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INTRODUCTION |
The genome of HIV-11 is
transcribed as a full-length 9-kilobase mRNA that can follow two
different "pathways" (1-4). Early in infection, the primary HIV
transcript is fully spliced to mRNAs of approximately 2 kilobases
in length and then transported from the nucleus to the cytoplasm, where
the mRNAs are translated into a variety of small accessory
proteins, such as Tat, Rev, and Nef (3-8). Later in the virus
lifecycle, unspliced or partially spliced RNAs are exported from the
nucleus to the cytosol, where the unspliced RNA is packaged into virus
particles; the different partially spliced mRNAs are translated into
Gag, Pol, Env, and smaller proteins, such as vif, vpR, and vpU (4,
6-10). This temporal switch in the HIV RNA complexity and the coding
potential is mediated by virus coded Rev protein (11-19), an
approximately 16-kDa basic RNA-binding protein with two functional
domains (Fig. 1A). The N-terminal domain is involved in RNA
binding and oligomerization (20-25), whereas the more C-terminal
activation domain (13, 26-30) is thought to interact with the nuclear
pore-associated proteins, such as hRIP/Rab (31-35), CRM1 (36-39), and
nuclear eIF-5A (40, 41). In the absence of Rev, the RNA transcript is
fully spliced; the full-length transcript is never observed in the
cytoplasm (3, 8, 9, 13, 16-19).
Rev binds specifically to a highly structured 244-nucleotide RNA
sequence, the Rev response element (RRE), located in the env
gene of the primary transcript (16, 42, 43); this RNA binding (43-48)
is essential for the nuclear export of unspliced and partially spliced
HIV mRNAs (8, 9, 15, 16,). Lack of functional Rev or RRE completely
blocks viral replication. The secondary structure of the RRE RNA is
presumed to fold into four stem-loops, designated A, C, D, and E (47)
or stem-loops I, III, IV, and V (16), which have branched stem-loop
structure, B/B1/B2 (47); or into stem loop II A/B/C (16), linked by a
central loop (Fig. 1B). Several studies have shown that most
of this RRE structure is dispensable for Rev activity, that a minimal
structure composed of the B/B1/B2 (or stem loop II A/B/C) subdomain was
active both in vitro and in vivo, and that
specific mutations within this region eliminate Rev binding in
vitro and trans-activation in vivo (47-58).
Notwithstanding the functional definition of the minimal Rev-responsive
sequence, Mann et al. (59) have reported that a larger
RRE-RNA structure of 351 nucleotides is required for complete
biological activity. According to their model, the larger RRE is deemed
to act as a "molecular rheostat," which binds multiple Rev monomers
up to a functionally optimal threshold of 10-12 Rev monomers per RNA
molecule. Furthermore, although a number of groups have demonstrated
that a single Rev monomer bound to a high affinity site on the RRE,
after which additional Rev molecules were recruited through protein-RNA
and protein-protein interactions (20, 58, 60, 61), others have shown
that Rev bound to its target RNA in an oligomeric form (62, 63).
Although Rev-RRE interaction has been extensively studied, a number of
fundamental questions remain to be answered. These include whether Rev
acts solely to transport RRE-containing RNA to the cytoplasm or also
actively inhibits splicing, the precise number of Rev monomers bound to
each RRE, whether Rev monomers bind sequentially or in oligomeric form
and whether this binding is cooperative, and precise knowledge of the
various rate constants of Rev-RRE binding. We used surface plasmon
resonance (SPR) measurements (see Refs. 63 and 64 for review) to
determine the kinetics of Rev-RRE binding. We developed a novel
approach to bind RNA to the sensor surface. Rev in solution was then
passed through the flow cell under conditions that allowed Rev-RRE
interaction. The accumulation of macromolecules on the sensor surface
alters the refractive index of the solution in the vicinity of the
surface, which is detected and quantified by the instrument, Biacore
2000 (Biacore Inc., Piscataway, NJ). Depending on the binding
conditions, initial velocity or steady-state kinetic data can be
collected in real time. Our results show that RRE can be subdivided
into the following: 1) a minimal site that interacts with one molecule of Rev, 2) a core site that can bind at least two Rev monomers, and 3)
the full-length RRE that binds four Rev molecules. With excess Rev
concentration, additional binding to other low affinity sites was
observed. The high affinity interactions were very strong and specific,
with KD values in the 10
10 to
10
11 M range. Finally, this method can be
used to rapidly and conveniently investigate RNA-protein interactions
in general and was quite useful in rapidly evaluating the effects RNA
and protein mutations on the binding, as well as inhibitors of these interactions.
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EXPERIMENTAL PROCEDURES |
Construction and Amplification of RRE Element DNA
Fragments--
The RRE element had been subcloned form the proviral
plasmid pNL432 (positions 7749-7992, 244 base pairs) as described
(47). RRE mutants used here have been described in Refs. 47 and 56, except for the B/B1/B2 stem-loop RRE fragment (RRE/SLIIB, spanning nucleotides 53-110, but deleting nucleotides 90-104) (Fig.
2A), which was generated by hybridization of two
oligonucleotides tagged with the bacteriophage T7 RNA
polymerase promoter and (dT)18 at the 5'- and 3'-ends,
respectively. All other RRE DNA elements were amplified and tagged at
the 5'-end with a T7 promoter and at the 3'-end with
(dT)18 by PCR using oligonucleotides complementary to the
5'- and 3'-ends of the desired RRE element at the following positions
(47, 56): RRE, RRE-3A, and RRE/Z, 5'-positions 1-18, 3'-positions
227-244; RRE/T, 5'-positions 40-57, 3'-positions 196-213; RRE/HH,
5'-positions 50-68, 3'-positions 96-113. The TAR-DNA element used was
generated by hybridization of the following oligonucleotides
tagged at the 5'-end with the T7-RNA polymerase promoter and with a (A)18 tail at the 3'-end:
5'-TGAATTGTAATACGACTCACTATAGGCTTTTTGCCTGTACTGGGTCTCT CTGGTTAGACCAGATCTGAGCCTGGGAGAAAAAAAAAAAAAAAAAAA-3' and
5'-TTTTTTTTTTTTTTTTTTCTCCCAGGCTCAGATCTGGTCTAACCAGAGAGACCCAGTACAGGCAAAAAGCCTATAGTGAGTCGTATTACAATTCA-3'.
In Vitro RNA Synthesis and Purification--
T7
promoter-tagged PCR amplified RRE DNA fragments (0.5 pmol) were used as
templates to transcribe RRE RNA (47, 56) using the Riboprobe Gemini
Systems kit from Promega Corp. (Madison, WI). Full-length RNA tagged at
the 3'-end with (A)18 was purified by hybridization to
oligo(dT)25 linked magnetic beads (Dynal Corp., A.S., Oslo,
Norway) as follows. The beads (50 µl) were washed twice as described
by the manufacturer in deionized water and once in hybridization buffer
(30 mM sodium phosphate, pH 7.4, 450 mM NaCl, 3 mM EDTA, 0.1% Triton X-100) and suspended in 25 µl of
the same buffer. Freshly transcribed RNA (1 µmol) in 25 µl of
hybridization buffer was added to the beads and incubated with
occasional mixing for 30 min at room temperature. The beads were
immobilized with a magnet and washed twice with 100 µl of fresh
hybridization buffer. Bound RNA was eluted by adding 50 µl of
deionized water and incubating for 5 min at room temperature. The beads
were again immobilized using a magnet and the eluted RNA was ethanol
precipitated and made up to a final concentration of 0.5 mg/ml in TE
buffer (10 mM Tris-HCl, pH 8.0, and 1 mM EDTA).
Proteins and Peptides--
HIV-1 Rev and Nef proteins were
expressed in Escherichia coli from the thermally inducible
coliphage
pL promoter and purified as described (66).
Before use in these experiments, the previously purified Rev protein
was reassessed by gel electrophoresis and gel filtration under
denaturing conditions. The RRE RNA binding potential was determined by
electrophoretic mobility shift assay (EMSA) and filter binding assays
as described (47, 56). The in vivo functional potential of
Rev protein and Rev domain peptides were determined by protein
transfection (67) in Rev-dependent HIV-1 gag expression
assay essentially as described (47, 56). Wild type Rev fused to MS
coliphage-2 coat protein (MS2-C) and mutant versions of Rev in the same
context were expressed in E. coli (68), molecularly tagged
to maltose-binding protein. For SPR use, the maltose-binding protein
moiety was excised by protease X digestion and further purified as
described (68). The different Rev peptides used in this study were
purified by two cycles of reversed phase high pressure liquid
chromatography on C18 columns, and their purity was checked by
automated Edman degradation through 30 or 40 cycles and mass
spectrometry. RRE RNA binding potential of individual peptides was also
determined by EMSA and filter binding assay in the presence of heparin,
but without tRNA competitor (47, 56).
SPR Analysis of Rev-RRE Interactions--
A Biacore
2000TM instrument was used throughout this study. A
5'-biotin U-(dT)18 oligomer (5 µM in 20 mM Tris-HCl, pH 7.4, containing 0.15 M NaCl)
was bound to the surface of streptavidin-coated sensor chips (Biacore
Inc.) at a flow rate of 5 µl/min for 10 min. The chips were then
washed 5 times with 25 µl each of 50 mM NaOH at a flow
rate of 5 µl/min to stabilize the surface prior to the binding of
RNA. (A)18 containing RRE RNAs were immobilized in each
flow cell by hybridization to the 5'-biotin U-(dT)18
oligomer in running buffer (10 mM HEPES, pH 7.4, 450 mM NaCl, 3 mM EDTA, 0.1% Triton X-100) at a
flow rate of 1 µl/min at 25 °C. The duration of each injection was
varied depending on the surface RNA density that was desired for each
individual experiment. All RNAs were activated by sequential incubation
at 42 °C, at room temperature, and on ice, each for 10 min in
running buffer prior to injection over the sensor chip. Following RNA
loading, 20 µl of protein dissociation buffer (10 mM
HEPES, pH 7.4, 250 mM NaCl, 3 mM EDTA, 0.05%
Triton X-100, 0.1% SDS) was passed over the surface at a flow rate of
10 µl/min to ensure that all RNAs were tightly bound. In general, RNA
chips with between 10 response units (RUs) (low density) and 200 RUs
(high density) immobilized RNA were used in the various experiments.
Purified protein in running buffer was passed over the RNA surface
through multiple rounds at a flow rate of 10-100 µl/min depending on
whether data pertaining to steady-state or initial velocity conditions
were to be collected. Bound proteins were completely removed after each
round of injection by passing 20 µl of dissociation buffer at a flow
rate of 10 µl/min over the sensor surface. This treatment did not
remove more than 1% of bound RNA over the course of the experiment. To
regenerate the surface, the RNA was completely removed by injecting 75 µl of 50 mM NaOH at a flow rate of 25 µl/min, allowing
the sensor chip to be reused multiple times.
Data Analysis--
Nonlinear fitting of the primary sensogram
data was used to calculate the association and dissociation rate
constants using the BIAevaluation 2.1 software (Biacore Inc.) and as
recommended (69, 70). The dissociation rate constants,
kd1 and kd2, for two parallel dissociation
reactions, were derived for Rev binding to RRE, RRE/T, and RRE/HH using
the equation,
|
(Eq. 1)
|
where R(t) is the response at time
t (in seconds), Ro is the total response
at the start of dissociation at to, R1 is the contribution to
R0 from component 1, (Ro
R1) is the contribution to Ro
from component 2. The association rate constants,
ka1 and
ka2, for Rev binding to two ligand
components on RRE, RRE/T, and RRE/HH were then derived using the
equation,
|
(Eq. 2)
|
where Req1 and
Req2 are the steady state response levels for
components 1 and 2, respectively, C is the molar
concentration of the analyte, and to is the start
time for association. The dissociation rate constant,
kd, for Rev binding to the single site on RRE/SLIIB
was derived using the equation,
|
(Eq. 3)
|
where R(t) is the response at time
t (in seconds), Ro is the response at the
start of dissociation, and to is the start time for
dissociation. The association rate constant, ka, for
the single site Rev-RRE/SLIIB binding reaction was derived using the
equation,
|
(Eq. 4)
|
where Req is the steady state response
level, C is the molar concentration of the analyte, and
kd is the dissociation rate constant. Affinities
were calculated from the rate constants; KD = kd/ka.
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RESULTS |
RNA Preparation and Binding Protocol--
The HIV-1 Rev protein is
crucial for virus replication; viral replication is abolished in the
absence of functional Rev. Fig. 1A shows the crucial
functional domains of Rev located between amino acids 22 and 85, and
the putative secondary structure of the cognate RNA target, the RRE, is
shown in Fig. 1B. Kinetic analysis of Rev-RRE interactions
was done by means of SPR measurements using a biosensor instrument,
Biacore 2000. For this purpose, we bound the different RNA targets to
the sensor surface, rather than Rev, because the locally high density
of multiple immobilized Rev monomers may induce unwanted aggregation
and trap free-flowing RRE-RNAs nonspecifically. To immobilize different
RNAs in an easily reversible manner, we developed a novel hybridization
technique, shown diagrammatically in Fig. 1B, that is a
modification of the technique described by Wood (71) for DNA-DNA
hybridization. Briefly, RRE-DNA fragments tagged at the 5'-end with the
T7-RNA polymerase promoter, and 18 dT nucleotides at the
3'-end were generated by PCR (see under "Experimental Procedures").
3'-poly(A)-tailed RRE RNA was synthesized by T7 transcription, purified
by oligo(dT) chromatography, and then hybridized to a
biotin-U-(dT)18 oligomer that had been previously bound to
a streptavidin-derivatized sensor chip. Using this protocol, the RNA
molecules were all presented in the same orientation, which could
freely interact with Rev. Injection of a low salt buffer containing
0.1% SDS effectively removed bound Rev at the end of each sensogram
run without removing the RNA. The total amount of RNA lost from the
sensor surface over the course of a typical experiment was less than
1% (data not shown). To completely regenerate the sensor chip, the RNA was removed by brief injection of 50 mM NaOH, allowing the
chip to be re-used multiple times with different RNA preparations.

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Fig. 1.
Schematic diagram of Rev, secondary structure
of RRE RNA, and overview of the RNA capture and SPR protocol.
A, schematic illustration of HIV-1 Rev protein with the
putative functional domains involved in nuclear localization and RNA
binding, oligomerization, and activation highlighted and identified.
B, (dT)18 oligomer biotinylated at the 5'-end
was bound to streptavidin covalently linked to a carboxymethyl dextran
matrix attached to a thin gold film over a glass support. RRE-RNAs were
immobilized by hybridization between the 3'-poly(A) tail and the
surface-bound oligo(dT) followed by multiple rounds of Rev
injection.
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Specificity of Rev-RRE Binding--
To test the specificity of Rev
binding in this protocol, Rev was injected over a sensor surface to
which either the HIV-1 RRE or the HIV-1 TAR RNA had been bound. As seen
in Fig. 3A, Rev binds efficiently to RRE-RNA, but not to the
TAR-RNA. A non-RNA-binding protein, HIV-1 Nef, was also tested with
both RNA chips as an additional control for nonspecific interactions.
As expected, Nef did not bind to these surfaces. Rev also did not bind
to the surface of flow cells lacking bound RRE-RNA. An "empty" flow
cell was used routinely as a background control (not shown).
The specificity of Rev-RRE interaction was also tested by competitive
inhibition of Rev binding. 50 nM Rev was preincubated with
increasing concentrations of either RRE/HH (nucleotides 50-113 of RRE
(Fig. 2A)) lacking the poly(A)
tail or yeast tRNA prior to injection over a sensor surface to which
RRE/T (nucleotides 39-214 of RRE) (Fig. 2A) had been
previously bound. A representative experiment is shown in Fig.
3B. Increasing the
concentration of free RRE/HH RNA significantly inhibited the binding of
Rev to the surface bound RNA, resulting in a maximal 75% inhibition of Rev binding to the surface bound RRE/T, whereas tRNA had no effect. Relatively high concentration of competitor RNA in solution was required to obtain significant inhibition because under the conditions used in SPR, the local concentration of immobilized RNA was higher than
that of the competitor RNA (1-2 µM RRE/HH in solution)
required for ~50% inhibition. Furthermore, RRE/HH bound to Rev in
solution was in equilibrium and could have dissociated during the
analysis, allowing Rev to bind to unoccupied sites on the surface bound RRE/T. In addition, RRE/T has four Rev binding sites, whereas RRE/HH
has only two (see below). RRE/HH was used as the competitor RNA here
because sufficiently high concentrations of the larger RNAs proved too
difficult to manipulate under these conditions.


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Fig. 2.
Structure and sequence of RNAs and
proteins. A, putative secondary structures of various
RRE RNAs used in this study. Full-length HIV-1 RRE, a 244-nucleotide
sequence used in this study, is shown; RRE/T is a truncation
mutant containing nucleotides 40-213 of the RRE; RRE/HH
denotes nucleotides 50-113 (stem/loops B/B1/B2); RRE/SLIIB
is composed of stem/loop B/B1 includes nucleotides 51-89 appended to
nucleotides 105-110; RRE/Z is RRE from which the stem/loops
B/B1/B2 (nucleotides 50-113) had been excised; and RRE-3A
is a mutant RRE in which three Gs (denoted by arrows) at
positions 56-58 had been exchanged for three As. B, amino
acid sequence of different Rev proteins described in the text. Wherever
appropriate, deleted residues are indicated by dashes.
REV T 87 refers to Rev protein truncated at the 87th
residue. REV 25-34/MS-C denotes a REV/MS2 coat protein
fusion containing a deletion of REV residues 25-34. The Rev sequence
with the deletion (denoted by dashes) stops at the
arrow, and italics denote the MS-2 coat protein
sequence beyond this point. REV 12-88 and REV
17-87 are synthetic peptides encompassing residues 12-88 and
17-87 respectively. Other peptides shown include REV
22-86 25-34, a REV peptide of residues 22-86 and containing a
deletion of residues 25-34; REV 22-86 53-66, sequence
22-86 with a deletion of residues 53-66; REV
22-86 24-34 53-66, sequence 22-86 with deletions of
residues 24-34 and 53-66; REV 22-40:DLRE:45-60, sequence
between residues 22 and 60 and containing a DLRE (underlined
italics) substitution for residues 41-44; and REV
22-40:KKKK:45-60, sequence between residues 22 and 60 and
containing a KKKK (underlined italics) substitution for
residues 41-44.
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Fig. 3.
Specificity of Rev-RRE binding and
competitive inhibition. A, overlay sensogram plots of
100 nM HIV-1 Rev or Nef proteins injected at 100 µl/min
over a sensor surface to which either HIV-1 RRE or TAR RNAs had been
bound. The arrow indicates the end of the association phase
and start of the dissociation phase. B, Rev interaction with
the RRE sensor in the presence of increasing amounts of RRE/HH RNA or
yeast tRNA in solution. Data are plotted as percentage of resonance
signal obtained with 100 nM Rev in the absence of any RNAs
in solution. .
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Equilibrium Binding and Stoichiometry of Rev-RRE
Interactions--
The stoichiometry of Rev-RRE binding was determined
by equilibrium binding protocols described under "Experimental
Procedures." Briefly, increasing concentrations of Rev was injected
at low flow rates for extended times (5 µl/min for 20 min) over a
sensor chip to which precisely known quantities of RNA had been bound. A typical example of sensogram series of Rev interacting with the
full-length RRE-RNA is shown in Fig.
4A. The total amount of bound
Rev was measured 5 s before the end of the injection/association phase (arrow), which was then followed by a short
dissociation phase. For clarity, the regeneration phase of each run has
been omitted in this figure. Because the amounts of bound RNA and
protein are both known, the ratio of protein:RNA can be calculated
using the conversion factors; 1 RU of RNA bound = 0.8 pg of
RNA/mm2 flow cell surface area, and 1 RU of protein
bound = 1 pg of protein/mm2 (65, 72,
73).2 Fig. 4B
shows the results of these analyses with different RRE-RNAs used in
this study. Each point on the different plots represents the average of
at least three independent runs using different batches of the
respective RNAs. As expected, Rev associated efficiently with RRE and
RRE/T RNAs, but the amount of Rev bound leveled off at approximately
four Rev monomers per molecule of RNA at about 350-400 nM
of injected Rev. However, at 400 nM Rev, a breakpoint was
reached at which the amount of Rev bound to the full-length RRE
increased rapidly to a ratio of more than 10:1 at 1 µM
injected Rev. This was also obvious from closer inspection of the
sensograms in Fig. 4A. Unlike the sensogram run at 400 nM Rev, in which a state of equilibrium was reached
relatively early in the injection phase, sensograms at Rev inputs
greater than 400 nM display an increase in the resonance
signal as additional Rev is bound throughout the time course of the
experiment. Runs at lower analyte (Rev) concentrations also have a
gently rising slope as the core (high) affinity sites were loaded; runs
at 50 or 100 nM Rev have, in addition, a rapidly increasing
signal early in the association phase as the core RRE site becomes
saturated. With a truncated RRE, RRE/T (Fig. 2A), which
lacked most of stem/loop A, no break point between two classes of
binding sites was evident (Fig. 4B). RRE/T RNA bound only
four Rev monomers. In contrast, two other RRE mutants, namely RRE/Z,
deleted for the core binding site, and RRE-3A, wherein the three Gs at
the core site were changed to three As (Fig. 2A), bound
little or no Rev at low concentrations of the protein (as expected from
their null phenotypes). However, with high Rev input; i.e.
300 nM for RRE/Z or 400 nM for RRE-3A), there
was significant protein binding. This suggested that although initial
nucleation of these mutant RNAs is inefficient, once the Rev monomer
bound to these RNAs, the protein could quickly accumulate, possibly
aided by protein-protein interactions. As the size of the RNA
decreased, the number of Rev monomers that can bind was also reduced.
Even at the highest concentrations of injected Rev, no more than one
Rev monomer was bound to the minimal RNA, RRE/SLIIB (Fig.
2A), whereas the RRE/HH bound two Rev monomers (Fig.
4B). Taken together, these results suggested the presence of
two classes of Rev binding sites on the RRE, which are designated as
high and low affinity binding sites. They also showed that protein interactions through the Rev oligomerization interphase were in and of
themselves insufficient to permit the accumulation of Rev on the sensor
surface unless a sufficiently large RNA target was available.

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Fig. 4.
Equilibrium binding of Rev to RRE and
calculated Rev-RRE stoichiometries. A, sensogram runs
at different Rev concentrations under conditions approaching
equilibrium (flow rate < 10 µl/min). The arrow
denotes the point at which injection was switched to dissociation
buffer. B, plot of relative stoichiometry of Rev per
immobilized RRE RNA species. Individual graphs represent the number of
Rev monomers bound to the respective RNAs plotted as a function of Rev
concentration (in nM). Each point (with error
bars) represents the average of three independent experiments with
different preparations of each RNA species.
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Rev-RRE Binding Kinetics--
The presence on RRE of both high and
low affinity binding sites for Rev had a significant effect on the
calculation of the kinetics of Rev-RRE interaction. As seen in Fig.
5, A and B, a very
close fit to the primary sensogram was calculated for Rev binding to
the truncated RNAs, RRE/SLIIB and RRE/HH, respectively, with the best
fit curves generated using the BIAevaluation 3.0 (Biacore Inc.)
software (heavy dashed lines) almost completely superimposable over the experimental plots. The association kinetics were not significantly affected by changes in the flow rate or density
of RNA on the sensor surface (data not shown), indicating that mass
transport limitations were not a factor under these experimental
conditions. However, for the larger RNAs, RRE/T (Fig. 5C)
and RRE (Fig. 5D), the calculated fits deviated
significantly from the experimental plots, thus contributing
significantly to errors in the determination of binding affinities of
Rev for these large RNAs. The presence of more than one on-rate for the
binding of Rev to the full-length RRE may account for these
discrepancies. Although not obvious in the primary sensograms,
transformation of the association phase of the sensogram revealed a
significant increase in the rate of Rev binding to RRE as the
concentration of Rev was progressively increased (Fig.
6A). Whether this resulted from increased accessibility of Rev to the low affinity binding sites
on RRE, reflected potential cooperativity in Rev binding (mediated
through the oligomerization domains of the protein), or a combination
of both could not be determined.

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Fig. 5.
Binding kinetics of Rev to target RNAs and
calculated best fits. Sensogram profiles of Rev binding to
RRE/SLIIB (A), RRE/HH (B), RRE/T (C),
and full-length RRE (D) RNA sensor surfaces. In each panel,
sensograms obtained with different Rev inputs (denoted inside each
panel) are overlaid. The best theoretical fit (denoted by the
heavy dashed line) for each sensogram was calculated using
BIAevaluation 3.0 software and is overlaid on each plot.
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Fig. 6.
Analysis of association and dissociation
phases of Rev binding to RRE. A, data points
corresponding to the association phase of each sensogram are presented
as derivative semi-log plots (ln(dRU/dT)) versus time (in
seconds). Individual plots correspond to results obtained for sensogram
runs with different Rev inputs. B, data points corresponding
to the dissociation phase of each sensogram are presented as semi-log
plots of [ln(RU0/RUn)] versus time.
Individual plots correspond to results obtained for sensogram runs with
different Rev inputs.
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Transformation of the data from the dissociation phase of the
sensograms representing Rev interaction with the larger RNAs also
revealed significant deviations in the observed off-rates (as indicated
by the slopes) that were more pronounced for sensograms generated at
high Rev inputs (Fig. 6B). The apparent initial rates of
dissociation of Rev from RRE increased significantly for all sensogram
runs performed at Rev concentrations exceeding 500 nM. Presumably, at these higher protein concentrations, the dissociation rates of the low affinity binding sites predominate. As a consequence, the estimates of the dissociation rate constants were made using data
generated with less than 400 nM injected Rev. However, even at these levels, two different off-rates could be calculated in which
the rate of dissociation from the low affinity site is 3 orders of
magnitude faster (~10
2 versus
~10
5, Table I) than that
seen for the high affinity site. The above considerations resulted in
some ambiguities in the calculated affinity constants for Rev
interaction with the larger RNAs, shown in Table I. Similar analyses of
sensograms of Rev interaction with the truncated targets, RRE/HH and
RRE/SLIIB, did not reveal such discrepancies in the off-rates at
different Rev inputs (data not shown). But experiments with the
truncated RNAs allowed us to calculate a binding affinity constant
(KD) of 4 × 10
11 (Table I) for
Rev interaction with the high affinity site with high confidence in
light of the close theoretical fits obtained for these interactions
(Fig. 5, A and B). The substantially lower affinities of Rev for RRE and RRE/T implied in these calculations are
most likely a reflection of the contributions of low affinity site
binding. The apparent association rate constants
(ka1 and
ka2) for the low and high affinity sites
are not too different, which may be due to the rapid association rate of Rev for the high affinity site masking the potential differences in
the association rate for the low affinity sites.
Mass transport of the analyte, in this case, Rev to the surface bound
ligand can, under certain conditions be rate-limiting. This may
introduce errors in the calculation of molecular on and off rates. To
determine whether Rev was mass transport limited, the Rev-RRE
interaction was examined at various flow rates and RNA surface
densities. No significant difference in the Rev-RRE binding kinetics
could be detected (data not shown), indicating that mass transport
limitations were not a factor under our experimental conditions.
Effect of Rev Mutations on RRE Binding--
As shown in Fig.
1A, Rev contains multiple functional domains, with the
extreme N and C termini considered nonessential to Rev function. To
test the effects of mutations in these functional regions on Rev-RRE
interactions, we injected a variety of Rev mutant proteins or peptides
over surfaces to which the different RNAs had been bound. As shown in
Fig. 7A, the C-terminal region of Rev is not essential for RNA binding because the termination mutant
RevT87 (Rev terminated at amino acid 87) bound RRE very efficiently.
This mutant protein behaved in all respects like the full-length Rev
(not shown). By contrast, even short deletions of the N-terminal
portion of Rev had a profound effect on the behavior of the protein.
The two peptides Rev12-88 and Rev 17-87 appear to bind very well to
the RRE loaded flow cell used here (see Fig. 7, C and
E, respectively). However, these peptides also bound almost
as well to the empty flow cell, which lacked even the
biotin-U(dT)18 linker (see Fig. 7, D and
F). This is in marked contrast to RevT87, which does not
bind to the empty flow cell (Fig. 7B). The deflection
observed in Fig. 7B is due to the increased refractive index
of the injected solutions that contain increasing concentrations of
protein, and not to specific binding. Proteins or peptides containing
deletion mutations in the oligomerization domains (M42
(Rev
25-34/MS-C), Rev22-86
25-34, and Rev22-86
53-66 or
Rev22-86
24-34
53-66) did not bind to any RNA used (data not shown). Similarly, peptides with mutations in the RNA binding domain
(e.g. 22-40:DLRE:45-60 or 22-40:KKKK:45-60, each
replacing four arginines at positions 41-44) also did not bind to any
of the RNAs used here (data not shown). This confirmed previous results (13, 20-25, 75) demonstrating the importance of this region of Rev to
stable RNA binding.

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Fig. 7.
Specificity of binding of Rev peptides to
sensor surfaces with or without immobilized RNAs. Sensograms
profiles of Rev protein truncated at residue 87 binding to RRE
(A) or a control chip (B); synthetic Rev peptide,
Rev12-88 of residues 12-88 with RRE (C) or control chip
(D); and synthetic Rev peptide, Rev17-87 with RRE
(E) or control chip (F). The different plots in
each panel represent sensogram runs at different Rev or peptide
concentrations.
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Evaluation of Inhibitors of Rev-RRE Interactions--
It has been
previously reported (76) that small molecules that bind RNA, such as
the aminoglycoside antibiotics (e.g. neomycin B), can
inhibit the binding of Rev to RRE. To investigate whether this method
could be used to rapidly evaluate this inhibitory reaction, Rev, at a
concentration of 100 nM, was incubated with increasing
concentrations of either neomycin B or hygromycin B prior to injection
over a sensor surface to which RRE/T had been bound. As seen in Fig.
8A, neomycin B efficiently
inhibited the binding of Rev to the RNA, with maximal inhibition
between 10 and 25 µM of the antibiotic. In contrast,
hygromycin B did not inhibit Rev binding at any concentration tested
(Fig. 8B). Kanamycin A and tetracycline were tested as well,
and neither one inhibited Rev binding (data not shown). These results
demonstrated that this technique could be used to screen rapidly and
conveniently the effectiveness of potential inhibitors of Rev-RRE and,
by extension, any RNA-protein interactions in real time.

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Fig. 8.
Inhibition of Rev-RRE interaction by
antibiotics. Sensogram overlay plots of 100 nM Rev
preincubated with increasing concentrations of neomycin B
(A) or hygromycin B (B) binding to RRE/T.
Specific concentrations of the indicated antibiotics are shown.
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DISCUSSION |
Using SPR to investigate RNA-protein interactions has a number of
advantages over traditional methods, such as EMSA (77) or filter
binding assay (78). The SPR technique provides kinetic data more
conveniently than other techniques and allows a rapid evaluation of
potential inhibitors of these interactions. Besides these obvious
advantages, the SPR method provides a more precise determination of
both initial velocity and steady-state parameters. In the EMSA and the
filter binding method, substantial time elapses before the RNA-protein
complexes are quantitated. As such, in a complex interaction involving
multiple binding sites on the RNA, the kinetic parameters evaluated by
these methods may reflect a pseudo-steady-state averaging of individual
interactions. This results in a wide distribution of RRE-Rev complexes
of different stoichiometries appearing as discrete bands in EMSA gels.
Whereas this pseudo-steady-state analysis may not be a problem if the low affinity interactions have fast off rates, the high levels of Rev
(µM versus nM in the SPR) used in
the EMSA may spuriously reinforce the low-affinity interactions through
nonspecific trapping of RNA by Rev oligomers. Filter binding assays in
general use a lower range of protein concentration and are therefore
comparable with the SPR assays. Although individual
kd for RRE-Rev interactions can be estimated fairly
well by filter binding assays, this method is inadequate for obtaining
directly, precise estimates of the chemical on rates
(ka), because even at low concentrations of Rev, the
system approaches equilibrium within 5 min (Fig. 4A).
The RNA capture technique described here resulted in a stable sensor
surface that could be used through multiple rounds of protein binding
but still allowed complete regeneration of the surface, permitting
reuse with different RNAs. Immobilizing the RNA at its 3'-end through a
rigid RNA-DNA hybrid presents the ligand in the same orientation,
unlike the situation when proteins or RNAs are bound directly to the
dextran surface of the chip. This is particularly crucial in systems
such as this, in which multiple Rev monomers bind to the RNA. As such,
it would be difficult, if not impossible to determine the stoichiometry
of Rev-RRE binding if Rev was constrained by being bound to the chip.
In fact, attempts to link Rev to the dextran matrix followed by
injection of soluble RRE were unsuccessful (data not shown).
One area of continued controversy involves the stoichiometry of Rev-RRE
interactions. Several studies have shown that between 8 and 10 Rev
molecules bind to the 244-nucleotide RRE in vitro (44, 45,
47, 50, 52, 53, 60) and Rev multimers on RRE may be necessary for
trans-activation (54, 61, 62). Mann et al. (59) have claimed
that the functional RRE sequence is substantially larger than
previously described and that complete trans-activation requires 10 or
more Rev molecules bound per RRE. Although our full-length RRE
(described as truncation 3 by Mann et al. (59)) was capable
of binding 10 or more Rev monomers (Fig. 4B), these high
ratios represent an extreme case with high Rev inputs. Our RRE RNA
bound Rev very efficiently, reaching a plateau of 4:1 (Rev-RRE) at
approximately 400 nM injected Rev, and showed significant
activity in vivo. On the other hand, both RRE-3A and RRE/Z,
mutant RNAs that inactivate or delete the core binding site for Rev
(Fig. 2A), bound little or no Rev at low concentrations of
the injected protein, but at higher concentrations (300-400
nM), Rev accumulates on these RNAs. Because both of these mutant RNAs are inactive in vivo (47, 56), we feel that the Rev binding to the mutant RNAs seen in SPR assay is directed to the low
affinity sites that may be unavailable under conditions that exist in
infected cells. In other words, although it is possible to force the
RNAs to bind large amounts of Rev in vitro, the in vivo relevance of these results is questionable.
Another difference between these results and those of Mann et
al. (59) is that they only detected a 2:1 Rev:RNA ratio with their
truncation 5 mutant, which is a rough equivalent of RRE/T species
described in this manuscript. We obtained a 4:1 Rev:RNA ratio for RRE/T
RNA with binding kinetics essentially identical to that of Rev binding
the full-length RRE. A maximal 2:1 ratio was obtained in our assay only
with a considerably smaller target RNA such as RRE/HH (stem/loops
B/B1/B2). Reducing the size further (RRE/SLIIB) yielded an RNA molecule
that bound Rev at a 1:1 ratio and still retained a somewhat reduced
in vivo Rev response (54, 56, 58). Kinetic analysis (see
below) confirmed that this was the core high affinity Rev binding site.
There has been some dispute regarding whether Rev binds the RRE as a
monomer or whether protein multimerization is a prerequisite for RNA
binding. Although the kinetics presented here cannot settle this
dispute, the fact that the minimal high affinity binding site
(RRE/SLIIB) bound Rev at a 1:1 ratio and the slightly larger RRE/HH
yields a 2:1 protein:RNA ratio suggested that the initial binding to
RRE must involve a Rev monomer, followed by multimerization in
situ, in agreement with previous reports (10, 58, 81, 82). It has
also been suggested that a critical threshold level of Rev on RRE is
required for trans-activation (59, 61, 79, 80) and also that the RRE
acts as a molecular rheostat (59) regulating the sequential addition of
Rev monomers to achieve the critical threshold. Although the results
presented here tend to support these suggestions, we disagree with some
of the details. As discussed above, the number of Rev monomers bound to
each RRE is unlikely to be more than four in vivo. EMSA of
Rev binding to stem II RNA (RRE/HH equivalent) suggested that Rev bound
this RNA with a stoichiometry of 3:1 (53). Because RRE/HH preserved only 50% of the in vivo activity obtained with full-length
RRE (54, 56), it may be reasoned that four Rev monomers have to be
bound to the target for maximal Rev response in vivo. In
point of fact, full Rev response was obtained with RNA targets
containing a RRE/HH dimer (54, 56), which had a potential to bind four Rev monomers.
A precise determination of Rev binding kinetics was made difficult by
the contribution of both high and low affinity binding sites on the RNA
and by the presence in Rev of oligomerization domains that may result
in cooperative RNA binding. These limitations underlay the inability of
the BIAevaluation 3.0 program to provide a good theoretical fit to the
sensograms of Rev binding to RRE and RRE/T, whereas excellent fits were
possible when the truncated RNA targets RRE/HH and RRE/SLIIB were used.
Use of these shorter RNAs reduced or eliminated the contributions of
the low affinity binding sites in the larger RNAs and probably
precluded the contributions by Rev oligomer binding. Both RRE/HH and
RRE/SLIIB showed affinities for Rev on the order of 4 × 10
11 M despite being calculated using
different models (double reciprocal plots for RRE/HH and single site
model for RRE/SLIIB). The difference in the affinity constants for RRE
and RRE/T compared with those for RRE/HH and RRE/SLIIB is undoubtedly
due to the residual contributions of the low affinity sites in the
larger RNAs in the calculation of the koff
functions (Table I, low affinity versus high affinity kd values). The low affinity binding constant,
KD, differed by 3-4 orders of magnitude from the
high affinity counterpart. The contribution of the low affinity sites
is shown by transformation of the dissociation phase of Rev binding
(Fig. 6B). The dissociation rates increase substantially at
higher concentrations of injected Rev, when the contribution of the low
affinity sites might be expected to predominate. Furthermore, the
KD values for the RRE and RRE/T, representing the
contribution of the high affinity binding site in the double reciprocal
plots, imply affinities of Rev for these larger RNAs to be at least an
order of magnitude more than the previous estimates (44-47, 49,
81).
Although the calculated association rates of Rev did not vary
substantially no matter which RNA was used (Table I), transformation of
these data showed changes in the rates of association with increasing
concentrations of Rev (Fig. 6A). As the Rev concentration increased, so did the apparent association rate. This phenomenon was
not observed with the RRE/SLIIB (data not shown). We interpret these
results to be a consequence of cooperative binding of Rev to the RRE,
mediated by Rev oligomerization. Because RRE/SLIIB can only bind one
molecule of Rev, cooperative binding was not an issue. Furthermore,
because Rev bound with high affinity to RRE/SLIIB, this showed that
high affinity binding is possible in the absence of multimerization, in
agreement with the suggestion of Daly et al. (81, 82).
It has been demonstrated that the C-terminal residues beyond amino acid
87 were dispensable for Rev function (60, 74), and consistent with
this, the Rev termination mutant (RevT87) showed good discrimination
between RRE/T and the unsubstituted flow cell (see Fig. 7, A
and B). In addition, this protein behaved identically to the
full-length Rev in all the kinetic measurements (data not shown). This
was in contrast to an earlier report that showed by EMSA that residues
near the C terminus of Rev beyond position 91 might be required for
oligomerization of Rev on the RNA (81). Other studies (63, 82) have
suggested that the N-terminal domain of HIV-1 Rev was required for the
discrimination between specific and nonspecific RNA binding. The
results presented in Fig. 7 seem to support this, but possibly not for
the reasons cited by Daly et al. (82). Peptides in which the
first 11 or 16 amino acids had been deleted do bind to the sensor
surface to which RRE/T had been bound (Fig. 7, C and
E, respectively). However, these peptides also had
considerable affinity to a flow cell to which neither RNA nor biotin
U-(dT)18 linker had been bound (Fig. 7, D and
F). The differences in amplitude between the unsubstituted
flow cell and the one containing RRE/T could be attributed to at least
two factors: either 1) these peptides retain some residual specificity
for the RNA over the dextran matrix contained in the flow cell, or 2)
the underivatized chip has a larger number of charged sites available
for protein binding, or a combination of the two. Even if the observed
differences in the sensograms of peptide binding to the RRE
versus control chip (Fig. 7, C versus
D and E versus F) reflected
binding specificity, the rapid dissociation of peptides from the sensor
surface precluded derivation of kinetic parameters, reminiscent of the
situation when the steady state binding of Rev peptides to RRE was
evaluated by EMSA or filter binding assay (82). Although the in
vitro peptide binding appeared to be largely nonspecific, Rev
domain peptides encompassing residues 17-87 or residues 23-87 were
capable of trans-activating RRE RNA targets in vivo but not
the mutated RRE counterpart lacking the core
site.3 Interestingly, these
peptides also activated HTLV-I RexRE RNA and a RRE chimera that had
substituted the core site for the MS2 phage translational operator
RNA.3 The differences in the in vitro and
in vivo behavior of the peptides may be reconciled if it is
presumed that the RRE binding by the Rev domain peptide(s) reflects a
somewhat nonspecific lower order electrostatic interaction between the
arginine forks in the peptide and the phosphate backbone of RNA,
similar to the initial "searching and discriminatory" process in
the case of Rev as it finally docks on to the active site on RRE RNA
through higher order interactions. This idea is bolstered by
experiments in which peptides with mutations in the RNA binding domain
that replace several arginine residues (e.g.
22-40:DLRE:45-60 or 22-40:KKKK:45-60) do not bind to any surface
(data not shown), even though these short peptides are largely
unstructured and carry a net positive charge. In vivo, even
the lower order interaction between authentic Rev domain peptides and
RNA may be sufficiently bolstered by ancillary cellular factors to
enable trans-activation in the absence of stringent RNA sequence specificity.
Finally, it has been reported (76) that certain antibiotics can
selectively inhibit Rev-RRE interactions. Neomycin B completely inhibited Rev binding to RRE/T (Fig. 8A), whereas hygromycin
B (Fig. 8B), tetracycline, or kanamycin A (data not shown)
had no inhibitory effects. The maximum inhibitory effect occurred
between 10 and 25 µM neomycin B co-injected with Rev,
whereas Zapp et al. (76) reported that 1 µM
neomycin B produced maximal effects. These differences could be
attributed to the fact that in the SPR assay, approximately 12-fold
more Rev was used than was the case in gel-shift assay of Zapp et
al. (76) (16 ng of Rev versus 1.3 ng). A high Rev
concentration was necessary in our case to obtain a good signal in the
absence of neomycin. Furthermore, in the kinetic system of the
biosensor, Rev and neomycin B are competing for the same binding site
(76), and if Rev binds more tightly than the antibiotic to this site, a
higher concentration of the antibiotic might be required to obtain
maximal inhibition. Nonetheless, these results demonstrate the
usefulness of this system to rapidly evaluating potential inhibitors of
Rev-RRE interactions.
RNA-protein interactions are a crucial element in cellular metabolism.
Although a number of methods are available to investigate these
processes, most are tedious and time consuming. The technique outlined
here provides a more convenient means of assessing binding kinetics,
stoichiometry, specificity, and potential inhibitors of protein-RNA
binding in real time, and as such provides a significant advance over
the more traditional methods, even in a complex system such as that
seen for the HIV-1 Rev and its target, the RRE.