(Received for publication, February 26, 1997)
From the Department of Antiviral Research, Merck Research Laboratories, West Point, Pennsylvania 19486
Ribonuclease L (RNase L) is an endoribonuclease
that is activated upon binding of adenosine oligomers linked 2 to 5
to cleave viral and cellular RNAs. We recently proposed a model for
activation in which activator A binds to monomer, E, to
form EA, which subsequently dimerizes to the active form,
E2A2 (Cole, J. L., Carroll,
S. S., and Kuo, L. C. (1996) J. Biol. Chem. 271, 3978-3981). Here, we have employed this model to define the
equilibrium constants for activator binding (Ka)
and dimerization of EA to
E2A2 (Kd) by
equilibrium analytical ultracentrifugation and fluorescence
measurements. Multi-wavelength sedimentation data were globally fit to
the model above, yielding values of Ka = 1.69 µM and Kd = 17.8 nM for
2
,5
-linked adenosine trimer. Fluorescent conjugates of 2
,5
-linked
adenosine trimer with 7-hydroxycoumarin have been prepared. The
coumarin emission anisotropy shows a large increases upon binding to
RNase L. Analysis of anisotropy titrations yields values of
Ka and Kd close to those
obtained by sedimentation. The sedimentation parameters for unmodified
2
,5
-linked adenosine trimer also agree with those obtained by enzyme
kinetic methods (Carroll, S. S., Cole, J. L., Viscount, T., Geib, J.,
Gehman, J., and Kuo, L. C. (1997) J. Biol. Chem. 272, 19193-19198). Thus, the data presented here clearly define the
energetics of RNase L activation and support the minimal activation
model.
RNase L is an endoribonuclease involved in the interferon pathway.
Upon activation by adenosine oligomers linked 2 to 5
, RNase L
degrades viral and cellular mRNAs, leading to an inhibition of
protein synthesis in virally infected cells (1, 2). It has been
demonstrated by gel filtration, chemical cross-linking (3), and
analytical ultracentrifugation (4) that RNase L exists as a monomer in
solution but is dimerized in the presence of an activator, suggesting
that the catalytically active form of RNase L is a homodimer. Under
stoichiometric binding conditions, dimerization and activation of RNase
L require the binding of one activator molecule per ribonuclease L
monomer (4). However, in the absence of an activator, only monomer is
observed up to a protein concentration of at least 18 µM,
indicating that unliganded enzyme is unable to dimerize, or the
association is extremely weak. These observations support a minimal
model for RNase L activation depicted in Scheme 1. In
the context of this model, the energetics of activation of RNase L are
defined by two equilibrium dissociation constants,
Ka and Kd. Note that this model
is a subset of the full reaction scheme for a coupled ligand binding and enzyme dimerization equilibrium (5, 6). We have also extended this
model to include the substrate binding step and employed it to compare
the activation of RNase L by analogs of the native activator (7).
Scheme 1.
Under the conditions of our previous sedimentation equilibrium
measurements (4 °C), the enzyme concentrations were much higher than
either Ka or Kd. Thus, the
stoichiometry of the interaction was defined, but the association
constants were inaccessible. However, activator binding is highly
temperature-dependent, such that at 20 °C association is
weaker and directly accessible by biophysical measurements for several
activators containing a 5-OH group. In the present study, we have
characterized the binding of several activators with both
sedimentation equilibrium and fluorescence anisotropy
measurements. The results are compared with those obtained by enzyme
kinetic methods in the accompanying paper (7).
Human RNase L was expressed and purified as described previously
(8) and stored in 40% glycerol, 25 mM HEPES, pH 7.5, 100 mM KCl, 5.8 mM MgCl2, and 5 mM DTT.1 To reduce the UV
absorbance due to oxidized DTT, the sample buffer (11 mM
HEPES, pH 7.5, 104 mM KCl, 5.8 mM
MgCl2) was purged of oxygen by bubbling with argon prior to
adding 2 mM DTT. The enzyme was equilibrated into the
sample buffer using Bio-Rad Biospin 6 spin columns. Protein
concentration was measured spectrophotometrically using a molar
extinction coefficient at 280 nm of 8.41 ± 0.87 × 104 M1 cm
1 (4).
The 2,5
-linked adenosine trimer (HO-2
,5
-A3) was
obtained from the Midland Certified Reagent Company or from Sigma. The activators containing a free 2
-amino group,
HO-2
,5
-A3-2
-NH2 and
p-2
,5
-A3-2
-NH2, were obtained from NAPS
GmbH, Goettingen, Germany. The oligoribonucleotide concentrations were
determined spectrophotometrically using
260 nm = 1.53 × 104 M
1
cm
1/adenosine. The succinimidyl ester of
7-hydroxycoumarin-3-carboxylic acid (NHS-7HC) was obtained from
Molecular Probes.
Fluorescence activators were prepared by reacting NHS-7HC with
HO-2,5
-A3-2
-NH2 or
p-2
,5
-A3-2
-NH2. NHS-7HC was dissolved in
N,N-dimethylformamide at a concentration of 5 mM. The conjugation reactions were performed in 50 mM sodium phosphate, pH 7.65, at a concentration of 20 µM oligoribonucleotide, 200 µM NHS-7HC at room temperature for 2 h. The conjugate was purified on reverse phase HPLC using a Vydac C18 column with a linear gradient
of acetonitrile (0.5-30% in 30 min, flow rate = 1 ml/min) in 50 mM aqueous triethylammonium acetate, pH 6.8. Representative elution times are: NHS-7HC, 12.95 min;
HO-2
,5
-A3-2
-NH2, 10.13 min; HO-2
,5
-A3-7HC, 15.92 min.
Equilibrium analytical ultracentrifugation were performed using six-channel (1.2-cm path) charcoal-Epon cells with a Beckman XL-A centrifuge and an An-60 Ti rotor at a speed of 14,000 rpm and a temperature of 20 °C unless otherwise indicated. Samples of 110-µl volume were loaded under argon. Scans were recorded at 230 and 260 nm using 0.001-cm point spacing and averaging 10 readings at each point. Equilibrium was judged to be achieved by the absence of systematic deviations in a plot of the difference between successive scans taken 4 h apart.
For fluorescence experiments, 120-µl samples containing fluorescent
activator and RNase L were prepared in sample buffer and allowed to
equilibrate at least 2 h prior to measurement. Data were collected
at 20 °C in 10 × 2-mm Hellma microcuvettes using an ISS K2
fluorometer at an excitation wavelength of 404 (16 nm bandpass), and
emission was collected using a Schott KV-470 long pass glass filter.
Steady-state anisotropy was measured in the L-format, and
instrument G-factors were measured with the excitation polarizer in the
horizontal position. Data for both the sample and buffer blank were
collected for 2 s in each orientation, and at least three
measurements were averaged. Fluorescence lifetime measurements were
performed using the phase modulation method over a frequency range of
5-190 MHz. The lifetime reference was Me2POPOP in ethanol,
( = 1.45 ns) (9).
Sedimentation equilibrium data were globally fit using the Marquardt-Levenberg least squares algorithm with the SAS software package (SAS Institute, Cary, NC). Within the context of Scheme 1, the concentrations of EA and E2A2 can be expressed in terms of the dissociation constants Ka and Kd and the concentrations of E and A.
![]() |
(Eq. 1) |
![]() |
(Eq. 2) |
Our method to determine the dissociation constants relies on the differences in the absorption spectra of E and A and is similar to the approach described by Lewis and co-workers (10) to characterize protein-DNA interactions. We can express the absorbance at any wavelength as the sum of the contributions from each of the species participating in the equilibrium
![]() |
(Eq. 3) |
![]() |
(Eq. 4) |
![]() |
![]() |
(Eq. 5) |
The value of E was experimentally measured by
sedimentation equilibrium of RNase L alone at 14,000 rpm, and
A was obtained for each activator by sedimentation
equilibrium at 45,000 rpm. Because of the low molecular weight of the
activators,
260 could not be obtained by overspeeding
following the run. Therefore, we used a mass conservation method in
which
260 was measured as the difference in the
integrated absorption at 260 nm at equilibrium at 14,000 rpm and the
absorbance at 260 nm at the beginning of the run. Because the
activators do not contribute much absorbance at 230,
230
was measured by overspeeding to 45,000 rpm following the centrifugation
experiment at 14,000 rpm. For each experiment three channels containing
different activator concentrations were analyzed globally at 230 and
260 nm where C0A and
C0E were treated as local
fitting parameters, and Ka and Kd
were global parameters.
The fluorescence anisotropy titration data were fit using the Marquardt-Levenberg algorithm within the IGOR Pro software package (Wavemetrics, Inc.). From Equations 1 and 2 and mass conservation it can be shown that
![]() |
(Eq. 6) |
![]() |
(Eq. 7) |
The titrations of RNase L were performed at four activator
concentrations under conditions where [A]T 5[E]T. The value of R was take to
be 0.40 based on the limiting intensity decrease at high enzyme
concentration. The value of rf was measured
experimentally.
For both the sedimentation and fluorescence experiments the joint confidence intervals for the fitted values of Ka and Kd were obtained by a search of the error surface in which one parameter was fixed, and the all the other parameters were allowed to adjust to their best fit values. This method correctly accounts for the cross-correlation (nonorthogonality) of the fitting parameters and the nonlinearity of the fitting equation (11, 12).
Our previous studies of
activator-induced dimerization of RNase L by sedimentation equilibrium
were performed at 4 °C, using conditions where [E]
Ka and Kd, so that the stoichiometry could be characterized (4). In contrast, for the weaker
activators, such as HO-2
,5
-A3, activator-induced dimerization is not stoichiometric at the higher temperature of 20 °C.2 Thus, sedimentation equilibrium
data obtained under these conditions may be used to extract
Ka and Kd. The radial absorbance
profiles contain contributions from each of the species participating
in the equilibrium: E, A, EA, and
E2A2; these data can be fit to
Equation 5 to obtain Ka and Kd. To define the contributions of RNase L and activator at each radial position we have taken advantage of differences in their absorption spectra. Fig. 1 shows that at enzyme and activator
concentrations typically used in our experiments the 230 nm absorbance
is dominated by the contribution from the enzyme, whereas the 260 nm
absorbance is dominated by the activator. Thus, we record data for each
channel at both of these wavelengths and fit using a single value of
C0A and
C0E. To further constrain
the fits, data can be obtained at several concentrations of enzyme and
activator. We have found that three activator concentrations are
sufficient to define Ka and Kd
for the activators used in this study.
Fig. 2 shows the sedimentation equilibrium data obtained
at 230 and 260 nm for samples loaded at a concentration of 0.5 µM RNase L and 1, 3, and 10 µM
HO-2,5
-A3. The data have been fit to Equation 5; the
solid lines are the best fit to the data, the insets are the residuals, and the results are summarized in
Table I. There is a weak positive systematic deviation
in the residuals at 230 nm for the data obtained at 1 and 10 µM and a negative deviation for the data obtained at 3 µM. These deviations are likely due to errors in the
baseline offset term
230. However, outside of these
deviations the fit is quite good. Although baseline errors result in
higher r.m.s. deviations, in the range of 0-0.02 OD they do not affect
the fitted values of Ka or Kd significantly as defined by the 1 S.D. confidence
intervals.2
|
Fig. 3 shows a contour plot of the fit error surface for
the data in Fig. 1 as a function of lnKa and
lnKd. The values of lnKa and
lnKd were fixed, and the other parameters were
allowed to adjust to the best fit values. The inner contour is drawn at
1 S.D., and the outer contour is at 2 S.D. joint confidence intervals.
The shape of the contours is typical for a system exhibiting parameter
cross-correlation (12). In our case, there is negative correlation
between lnKa and lnKd. Despite
this correlation, the system exhibits a single, well defined fit
minimum. As shown in Table I, the 1 S.D. joint confidence intervals
correspond to about a 2-fold change in either Ka or
Kd.
The validity of this fitting procedure was also verified using
simulated data. Extinction coefficients and reduced molecular weights
were fixed at the experimental values for RNase L and HO-2,5
-A3. Data were simulated according to Scheme 1 with
a loading concentration of 0.5 µM enzyme and 1, 3, and 10 µM activator. Random Gaussian noise of amplitude 0.0062 r.m.s. was added to corresponding to typical data obtained with our
XL-A centrifuge. With values of Ka and
Kd fixed at the experimental determined values from
Table I, the fitting procedure recovers the correct values with 1 S.D.
joint confidence intervals of 1.31-1.89 µM for
Ka and 15.1-21.8 nM for
Kd. The slightly greater width of the experimental
confidence intervals presumably reflects the contributions of small
systematic errors in the fixed parameters.
Table I also shows values of Ka and Kd and r.m.s. errors obtained from fitting the same sedimentation data to an alternative association model in which E dimerizes to form E2, which is then competent to bind two activator molecules to form E2A2 (Scheme 2). In this case, Ka is found to be 7 orders of magnitude lower than in the previous fit, and Kd is significantly higher. The r.m.s. deviation is also elevated relative to the previous fit.
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![]() |
To compare directly the sedimentation equilibrium and fluorescence
anisotropy methods, we characterized one of the fluorescent activators,
HO-2,5
-A3-2
-7HC, using the sedimentation equilibrium protocol. Initial measurements indicated that Kd for the activator is significantly lower than for HO-2
,5
-A3.
Therefore we have obtained sedimentation equilibrium data at a lower
enzyme concentration of 0.2 µM and three activator
concentrations of 0.2, 0.6, and 2 µM. As shown in Table
I, the best fit parameters for HO-2
,5
-A3-2
-7HC are
Ka = 2.67 µM and Kd = 1.72 nM. The joint confidence intervals for this fit are
much broader than for HO-2
,5
-A3, so that the change in
Ka relative to HO-2
,5
-A3 is not
statistically significant. However, the decrease in
Kd is significant. As in the case of HO-2
,5
-A3, simulations were performed with the
experimentally derived parameters and Gaussian noise of 0.0062 OD
r.m.s. amplitude. Again, the correct values of Ka
and Kd are recovered from the fitting procedure with
joint confidence intervals of 1.50-8.62 µM for
Ka and 0.19-4.35 nM for
Kd. Thus, the increased breadth in the confidence
intervals for the fit of HO-2
,5
-A3-2
-7HC are also
reflected in the simulations.
A fluorescence activator binding
assay was developed to characterize independently the equilibria
depicted in Scheme 1. Fluorescent activators, synthesized from
2,5
-oligoadenylate precursors containing a single 2
-NH2
moiety, were prepared by standard amine coupling reactions. We found
that activator conjugates with the NHS ester of NHS-7HC exhibited very
useful fluorescence properties. Coupling reactions were characterized
by HPLC and spectrophotometry. The product was identified based on the
appearance of an HPLC peak with retention time different from either
unconjugated oligonucleotide or the reactive fluorophore. In all cases,
a single predominant product was formed in the reaction. No new HPLC
peaks were formed in a reaction of NHS-7HC with HO-2
,5
-A3
lacking a 2
-NH2 group. Thus, the chemistry is specific for
the 2
-NH2 group. The identity of the product was confirmed
by absorption spectrophotometry. In aqueous solution, the NHS-7HC
exhibits one predominant absorption band at 415 nm without appreciable
absorption near 260 nm. In contrast, the product
p-2
,5
-A3-7HC exhibits two absorption bands at 405 and 260 nm. Thus, the absorption spectral features are consistent with the
proposed structure.
The fluorescence emission parameters of p-2,5
-A3-7HC are
summarized in Table II. The emission maximum is at 446 nm (uncorrected). Addition of a slight molar excess of RNase L results
in about 60% quenching in the fluorescence amplitude and a slight
shift in the emission maximum to 451 nm. A dramatic effect is observed in steady-state fluorescence anisotropy, which increases from 0.05 to
0.33 upon binding RNase L. Further addition of RNase L does not result
in appreciable increase in the fluorescence changes. Table II also
shows that the fluorescence lifetime is 3.5 ns in the absence and 3.4 ns in the presence of RNase L. Because there is no marked change in the
lifetime, the observed increase in anisotropy of
p-2
,5
-A3-7HC observed upon binding to RNase L is clearly
due to an increase in the rotational correlation
time.3 Similar spectral perturbations and
anisotropy increases are observed for HO-2
,5
-A3-7HC,
which lacks a 5
phosphate.4 Because the
anisotropy increase is very large upon binding, we have chosen to use
this parameter for quantitative characterization of the dissociation
constants.
|
The interaction of p-2,5
-A3-7HC with RNase L is too
strong to be readily characterized by fluorescence anisotropy
experiments, where the lowest accessible fluorophore concentration is
about 2 nM. However, the values of Ka
and Kd for HO-2
,5
-A3-7HC, which lacks
a 5
-phosphate, are accessible to fluorescence measurements. Fig.
4 shows a series of RNase L titrations performed at four activator concentrations ranging from 5 to 50 nM. At each
activator concentration the anisotropy shows a pseudo-hyperbolic
increase with increasing enzyme concentration. The plateau value of the anisotropy observed at the highest enzyme concentration of 2 µM decreases with decreasing activator concentration. In
contrast, the half-maximum point is not strongly dependent on activator concentration. We have globally fit these data to Equations 6 and 7. To
reduce the number of floating parameters it is necessary to constrain
the value of the emission anisotropy of EA,
rEA. The two limiting
cases we have considered are
rEA = rf and
rEA = rb. Fig. 4 shows a fit of the data assuming
rEA = rf, and the fitted parameters are in Table
III. The fitted curves overlay the data quite well
(r.m.s. deviation of 0.0131), and the best fit values of
Ka = 1.32 µM and Kd = 1.09 nM agree well with those determined by
sedimentation; however, the joint confidence intervals are much
narrower for the fluorescence results. A much poorer fit is obtained
assuming rEA = rb (r.m.s. = 0.0229), and the values of
Ka = 44.1 µM and Kd
near 1 pM are not compatible with the sedimentation data.
We have also considered the intermediate model in which
rEA = rb/2. The quality of this fit is intermediate
between the two former cases (r.m.s. = 0.0204), and the values of
Ka and Kd are close to that
determined assuming rEA = rf. In all of these cases we have also assumed
that the 60% quenching only occurs upon dimerization of EA
to form E2A2. However, the values of
the equilibrium constants and the fit quality are not very sensitive to
the extent of quenching of EA. We conclude that the simplest
model capable of fitting the data is that the anisotropy of
HO-2
,5
-A3-7HC is not strongly changed upon complexation
to EA but increases dramatically upon dimerization of
EA.
|
The sedimentation equilibrium and fluorescence data presented here
define the energetics of RNase L activation by the 2,5
-linked adenosine trimers HO-2
,5
-A3 and
HO-2
,5
-A3-7HC and provide support for the minimal
activation model presented in Scheme 1. For these activators,
Ka is in the low micromolar range, and
Kd is in the low nanomolar range. The value of
Kd is decreased by about 10-fold for the latter
activator relative to HO-2
,5
-A3. Despite the correlation
of the Ka and Kd parameters in
fitting the sedimentation and fluorescence data, global analysis of
data obtained over a range of activator and/or enzyme concentrations
gives rise to well defined fit minima and joint confidence intervals.
For HO-2
,5
-A3-7HC, the lower value of
Kd gives rise to broad confidence intervals for the
sedimentation fitting parameters. However, the confidence intervals for
the anisotropy parameters are quite narrow, and there is good agreement
between the best fit values of Ka and
Kd for this activator determined by the two
independent methods.
The values of Ka and Kd obtained
by sedimentation equilibrium may also be compared with those determined by kinetic methods (7). Using a model that explicitly accounts for
substrate binding, we have found that for a given activator, Ka and Kd are dependent on the
identity of the substrate: Ka is about 3-fold higher
for C11U2C7 than for
C11UC8, whereas Kd is about
5-fold lower. The parameters obtained by sedimentation measurements for
HO-2,5
-A3 agree more closely with the kinetic data
obtained with poorer substrate C11UC8, such
that Ka and Kd determined by the
two methods are within a factor of 2.
The sedimentation equilibrium data are also capable of distinguishing the activation model presented in Scheme 1 from the alternative model in Scheme 2. In the latter case, the enzyme monomer is capable of dimerization prior to binding activator. Although both models are consistent with our earlier stoichiometry measurements, the absence of any detectable dimer in the absence of activator at enzyme concentrations up to at least 18 µM strongly suggested that activator binding precedes dimerization (4). In the present study, the data fit well to the model in Scheme 1, but the large increase in r.m.s. for the alternative model (Scheme 2) indicates that it does not describe the data adequately. Most importantly, the deduced value of Kd = 138 nM is not consistent with the absence of measurable dimerization of free enzyme at micromolar concentrations. Thus, Scheme 1 represents the minimal model capable of describing the sedimentation equilibrium data.
The fluorescence anisotropy data are also consistent with the model in Scheme 1, but they cannot easily be used to distinguish different models because of the need to constrain the value of rEA. The binding data obtained at several activator concentrations fit well to Scheme 1 if it is assumed that the anisotropy increase only occurs upon dimerization of EA and fit less well to models that invoke an increase in anisotropy upon binding of A to E to form EA. This observation is surprising, since one might expect that the mobility of the fluorophore would be at least partially restricted upon binding, especially given the short linker between the oligoadenylate moiety and the fluorophore. It is likely that some anisotropy increase upon formation of EA does actually occur but is masked by the very large increase observed upon dimerization to E2A2. In any case, the fitted values of Ka and Kd are not strongly dependent on the value of rEA.
It is noteworthy that RNase L is monomeric at protein concentrations up to at least 18 µM, whereas in the presence of activator the Kd for dimerization is in the nanomolar range. Thus, activator binding increases dimerization affinity by a factor of at least 105-106, corresponding to strong thermodynamic linkage. The structural basis of this linkage is not yet clear. The activator binding site may be distant from the dimerization interface and modulate dimerization via a long range conformational change. Alternatively, activator binding may occur at or near the dimerization interface and directly modify the interactions that govern dimerization. There is precedence for strong linkage between dimerization and ligand binding in the Escherichia coli Rep helicase system. In the absence of DNA, Rep exists as a monomer at concentrations of at least 8 µM (13). However, dimerization is induced upon binding DNA with a Kd of about 5 nM, indicating an enhancement of dimerization of at least 104-fold (14).
The strong coupling of enzyme dimerization to activator binding
presumably represents a physiological mechanism to control RNase L
activity. The enzyme kinetic data suggest that monomeric RNase L is
inactive (7), and the ability of various oligoadenylates to activate
RNase L correlates with enzyme dimerization (3). Synthesis of
2,5
-linked oligoadenylates is performed by a family of
double-stranded RNA-activated synthetases which are induced by
interferon treatment (15). Although there are low, basal levels of
RNase L expression in most mammalian cells (less than one part in
500,000 of the protein in mouse liver), it is also induced during
interferon treatment (16). According to Scheme 1, formation of
E2A2, and therefore the specific
activity of RNase L, is dependent on the concentrations of both
E and A. Coordinate induction of 2
,5
-linked oligoadenylate
synthetases and RNase L by interferon would serve to prime the system
for a high level of RNase L activity upon activation of the synthetases
by double-stranded RNA.
We thank Fritz Benseler for suggesting the
use of a 2-amino group for synthesis of fluorescent activators and
Marc Lewis for help with the mathematical formalism for the analysis of
heterogeneous associations.