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Molecular and Cellular Biology, September 1998, p. 5404-5413, Vol. 18, No. 9
0270-7306/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
The Exon Splicing Silencer in Human
Immunodeficiency Virus Type 1 Tat Exon 3 Is Bipartite and Acts
Early in Spliceosome Assembly
Zhi-hai
Si,
Dan
Rauch, and
C. Martin
Stoltzfus*
Department of Microbiology, University of
Iowa, Iowa City, Iowa 52242
Received 10 February 1998/Returned for modification 24 March
1998/Accepted 8 June 1998
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ABSTRACT |
Inefficient splicing of human immunodeficiency virus type 1 (HIV-1)
RNA is necessary to preserve unspliced and singly spliced viral RNAs
for transport to the cytoplasm by the Rev-dependent pathway. Signals
within the HIV-1 genome that control the rate of splicing include weak
3' splice sites, exon splicing enhancers (ESE), and exon splicing
silencers (ESS). We have previously shown that an ESS present within
tat exon 2 (ESS2) and a suboptimal 3' splice site together
act to inhibit splicing at the 3' splice site flanking tat
exon 2. This occurs at an early step in spliceosome assembly. Splicing
at the 3' splice site flanking tat exon 3 is regulated by a
bipartite element composed of an ESE and an ESS (ESS3). Here we show
that ESS3 is composed of two smaller elements (AGAUCC and
UUAG) that can inhibit splicing independently. We also show that ESS3
is more active in the context of a heterologous suboptimal splice site
than of an optimal 3' splice site. ESS3 inhibits splicing by blocking
the formation of a functional spliceosome at an early step, since A
complexes are not detected in the presence of ESS3. Competitor RNAs
containing either ESS2 or ESS3 relieve inhibition of splicing of
substrates containing ESS3 or ESS2. This suggests that a common
cellular factor(s) may be required for the inhibition of
tat mRNA splicing mediated by ESS2 and ESS3.
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INTRODUCTION |
Human immunodeficiency virus type 1 (HIV-1) is a complex retrovirus whose RNA splicing is dependent on the
host cell splicing machinery (12, 50). HIV-1 pre-mRNA
contains five 5' splice sites and nine 3' splice sites which are used
to generate more than 30 different mRNA species by alternative
splicing. In addition, a pool of unspliced RNA must be maintained to
serve as genomic RNA and as mRNA for the gag and
pol gene products (18, 21, 32, 34, 35, 38). To
achieve the balance between spliced and unspliced RNAs, splicing of
HIV-1 RNA is inefficient, a feature which is shared with all other
retroviruses (4, 7, 11, 20, 22, 45-47, 52). In contrast to
the major HIV-1 5' splice sites, which are strong, the 3' splice sites
are weak (33). These are characterized by the presence of
nonconsensus polypyrimidine tracts and branch point sites other than
the eukaryotic consensus adenosine residue (2, 13, 17, 33,
43). In addition, exon sequences downstream of the 3' splice
sites enhance or inhibit splicing at the 3' splice sites (2, 3,
39, 42, 51). Furthermore, the virus-encoded protein Rev plays an
essential role in facilitating the transport of unspliced and singly
spliced mRNAs, thus preventing the viral RNA from splicing to
completion (19, 37; for a review, see reference
12).
HIV-1 tat mRNA is formed by the splicing of two coding exons
(exon 2 and exon 3) and one or more upstream noncoding exons. Exon 3 is
also joined to rev exon 2 and is therefore referred to as
tat/rev exon 3 (34). The 3' splice sites flanking
exon 2 and exon 3 (3' splice sites 3 and 7, respectively [Fig.
1]) contain suboptimal polypyrimidine
tracts, and the 3' splice site flanking exon 3 has been reported to
contain a nonconsensus branch point site (17, 33, 39, 43).
In addition, we and others have demonstrated that tat/rev
exon 3 splicing is regulated by an exon splicing enhancer (ESE) and a
juxtaposed exon splicing silencer (ESS) within this exon (ESS3) (Fig.
1) (3, 42). An ESS is also present within tat
exon 2 (ESS2), which has been localized to the RNA sequence
CUAGACUAGA in the region of vpr encoding the C
terminus and in the region of tat encoding the N terminus.
It acts at an early step in the splicing pathway to selectively inhibit
splicing at the upstream 3' splice site flanking this tat
exon (3). Addition of competitor RNA containing ESS2 to
splicing reaction mixtures with HIV-1 RNA substrates containing ESS2
causes an increase in splicing at the upstream 3' splice site (3'
splice site 3). Based on these data, we hypothesized that the
inhibition by ESS2 is mediated by a negative cellular factor(s) which
binds to the ESS (3). In this study, we have delineated the
ESS3 element in tat/rev exon 3 and have investigated the
mechanism by which it inhibits splicing. The data suggest that the
inhibition by ESS2 and ESS3 involves binding of a common factor(s).
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FIG. 1.
(A) Schematic diagram of HIV-1 splice sites and
structure of tat mRNA. ESS2 and ESS3 are ESSs located within
tat exon 2 and tat/rev exon 3, respectively. 5'
splice sites are indicated above the line by arrows; 3' splice sites
are denoted below the line as solid lines. "Cap" indicates the RNA
initiation site, and "pA" represents the poly(A) site. The ESE is
juxtaposed to the ESS3 within tat/rev exon 3. The regions
which are spliced to form the tat mRNA are indicated by
dashed lines. (B) Sequences of ESS2 and ESS3.
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MATERIALS AND METHODS |
Plasmid constructions.
Mutations were made either by
PCR-mediated site-directed mutagenesis or by subcloning of annealed
oligonucleotide pairs (36). Mutations were confirmed by
sequencing. p
ESS has been described previously (2, 39)
and was used as the parent plasmid for all mutant HIV-1 ESS3 minigene
constructs shown in Fig. 3A. pHS1-X and pESS10 have been described
previously (2). pHS1-X+ESS3 was constructed by standard
cloning techniques by replacing the 20-nucleotide (nt) ESS2 sequence
with a 20-nt ESS3 sequence. All plasmids were linearized with
HindIII at nt 6026 of pNL4-3 (GenBank accession no.
M19921) and used as templates to synthesize RNA substrates by in vitro
transcription with T3 RNA polymerase.
pAdML was a generous gift from Dr. R. Reed (Department of Cell Biology,
Harvard University) (
29). pAdML+ESS3S and pAdML+ESS3A
were
created by subcloning the
AccI-
AccI fragments
containing
the ESS from pESS3S and pESS3A into the unique
AccI site downstream
from the AdML 3' splice site (see Fig.
4A). To prepare DNA templates
for T7 RNA polymerase transcription,
pAdML was linearized with
BamHI and pAdML+ESS3S and
pAdML+ESS3A were digested with
MfeI.
pTAT118 has been described previously (
3) and was used to
make RNA competitors containing ESS2. pTAT90S was constructed
by
subcloning the
AccI-
MfeI fragment from pESS3S
between the
AccI
and
EcoRI sites of pBluescript
SK (Stratagene, La Jolla, Calif.).
pEC4× was constructed according to
methods previously described
(
27); it contains four tandem
copies of the ESS2 sequence. Both
plasmids were linearized with
BamHI for transcription with T7
RNA polymerase. pHRR80
(obtained from Ambion, Austin, Tex.) contains
an 80-nt fragment of
human 18S rRNA (nt 715 to 794 the sequence
under GenBank accession no.
M10098), inserted downstream from
the T7 promoter sequence of pUC18 in
the antisense orientation.
The construct was linearized with
HindIII and transcribed with
T7 RNA polymerase to
generate a 116-nt RNA which was used as the
control RNA in RNA
competition assays.
In vitro RNA splicing and RNA competition assays.
RNA
substrates of pESS and its derivatives were synthesized with T3 RNA
polymerase and labeled with [32P]UTP. Uncapped RNA
competitors were synthesized with T7 RNA polymerase with a
MegaShortscript kit (Ambion). After DNase I treatment, the
reaction mixtures were extracted twice with phenol-chloroform and
passed through 1-ml Sephadex G-50 columns to remove the unincorporated nucleotides. The RNA was precipitated with ethanol in the presence of
0.3 M sodium acetate and dried under vacuum. The amount of RNA was
quantitated spectrophotometrically, and the RNA was shown to be intact
by agarose gel electrophoresis. HeLa cell nuclear extracts were
prepared and splicing reactions were performed as previously described
(2, 16, 25). For RNA competition assays, the substrates were
spliced for 2 h at 30°C after the nuclear extracts were
preincubated for 15 min with various amounts of nonradioactive
competitor RNAs at the same temperature. The products of the splicing
reactions were separated on 7 M urea-4% polyacrylamide gels. Gels
were scanned for radioactivity and quantitated with an Instantimager
analysis system (Packard, Meriden, Conn.). The amounts of spliced
product were calculated based on the specific activity of the
[32P]UTP used for labeling the precursor RNAs and were
expressed as femtomoles of spliced product.
Analysis of spliceosome complexes.
Complex E formation and
analysis were carried out as previously described (29). In
brief, [32P]UTP-labeled RNA substrates were transcribed
from linearized pESS3S and pESS3A DNA, labeled RNA was incubated under
splicing conditions for 30 min in the absence of both ATP and
MgCl2, and the complexes were separated on Sephacryl S500
columns (1.5 by 100 cm). Complex A assembly was analyzed by
nondenaturing gel electrophoresis as previously described
(23). The RNA substrates, ESS3S and ESS3A, were
incubated at 30°C for various times in the presence of ATP (0.5 mM)
and MgCl2 (3 mM). Complex formation was determined by
electrophoresis on 4% native polyacrylamide gels with an
acrylamide-to-N,N'-methylenebisacrylamide
ratio of 80:1.
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RESULTS |
ESS3 is functional when inserted downstream of a suboptimal
heterologous 3' splice site.
We showed previously that the 3'
splice site of tat exon 2 contains a nonconsensus
polypyrimidine tract and that the negative effect of ESS2 is
orientation dependent (2, 39). We have also shown that the
ESS2 in tat exon 2 is active when inserted downstream of a
suboptimal heterologous src splice site (2). We
examined whether ESS3 was also active when placed downstream of a
suboptimal heterologous 3' splice site. For this experiment, we used
construct pESS in which the ESS2 sequence had been deleted and the
20-nt ESS3 had been inserted in the sense and antisense orientations
within tat exon 2 to create pESS3S and pESS3A, respectively (Fig. 2A). These constructs were used as
templates to synthesize RNA substrates for in vitro splicing assays
which were carried out for various times (Fig. 2B). As shown in Fig.
2C, the rate of splicing of substrate ESS3S was approximately 25% that
of ESS3A. ESS3A, on the other hand, was spliced with an efficiency
similar to that of ESS (data not shown). These results indicate that ESS3 behaves similarly to ESS2 in that it inhibits the efficiency of
splicing when placed downstream of a suboptimal heterologous 3' splice
site.
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FIG. 2.
ESS3 is functional when inserted downstream of a
suboptimal heterologous 3' splice site. (A) In pESS the region
containing ESS2 has been deleted from pHS1-X and replaced with a
multiple cloning site (MCS). ESS3 was inserted into pESS in both the
sense (pESS3S) and antisense (pESS3A) orientations downstream of
tat exon 2. [32P]UTP-labeled RNA substrates
were synthesized as described in Materials and Methods and incubated in
HeLa cell nuclear extracts for 2 h at 30°C under standard
splicing conditions with 3 mM Mg2+ (25). T3,
phage T3 RNA polymerase promoter; SS, splice site. Cap and pA, same as
in Fig. 1. (B) Time course of splicing for ESS3S and ESS3A RNA
substrates. Products of in vitro splicing were analyzed with denaturing
polyacrylamide gels. The substrate for each lane is specified at the
top, and the time (in minutes) of the reaction is shown in parentheses.
The identities of bands are illustrated on the left side of the
autoradiogram (top to bottom: lariat exon, pre-mRNA, spliced product,
lariat, and 5' exon). (C) Quantitation of the spliced products from RNA
substrates ESS3S (open symbols) and ESS3A (filled symbols) shown in
panel B.
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ESS3 is composed of two subelements, and each element can inhibit
splicing independently.
We have previously defined the ESS3 as a
20-nt sequence (3). We next determined whether the
inhibitory activity of ESS3 was confined to certain sequences within
ESS3. To this end, 2-nt substitution mutations were created to examine
the significance of individual bases (Fig.
3A). Four mutant substrates (ESS4445, ESS4849, ESS5657, and ESS5859) showed significantly higher levels of
splicing than the other mutant substrates, indicating that these
nucleotides are important for the activity of ESS3 (Fig. 3B). However,
the two groups of essential nucleotides in ESS3 (AGAUCC and
UUAG) were separated by 6 nt instead of being in one contiguous region,
as in ESS2. Since all of the mutant substrates were spliced with
efficiencies lower than that of ESS3A, we determined the effect of
combining mutations at positions 4445, 4849, 5657, and 5859 (ESS3D)
(Fig. 3C). The level of splicing of this mutant was increased to
comparable to that of ESS3A. This suggested that the two regions of
ESS3 act additively to inhibit splicing.
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FIG. 3.
ESS3 is composed of two subelements. (A) Schematic
diagram of the ESS3 mutants derived from pESS (see Materials and
Methods). Abbreviations are defined in the legend for Fig. 2A. (B)
Effects of single 2-nt substitution mutations in the ESS3 sequence on
splicing efficiency. (C) Effects of the two regions of ESS3 and
combination mutation on the splicing of tat exon 2. Splicing
assay procedures and the identities of the band are as described in the
legend for Fig. 2.
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To test whether the two groups of essential nucleotides
represented two subelements within ESS3, we constructed templates
containing either the first or second 10 nt of ESS3 (pESS4453
or
pESS5463, respectively). RNA substrates synthesized from these
templates were analyzed by in vitro splicing (Fig.
3C). Splicing
of
both ESS4453 and ESS5463 mutant substrates was inhibited but
to a
lesser extent than that of the substrate containing the entire
ESS
sequence (ESS3S). These results implied that there are two
subelements
within ESS3 that act independently and additively
to inhibit splicing.
ESS3 is less active in the context of optimal 3' splice sites.
Results previously acquired at our laboratory have indicated that ESS2
activity is strongly influenced by the strength of the upstream 3'
splice site which is inhibited (39). Staffa and
Cochrane, who used in vivo transfection studies, came to similar conclusions regarding ESS3 (42). However, the DNA constructs used in their study contained only the upstream ESS3 subelement (42). To determine if ESS3 inhibits splicing in the
context of an optimal 3' splice site, the entire 20-nt ESS3 was
inserted into pTat3'C+ESS in either the sense or the antisense
orientation (Fig. 4A). RNA substrates
synthesized from these constructs have optimal polypyrimidine tracts
and consensus 3' splice sites flanking tat exon 2 (39). The effects of the improved 3' splice sites on the
function of ESS3 were tested by in vitro splicing assays carried out
for the various times shown in Fig. 4B. The rate of splicing of
substrate Tat3'C+ESS3S was approximately 75% that of Tat3'C+ESS3A.
Comparison of the data in Fig. 2C and 4C indicated that the effect of
ESS3 was significantly weakened in the presence of an optimal 3' splice
site.
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FIG. 4.
Effect of optimal 3' splice sites on the function of
ESS3. (A) Organization of DNA constructs (see Materials and Methods).
pTat3'C+ESS is derived from pESS and has an optimized
polypyrimidine tract and a consensus 3' splice site (39).
ESS3 was inserted into pTat3'C+ESS in both the sense (pTat3'C+ESS3S)
and antisense (pTat3'C+ESS3A) orientations downstream from the 3'
splice site. MCS, multiple cloning site. (B) Time course of splicing
for Tat3'C+ESS3S and Tat3'C+ESS3A RNA substrates. The corresponding RNA
substrates were spliced and analyzed and the identities of the bands
(left) are as described in the legend for Fig. 2. (C) Quantitation of
the spliced products from the RNA substrates, Tat3'C+ESS3S (open
symbols) and Tat3'C+ESS3A (filled symbols), shown in panel B. (D) ESS3
was inserted downstream of the 3' splice site of pAdML in both the
sense (pAdML+ESS3S) and antisense (pAdML+ESS3A) orientations. Plasmid
linearization and transcription were performed as described in
Materials and Methods. RNAs were spliced in vitro for various times as
described in the legend for Fig. 2 with Mg2+ at a final
concentration of 1 mM. (E) Quantitation of the spliced products from
the RNA substrates, AdML+ESS3S (open symbols) and AdML+ESS3A (filled
symbols), shown in panel D.
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To further examine the effect of ESS3 in the context of an unrelated
RNA substrate that is not subject to regulation, we placed
ESS3 in both
orientations within exon 2 of an adenovirus major
late gene construct
(pAdML) (Fig.
4A). RNA substrates synthesized
with this
construct have been widely used to investigate basic
splicing
mechanisms, since these substrates contain consensus
polypyrimidine
tracts and branch point sequences (
5,
9,
10,
30,
44). As
shown in Fig.
4D and E, when the splicing
assays were carried out with
1 mM Mg
2+ for various times, the rate of splicing of
substrate AdML+ESS3S
was found to be approximately 25% that of
substrate AdML+ESS3A.
These data support the notion that ESS3 is
active in the context
of a heterologous RNA substrate.
Spliceosome assembly is inhibited prior to the formation of a
complex (complex A) with an RNA substrate containing ESS3.
Previous results indicated that splicing is inhibited at an early step
of spliceosome formation by ESS2 (3). Splicing of ESS3S
appeared to be inhibited before the first step of the splicing reaction, since no lariat-exon intermediate accumulated even at the 2-h
point (Fig. 2B). To further define the mechanism by which ESS3 acts to
inhibit splicing, we investigated the step at which spliceosome
assembly is blocked by ESS3. Functional spliceosomes are assembled in a
stepwise manner through a series of intermediate complexes, termed
complexes E, A, B, and C (24, 29). Formation of complex E,
or the commitment complex, appears not to require the presence of ATP
(6, 29). This is followed by a series of ATP-dependent steps
leading to formation of complexes A, B, and C (26, 31).
Two RNA substrates, ESS3S and ESS3A, in which ESS3 was inserted
in the sense and antisense orientations, respectively, were
used to
study formation of spliceosome complexes. Both substrates
were
incubated with HeLa cell nuclear extracts under standard
splicing
conditions in the absence of ATP. The reaction mixtures
were then
subjected to gel filtration analysis under conditions
previously shown
to separate complex E and a nonspecific RNA-protein
complex (complex H)
(
29). Incubation of control RNA lacking
splice sites
resulted in formation of only complex H, whereas
the control
substrate AdML formed both complexes H and E (data
not shown) as
described by Michaud and Reed (
29). The elution
profiles of
the RNA substrates ESS3S and ESS3A are shown in Fig.
5A. These results indicated that two
distinct peaks were present
when either RNA substrate was used. One
peak was eluted at a position
corresponding to complex H, and the other
was eluted at the position
expected for complex E. These results
suggested that the step
of splicing inhibited by ESS3 occurred
subsequent to formation
of complex E.
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FIG. 5.
Inhibition of spliceosome complex formation by the HIV-1
ESS3. (A) RNA substrates (ESS3S and ESS3A) were incubated with HeLa
cell nuclear extracts. The extracts were preincubated at room
temperature to deplete endogenous ATP levels. Incubation was carried
out in the absence of ATP and Mg2+ at 30°C for 25 min.
The reaction mixtures were applied to a Sephacryl S500 column under
conditions previously described (29). The positions of
complexes E and H are shown. Numbers on the abscissa and ordinate
indicate the fraction number and the amount of radioactivity in the
samples (counts per minute), respectively. (B) RNA substrates were
incubated with HeLa cell nuclear extracts in the presence of ATP and
Mg2+ for the indicated times, and the complexes were
analyzed on a 4% nondenaturing polyacrylamide gel.
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To further test the effect of ESS3 on spliceosome assembly, we
determined whether complex A was assembled with either or both
substrates. After incubation of RNAs with nuclear extracts in
the
presence of ATP for various times, the reaction mixtures were
applied
to native polyacrylamide gels to examine the assembly
of complexes
(Fig.
5B). Complex A formation was detected for substrate
ESS3A after
10 min. Substrate ESS3S, on the other hand, did not
form a significant
amount of complex A, even after a 30-min incubation.
It appears,
therefore, that assembly of both substrates can proceed
to complex E
formation but that the ESS3s substrate cannot form
complex A. Thus, the
inhibition of splicing caused by ESS3 takes
place prior to the
formation of complex A.
Evidence that ESS2 and ESS3 share a cellular factor(s)
necessary for the inhibition of tat mRNA splicing.
Previous assays with unlabeled competitor RNAs containing ESS2
suggested that the inhibition is mediated by a cellular factor(s) which binds to ESS2 (3). We next tested whether the
inhibition of ESS3 was also mediated by a cellular factor and, if so,
whether ESS3 and ESS2 bound to the same factor. RNA competition assays were carried out with three different RNA substrates (Fig.
6A). Substrates HS1-X and
ESS10 had been previously used to study the function of ESS2
(2, 3). HS1-X contains the wild-type ESS2 sequence, whereas
ESS10 contains a 10-nt heterologous sequence inserted in place of
ESS2. The former substrate is spliced inefficiently whereas the latter
substrate, because of the mutated ESS2 sequence, is spliced
efficiently. The third RNA substrate was synthesized from HS1-X+ESS3,
in which the 20-nt ESS3 sequence of HS1-X had been substituted for the
20-nt ESS2 sequence. As shown in Fig. 6B, splicing of HS1-X+ESS3 was
very inefficient. Addition of competitor RNA containing ESS3 (TAT90S)
derepressed splicing of HS1-X+ESS3 but had little effect on the
efficiency of splicing of the ESS10 substrate. This suggested that a
cellular factor specifically binds to ESS3 to mediate the inhibition of
splicing. Interestingly, addition of TAT90S RNA also derepressed
splicing of substrate HS1-X, which contains the ESS2 sequence. This
suggests that the two ESS sequences bind to a common factor or factors.
Comparison of the amounts of spliced product at the different molar
excesses of competitor shown in Fig. 6B indicated that TAT90S competed approximately twofold more efficiently with the ESS3 substrate than
with the ESS2 substrate. Splicing assays carried out at lower competitor concentrations (from 50- to 250-fold molar excesses) confirmed this small difference in efficiency (Fig.
7A). This suggests that ESS2 may have a
slightly higher affinity for the putative common factor than does ESS3.
A second RNA competitor containing the ESS2 sequence (TAT118) was
tested against the same set of RNA substrates. As shown
previously (3), this RNA specifically derepressed
splicing of substrate HS1-X but had no effect on the splicing of
ESS10, containing a mutated ESS2 (Fig. 6C). Addition of the ESS2
competitor also resulted in an increase in the splicing efficiency
of substrate HS1-X+ESS3. Like the ESS3 competitor, TAT118 RNA
competed more efficiently with the ESS3 substrate than with the
ESS2 substrate. Splicing assays carried out at lower competitor concentrations confirmed this small difference (Fig. 7B).
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FIG. 6.
ESS2 and ESS3 share a cellular factor(s) necessary for
the inhibition of tat mRNA splicing. (A) Diagrams of
constructs used to synthesize RNA substrates for competition assays.
SS, splice site. (B to E) In vitro splicing assays were carried out in
the presence of increasing amounts of unlabeled RNA (250, 500, and
1,000× specify the molar excesses relative to the substrate
concentrations). The identities of bands are as described in the legend
for Fig. 2B. (B) TAT90S RNA (ESS3 plus the same flanking sequence as
ESS2). (C) TAT118 RNA (ESS2 plus the flanking sequence). (D) EC4× RNA
(four tandem copies of ESS2 without the flanking sequence). (E) HRR80
RNA (control). This RNA contains an 80-nt fragment of human 18S rRNA
sequence but does not contain either ESS sequence.
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FIG. 7.
Competition assays against the same set of RNA
substrates as shown in Fig. 6A but with lower molar excesses of
competitor RNAs. 50, 100, and 250× specify the molar excesses relative
to the substrate concentrations. The identities of the bands are as
described in the legend for Fig. 2B.
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Because the ESS2 in TAT118 and the ESS3 in TAT90S share flanking
sequences, we tested a second RNA competitor (EC4×), which
contains four tandem copies of the ESS2 sequence with no flanking
sequence. As shown in Fig.
6D and
7C, this competitor produced
derepression of splicing of HS1-X and HS1-X+ESS3 similar to
that
by TAT118, indicating that no sequence except that of ESS2
is
required for the effect. Surprisingly, the concentration dependence
of derepression by EC4×, with four tandem copies of the ESS2 sequence,
was not significantly different from that of TAT118, with one
copy
(compare Fig.
6D with Fig.
6C and Fig.
7C with Fig.
7B).
This suggests
that some flanking sequence may be necessary for
binding the
putative factor(s).
To further test the specificity of the competitor RNAs, we used an
irrelevant RNA, HRR80, which contains an 80-nt sequence
derived
from human 18S rRNA. This RNA did not significantly affect
the
splicing efficiencies of the three RNA substrates (Fig.
6E).
This
indicated that the derepression by the ESS2 and ESS3 competitor
RNAs
was specific. These data strongly suggest that the inhibitory
effect of ESS3 is mediated by a cellular factor(s) and that ESS3
and
ESS2 may bind to a common cellular factor.
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DISCUSSION |
Our results, based on in vitro splicing assays, have
indicated that the ESS present in the second tat coding exon
of HIV-1 (ESS3) is comprised of two smaller elements and that each of
these smaller elements can inhibit splicing when placed in the exon downstream of a heterologous weak 3' splice site. We have shown that
the core sequences necessary for ESS3 activity are AGAUCC and UUAG and that the core sequence necessary for ESS2 activity is CUAGACUAGA (39). The sequences necessary for
ESS activity appear to have some common sequence features. In the first
part of the sequence AGAPyPy is common, and in the downstream regions PyUAG is common. It is possible that two binding sites on the RNA are
necessary for full activity of an ESS. ESS2 has two repeats of the
sequence CUAGA, and this may also indicate the presence of two binding
sites.
The sequences of several other potential ESS elements are known, and it
is of interest to compare them to those of ESS2 and ESS3. HIV-1 cryptic
exon 6D is a small exon present in the env gene. The
inclusion of this exon results in the production of a chimeric
Tat-Env-Rev fusion protein called Tev (28). Recently it has
been shown that this exon contains a potential ESS which may act to
inhibit splicing at the upstream cryptic 3' splice site. The sequence
of this ESS is CAAUAGUAGUAG (51). The human fibronectin alternative EDA exon contains a cis splicing
element comprised of an ESE (GAAGAAGA) and an ESS (CAAG)
(8). The K-SAM alternative exon of the human fibroblast
growth factor receptor 2 gene has been reported to contain an ESS
element with the sequence UAGGGCAGGC. Further experiments
suggested that the functional portion of this element is limited to the
sequence UAGG and that the dinucleotide AG appears to be of particular
importance to its activity (14, 15). Although the sequence
homology among these reported ESS elements is minimal, most of them
contain UAG, and all elements contain at least one AG.
Data obtained by our laboratory have suggested that both ESS2 and ESS3
act to inhibit splicing by blocking the formation of functional
spliceosomes at an early step (3). We showed in Fig. 5A that
a substrate containing a functional ESS3 forms a complex that is eluted
at the position expected for complex E. However, since the transition
from complex E to complex A may involve several intermediate steps
(9), we cannot determine whether the complexes shown in Fig.
5A are functional or aberrant complexes E, and further
characterization of these complexes will be required to answer this
question.
RNA competition assays suggested that a cellular factor or factors bind
to the ESS sequence to mediate inhibition of splicing. There are a
number of examples of RNA-binding proteins that inhibit splicing at an
early step. In Saccharomyces cerevisiae, the ribosomal protein L32 binds to the first exon of its own pre-mRNA to prevent the
formation of functional complex A (49). In
Drosophila, Sxl protein promotes the female-specific
splicing of tra pre-mRNA by blocking the utilization of the default
tra 3' splice site. Sxl regulation requires a poly(U)
sequence located in the polypyrimidine tract of the default 3' splice
site. Thus, it may inhibit splicing at an early step by competing with
U2AF for the binding to the polypyrimidine tract (48).
Splicing of the Drosophila P-element third intron is
repressed in the soma at an early step by blocking of U1 snRNP binding
to the actual 5' splice site and stabilization of U1 snRNP binding to
an inactive pseudo-5' splice site. This process is regulated by an
alternative splicing factor, PSI (P-element somatic inhibitor), and a
general splicing factor, hrp48 (homologous to mammalian hnRNPA1)
(1, 40, 41).
Our data suggest that inhibition of splicing by ESS2 and ESS3 may
involve the action of a common factor or factor(s). We showed that ESS2
RNA competitors are able to abrogate the inhibition by ESS3 and vice
versa. This is not surprising, since the sequences necessary for ESS2
and ESS3 activities have some common features. The small but
reproducible difference between the efficiencies of derepression of the
two competitor RNAs could be explained by different binding affinities
resulting from the sequence difference between ESS2 and ESS3.
Alternatively, it is possible that inhibition by ESS2 and ESS3 may
require the formation of a complex by a number of proteins. Thus, a
unique factor may bind to each of ESS2 and ESS3, but the complexes may
share one or more limiting factors.
The accumulation of unspliced and partially spliced HIV-1 RNA is
necessary for HIV-1 replication. It is of interest that splicing at
both tat 3' splice sites appears to be inhibited at an early step by ESS elements by similar mechanisms and with mediation by a
common cellular factor(s). This would allow a coordinated response of
the two splice sites to the cellular environment, with the result being
the preservation of the unspliced and singly spliced RNAs necessary for
Rev-dependent transport to the cytoplasm.
| |
ACKNOWLEDGMENTS |
We thank Stanley Perlman, Richard Roller, and Brad Amendt for
critical reading of the manuscript.
This research was supported by PHS grant AI36073 from the National
Institute of Allergy and Infectious Diseases.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Microbiology, University of Iowa, 3770 Bowen Science Building, Iowa
City, IA 52242. Phone: (319) 335-7793. Fax: (319) 335-9006. E-mail: marty-stoltzfus{at}uiowa.edu.
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Abstract
Introduction
