow the ribozyme to resolve
inactive conformations rapidly in order to adopt the active ribozyme
conformation more quickly in vitro (31). Evaluation of these
and other derivatives of the Tetrahymena intron in mammalian cells by using the experimental system presented here should greatly enhance our understanding of the factors which influence the ribozymes propensity to fold into a catalytically competent conformation in a
therapeutically relevant setting.
Recently, a few studies have reported that trans-splicing
versions of the Tetrahymena ribozyme can be employed to
revise targeted RNAs in E. coli (18, 29) and in
mammalian cells (15, 16, 19, 22). Moreover, two of these
reports describe the use of such trans-splicing to repair
clinically relevant transcripts associated with myotonic dystrophy
(22) and sickle cell disease (19) in mammalian
fibroblasts and in erythrocyte precursors from sickle cell patients,
respectively. Although these two reports are extremely encouraging with
regard to the potential utility of RNA repair, since they demonstrate
that ribozymes can revise endogenous transcripts, the levels of
trans-splicing and RNA repair in these studies were
apparently very low. Unfortunately, information about the catalytic
efficiency of the ribozymes utilized in these experiments is very
difficult to obtain. Nevertheless, for trans-splicing ribozymes to become useful in the clinic, the efficiency with which
ribozymes can repair mutant RNAs will need to be assessed and will
likely have to be increased. One parameter of the
trans-splicing reaction that may significantly impact on a
group I ribozyme's ability to repair target RNAs in vivo is the
propensity of the ribozyme to fold into a catalytically competent
conformation when it is expressed as part of a longer transcript inside
human cells. Unfortunately, we are currently unable to predict if group
I ribozymes will tend to form a catalytically active or inactive
structure when the intron is embedded in flanking exon sequences.
However, the strategy that we employed herein to assess the catalytic
efficiency of self-splicing in vivo should prove to be valuable for
evaluating and potentially enhancing the catalytic potential of a
variety of group I ribozyme-containing transcripts for therapeutic
applications in mammalian cells.
We thank P. Zarrinkar, C. Rusconi, and N. Lan for critical
reading of the manuscript and J. Jones, P. Zarrinkar, and C. Rusconi for helpful discussions. Plasmid pGST7 was generously provided by T. Cech. G. Nolan graciously provided the amphotrophic Phoenix cell line.
This material is based upon work supported under a National Science
Foundation Graduate Research Fellowship (M.B.L.) and by NIH grant GM
53525 (B.A.S.).
| 1.
|
Bass, B. L., and T. R. Cech.
1984.
Specific interaction between the self-splicing RNA of Tetrahymena and its guanosine substrate: implications for biological catalysis by RNA.
Nature
308:820-826[Medline].
|
| 2.
|
Been, M. D., and T. R. Cech.
1986.
One binding site determines sequence specificity of Tetrahymena pre-rRNA self-splicing, trans-splicing and RNA enzyme activity.
Cell
47:207-216[Medline].
|
| 3.
|
Bertrand, E. L., and J. J. Rossi.
1994.
Facilitation of hammerhead ribozyme catalysis by the nucleocapsid protein of HIV-1 and the heterogeneous nuclear ribonucleoprotein A1.
EMBO J.
13:2904-2912[Medline].
|
| 4.
|
Brehm, S. L., and T. R. Cech.
1983.
Fate of an intervening sequence ribonucleic acid: excision and cyclization of the Tetrahymena ribosomal ribonucleic acid intervening sequence in vivo.
Biochemistry
22:2390-2397[Medline].
|
| 5.
|
Cech, T. R.
1988.
Ribozymes and their medical implications.
JAMA
260:3030-3034[Abstract].
|
| 6.
|
Cech, T. R.
1990.
Self-splicing of group I introns.
Annu. Rev. Biochem.
59:543-568[Medline].
|
| 7.
|
Cech, T. R.
1993.
Structure and mechanism of the large catalytic RNAs: group I and group II introns and ribonuclease P, p. 239-269.
In
R. F. Gesteland, and J. F. Atkins (ed.), The RNA world. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
|
| 8.
|
Cech, T. R., and D. Herschlag.
1996.
Group I ribozymes: substrate recognition, catalytic strategies, and comparative mechanistic analysis, p. 1-17.
In
F. Eckstein, and D. M. J. Lilley (ed.), Catalytic RNA. Springer-Verlag, Berlin, Germany.
|
| 9.
|
Celander, D. W., and T. R. Cech.
1991.
Visualizing the higher order folding of a catalytic RNA molecule.
Science
251:401-407[Abstract/Free Full Text].
|
| 10.
|
Coetzee, T.,
D. Herschlag, and M. Belfort.
1994.
Escherichia coli proteins, including ribosomal protein S12, facilitate in vitro splicing of phage T4 introns by acting as RNA chaperones.
Genes Dev.
8:1575-1588[Abstract/Free Full Text].
|
| 11.
|
Emerick, V. L.,
J. Pan, and S. A. Woodson.
1996.
Analysis of rate-determining conformational changes during self-splicing of the Tetrahymena intron.
Biochemistry
35:13469-13477[Medline].
|
| 12.
|
Emerick, V. L., and S. A. Woodson.
1993.
Self-splicing of the Tetrahymena pre-rRNA is decreased by misfolding during transcription.
Biochemistry
32:14062-14067[Medline].
|
| 13.
|
Hantzopoulos, P. A.,
B. A. Sullenger,
G. Ungers, and E. Gilboa.
1989.
Improved gene expression upon transfer of the adenosine deaminase minigene outside the transcriptional unit of a retroviral vector.
Proc. Natl. Acad. Sci. USA
86:3519-3523[Abstract/Free Full Text].
|
| 14.
|
Herschlag, D.
1995.
RNA chaperones and the RNA folding problem.
J. Biol. Chem.
270:20871-20874[Free Full Text].
|
| 15.
|
Jones, J. T.,
S.-W. Lee, and B. A. Sullenger.
1996.
Tagging ribozyme reaction sites to follow trans-splicing in mammalian cells.
Nat. Med.
2:643-648[Medline].
|
| 16.
|
Jones, J. T., and B. A. Sullenger.
1997.
Evaluating and enhancing ribozyme reaction efficiency in mammalian cells.
Nat. Biotechnol.
15:902-905[Medline].
|
| 17.
|
Kinoshita, S.,
B. K. Chen,
H. Kaneshima, and G. P. Nolan.
1998.
Host control of HIV-1 parasitism in T cells by the nuclear factor of activated T cells.
Cell
95:595-604[Medline].
|
| 18.
|
Kohler, U.,
B. G. Ayre,
H. M. Goodman, and J. Haseloff.
1999.
Trans-splicing ribozymes for targeted gene delivery.
J. Mol. Biol.
285:1935-1950[Medline].
|
| 19.
|
Lan, N.,
R. P. Howrey,
S.-W. Lee,
C. A. Smith, and B. A. Sullenger.
1998.
Ribozyme-mediated repair of sickle -globin mRNAs in erythrocyte precursors.
Science
280:1593-1596[Abstract/Free Full Text].
|
| 19a.
| Long, M. B., and B. A. Sullenger.
Unpublished results.
|
| 20.
|
Nolte, A.,
G. Chanfreau, and A. Jacquier.
1998.
Influence of substrate structure on in vitro ribozyme activity of a group II intron.
RNA
4:694-708[Abstract].
|
| 21.
|
Pan, J., and S. A. Woodson.
1998.
Folding intermediates of a self-splicing RNA: mispairing of the catalytic core.
J. Mol. Biol.
280:597-609[Medline].
|
| 22.
|
Phylactou, L. A.,
C. Darrah, and M. J. A. Wood.
1998.
Ribozyme-mediated trans-splicing of a trinucleotide repeat.
Nat. Genet.
18:378-381[Medline].
|
| 23.
|
Price, J. V., and T. R. Cech.
1985.
Coupling of Tetrahymena ribosomal RNA splicing to -galactosidase expression in Escherichia coli.
Science
228:719-722[Abstract/Free Full Text].
|
| 24.
|
Roman, J.,
M. N. Rubin, and S. A. Woodson.
1999.
Sequence specificity of in vivo reverse splicing of the Tetrahymena group I intron.
RNA
5:1-13[Abstract].
|
| 25.
|
Roman, J., and S. A. Woodson.
1998.
Integration of the Tetrahymena group I intron into bacterial rRNA by reverse splicing in vivo.
Proc. Natl. Acad. Sci. USA
95:2134-2139[Abstract/Free Full Text].
|
| 26.
|
Rossi, J. J.
1992.
Ribozymes.
Curr. Opin. Biotechnol.
3:3-7[Medline].
|
| 27.
|
Semrad, K., and R. Schroeder.
1998.
A ribosomal function is necessary for efficient splicing of the T4 phage thymidylate synthase intron in vivo.
Genes Dev.
12:1327-1337[Abstract/Free Full Text].
|
| 28.
|
Sullenger, B. A.
1995.
Revising messages traveling along the cellular information superhighway.
Chem. Biol.
2:249-253[Medline].
|
| 29.
|
Sullenger, B. A., and T. R. Cech.
1994.
Ribozyme-mediated repair of defective mRNA by targeted trans-splicing.
Nature
371:619-622[Medline].
|
| 30.
|
Sullenger, B. A., and T. R. Cech.
1993.
Tethering ribozymes to a retroviral packaging signal for destruction of viral RNA.
Science
262:1566-1569[Abstract/Free Full Text].
|
| 31.
|
Treiber, D. K.,
M. S. Rook,
P. P. Zarrinkar, and J. R. Williamson.
1998.
Kinetic intermediates trapped by native interactions in RNA folding.
Science
279:1943-1946[Abstract/Free Full Text].
|
| 32.
|
Tsuchihashi, Z.,
M. Khosla, and D. Herschlag.
1993.
Protein enhancement of hammerhead ribozyme catalysis.
Science
262:99-102[Abstract/Free Full Text].
|
| 33.
|
Waring, R. B.,
J. A. Ray,
S. W. Edwards,
C. Scazzocchio, and R. W. Davies.
1985.
The Tetrahymena rRNA intron self-splices in E. coli: in vivo evidence for the importance of key base-paired regions of RNA for RNA enzyme function.
Cell
40:371-380[Medline].
|
| 34.
|
Woodson, S. A.
1992.
Exon sequences distant from the splice junction are required for efficient self-splicing of the Tetrahymena IVS.
Nucleic Acids Res.
20:4027-4032[Abstract/Free Full Text].
|
| 35.
|
Woodson, S. A., and T. R. Cech.
1991.
Alternative secondary structures in the 5' exon affect both forward and reverse self-splicing of the Tetrahymena intervening sequence RNA.
Biochemistry
30:2042-2050[Medline].
|
| 36.
|
Woolf, T. M.
1998.
Therapeutic repair of mutated nucleic acid sequences.
Nat. Biotechnol.
16:341-344[Medline].
|
| 37.
|
Yu, M.,
E. Poeschla, and F. Wong-Staal.
1994.
Progress towards gene therapy for HIV infection.
Gene Ther.
1:13-26[Medline].
|
| 38.
|
Zarrinkar, P. P., and J. R. Williamson.
1994.
Kinetic intermediates in RNA folding.
Science
265:918-924[Abstract/Free Full Text].
|
| 39.
|
Zaug, A. J.,
C. A. Grosshans, and T. R. Cech.
1988.
Sequence-specific endoribonuclease activity of the Tetrahymena ribozyme: enhanced cleavage of certain oligonucleotide substrates that form mismatched ribozyme-substrate complexes.
Biochemistry
27:8924-8931[Medline].
|
| 40.
|
Zhang, F.,
E. S. Ramsay, and S. A. Woodson.
1995.
In vivo facilitation of Tetrahymena group I intron splicing in Escherichia coli pre-ribosomal RNA.
RNA
1:284-292[Abstract].
|
![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
Top
Abstract
Introduction
