From the Division of Bioengineering and Physical
Science, ORS, OD, ¶ Laboratory of Biochemistry and
Genetics, NIDDK, and § Laboratory of Infectious
Diseases, NIAID, National Institutes of Health,
Bethesda, Maryland 20892
Received for publication, October 16, 2000, and in revised form, November 29, 2000
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The nonstructural protein NSP2 is a component of
the rotavirus replication machinery and binds single-stranded RNA
cooperatively, with high affinity, and independent of sequence.
Recently, NSP2 has been shown to form multimers and to possess an
NTPase activity, but its precise function remains unclear. In the
present study, we have characterized the solution structure of
recombinant NSP2 by velocity and equilibrium ultracentrifugation,
dynamic light scattering, and circular dichroism spectroscopy. We found
that NSP2 exists as an octamer, which is functional in the binding of
RNA and ADP. In the presence of magnesium, a partial dissociation of
the octamer into smaller oligomers was observed. This was reversed by
binding of ADP and RNA. We observed an increased sedimentation rate in
the presence of ADP and a nonhydrolyzable ATP analogue, which suggests
a change toward a significantly more compact octameric conformation.
The secondary structure of NSP2 showed a high fraction of Rotavirus is a significant cause of disease in humans and animals.
It is a member of the Reoviridae, and its genome consists of
11 segments of double-stranded RNA, which codes for six structural and
six nonstructural proteins. The structural proteins include VP2, VP6,
and VP7, which form the triple layered icosahedral virion capsid, the
spike protein VP4, the RNA polymerase VP1, and the multifunctional
capping enzyme VP3 (1) (for a review, see Ref. 2). Many studies have
addressed the properties of the structural proteins, for example their
spatial configuration in the virus particle using cryo-EM (3, 4), their
antigenicity, and their role in viral entry, replication, or
morphogenesis. Unfortunately, much less is known about the
nonstructural proteins that are expressed and left behind in the
infected cells.
Although it has been shown that these nonstructural proteins are not
essential for replicase activity in vitro (5), they are
important in several aspects of the replication cycle of the virus
in vivo. Some of the more intensively studied nonstructural proteins include, for example, NSP4, which has a membrane-destabilizing activity and assists in the budding of newly synthesized inner capsid
particles into the lumen of the endoplasmic reticulum (6, 7), where
they acquire the outer coat protein VP7. NSP3 binds to the 3'-end of
the viral mRNA and interacts with eukaryotic translation initiation
factor eIF4G to enhance efficiency of translation (8). NSP2 and NSP5
interact to form viroplasms, large inclusions in the cytoplasm where
core-like replication intermediates (core RIs)1 are assembled and RNA
replication takes place (2, 9, 10). As was shown in a study of a
temperature-sensitive mutant, NSP2 is required for the formation of the
viroplasm and is also essential in the synthesis of double-stranded RNA
in vivo (11). However, little is known about the mechanism
underlying this observation and the function of NSP2 on a molecular level.
NSP2 is a 35-kDa protein that forms homomultimers and interacts with
the RNA polymerase VP1 (12, 17). Furthermore, NSP2 interacts with and
induces hyperphosphorylation of NSP5 (13), which, in turn, interacts
with NSP6 (14, 15), possibly forming a regulatory multiprotein complex.
Both NSP2 and NSP5 possess a sequence-independent affinity for the
binding of RNA (17) and are associated with the structural proteins
VP1, VP2, and VP3 that form the core RIs that catalyze the synthesis of
double-stranded RNA (16). It is thought that NSP2 and NSP5 have an
active role in the packaging of RNA into cores (9). Recently, we have
shown that NSP2 possesses an NTPase activity, which suggests an
energy-dependent function for NSP2, possibly that of a
molecular motor involved in the packaging of mRNA into core RIs
(17).
In the present study, we have applied biophysical methods for the
characterization of recombinant NSP2, with the goal to better understand the molecular structure of NSP2 in solution and the interaction with its ligands. In particular, we have studied the secondary structure, the oligomeric state, and the hydrodynamic shape
of the protein in the presence of magnesium, nucleotide diphosphate and
triphosphate, RNA, and DNA. As will be described below, we found that
NPS2 self-assembles into relatively stable octamers, which can undergo
conformational changes in the presence of magnesium and ADP. This may
be of significance for understanding the function of NSP2.
Protein and Oligonucleotides--
Recombinant NSP2 was prepared
as described in detail in Ref. 17. In brief, a cDNA containing the
NSP2 open reading frame of the simian rotavirus SA11 was cloned into
the expression vector pQE60 (Qiagen). His-tagged rNSP2 was expressed in
Escherichia coli M15[pREP4] cells (Qiagen) and purified
using a Ni2+-nitrilotriacetic acid-agarose column. The
protein was dialyzed overnight against 10 mM Tris-HCl, pH
7.2, 10 mM NaCl, 0.5 mM dithiothreitol (DTT),
0.5 mM EDTA (except where noted otherwise). The protein was
obtained with a purity of >98%. The extinction coefficient and the
partial specific volume of the protein were calculated from the amino
acid composition using the program Sednterp (18). For the experiments
with phosphorylated NSP2, the protocol described previously was applied
(17), and the protein was incubated for 1 h at 37 °C in 5 mM MgCl2 and 0.5 mM ATP. This was
followed by dialysis against 10 mM Tris-HCl, pH 7.2, 10 mM NaCl, 0.5 mM DTT, and 0.5 mM EDTA.
For the study of nucleic acid binding of NSP2, a single-stranded 23-mer
of RNA (RNA23) (5'-UGAAAACGGCGACUGAGGAUACC3'; Dharmacon Research) and
single-stranded 23-mer of DNA (DNA23) (5'-TGAAAACGGCGACTGAGGATACC-3'; Life Technologies, Inc.) were used. The extinction coefficients of the
oligonucleotides were determined spectrophotometrically.
Sedimentation Equilibrium--
Sedimentation equilibrium studies
were conducted in a Beckman Optima XL-A centrifuge, using an
An50 Ti eight-hole rotor and absorbance optics. Double-sector
charcoal-filled epon centerpieces were filled with 120-130 µl of
sample at concentrations between 4 and 17 µM.
Sedimentation equilibrium was attained at a rotor temperature of
4 °C and at rotor speeds of 8,000 rpm and 12,000 rpm. Absorbance
profiles were acquired at wavelengths of 230, 250, and 280 nm, chosen
according to the protein concentration. In the presence of
oligonucleotide, ADP, or ATP Sedimentation Velocity--
Sedimentation velocity experiments
were conducted in a Beckman Optima XL-I ultracentrifuge, using an An50
Ti eight-hole rotor and the interference optical detection system.
Double-sector charcoal-filled epon centerpieces were filled with 350 µl of sample at concentrations between 1 and 10 µM. The
rotor temperature was set between 20 and 24 °C, chosen to minimize
temperature changes during a 1-2-h equilibration period of the rotor
in the vacuum chamber prior to the start of the run. The rotor was then
accelerated to 45,000 or 50,000 rpm at maximal rate, during which
already visible sedimentation took place, as judged by the camera
picture of the fringe profiles. Fringe displacement profiles were
acquired in intervals of 30-40 s. Data were analyzed with the software
SEDFIT, and errors reported are from replicate experiments.
The analysis of the sedimentation velocity profiles was performed by
direct boundary modeling by solutions of the Lamm equation,
The Lamm equation solutions were calculated by using the finite element
and moving frame of reference method described in Refs. 26-28, at a
radial increment of ~0.001 cm. The finite element simulations were
modified to include the consideration of the acceleration phase of the
rotor. An average acceleration rate and duration was calculated based
on the difference of the
To facilitate the analysis of systematic deviations in the fit, a
two-dimensional bitmap representation of the residuals was calculated
as described elsewhere.2 In
this picture, pixel rows represent the sequence of scans, and pixel
columns represent different radial values. The brightness at each pixel
is calculated by linearly mapping the values between
The sedimentation coefficient distributions were calculated by the
c(s) method by direct modeling with distributions
of Lamm equation solutions,
Dynamic Light Scattering--
Dynamic light scattering
experiments were conducted with a Protein Solutions DynaPro 99 instrument with DynaPro-MSTC200 microsampler (Protein Solutions,
Charlottesville, VA). Protein samples were drawn from the solution
column after equilibrium sedimentation after the ultracentrifuge was
stopped, and the cells were taken from the rotor with minimal
convection. Although it is difficult to control the protein
concentration in this way, this procedure is adequate for species
analysis, and because of the persistent gradient close to equilibrium
(31), it very effectively eliminates signal contributions from
particles outside the size range of interest. A 20-µl sample was
inserted in the 90° light scattering cuvette with the temperature
control set to 20 °C. The autocorrelation coefficients were exported
for analysis with the software SEDFIT, adapted for dynamic light
scattering analysis by replacing the single species Lamm equation
solutions by the intensity correlation function,
Circular Dichroism--
CD spectra of NSP2 and its complexes
with ADP, RNA, and DNA were measured in a Jasco J-715
spectropolarimeter at 25 °C, using 1-mm path length quartz cuvettes.
Control spectra were also measured on ADP, RNA, and DNA alone at the
same concentrations, and the spectrum of the protein in the complexes
was calculated as the difference between each sample and its control.
Protein concentrations were 130-150 µg/ml. Four scans were made
between 190 and 320 nm, at a speed of 50 nm/min, and with a time
constant of 1 s. The measured ellipticities (mdeg) were
converted into the mean residue ellipticity NSP2 Forms Octamers in Solution--
To characterize the state of
association of NSP2, we have performed sedimentation equilibrium,
sedimentation velocity, and dynamic light scattering experiments. Fig.
1 shows results from the hydrodynamic
measurements. The sedimentation velocity profiles can be fit very well
with a single species Lamm equation model with an s value of
s20 = 11.96 ± 0.1 S. The analysis of the
boundary spreading of the data shown in Fig. 1A gives a
molar mass of 301 kDa, which is virtually identical to the molar mass
of an octameric NSP2 of 301.4 kDa calculated from the amino acid
composition. However, the error associated with this estimate from
replicate experiments was observed to be approximately ±10-20 kDa.
Therefore, direct measurements of the diffusion coefficients were
performed separately by dynamic light scattering (Fig. 1D).
The autocorrelation data were well described by a single component with
D20 = 3.63 × 10
An independent confirmation of this result was obtained from
sedimentation equilibrium experiments. As shown in Fig.
2, the sedimentation equilibrium
distributions are well described at several loading concentrations and
rotor speeds by a model for a single octameric species. When the molar
mass is treated as an unknown parameter in the global analysis, a value
corresponding to 7.8 ± 0.2 subunits is obtained. From the
sedimentation equilibrium data alone, it cannot be ruled out that there
might be significant populations of both heptamers and nonamers.
However, the fact that the boundary spreading in sedimentation velocity
experiments is consistent with that of a single octameric species
strongly indicates the absence of heterogeneity of NSP2 and suggests
that octamer formation is a highly specific process.
Sedimentation velocity experiments with phosphorylated NSP2 led to
results indistinguishable from those of nonphosphorylated protein (data
not shown). We also performed sedimentation velocity and equilibrium
experiments with (nonphosphorylated) NSP2 in solutions of pH 5.2 and pH
9.2 and in varying concentrations of NaCl (0-100 mM).
These experiments also led to virtually identical results (data not
shown), indicating a high stability of the oligomer. Furthermore,
consistent with the apparent absence of a slower sedimentation boundary
in Fig. 1A, the fit to the sedimentation velocity data
cannot be improved by allowing for a smaller species in the model,
which indicates that the free NSP2 monomer concentration is below the
sensitivity of the interference optical systems, which corresponds to
concentrations of <0.1 µM.
After identifying the oligomeric state of NSP2 as an octamer, the
sedimentation coefficient of s20 of 11.96 S can
be used to calculate a frictional ratio of 1.30 and a Stokes radius of 5.8 nm. In terms of gross hydrodynamic shape, with an estimated hydration ratio of 0.383 g/g calculated from the
amino acid composition (18, 34) these values would correspond to an
equivalent prolate ellipsoid with major and minor axis of 22.5 × 6.9 nm or to an equivalent oblate ellipsoid with dimensions 15.4 × 4.5 nm. Qualitatively similar values for the dimensions of the
equivalent ellipsoids are obtained with a lower estimate for the
hydration (Table I).
NSP2 Octamers Dissociate in the Presence of Magnesium--
Since
it was shown recently that NSP2 has an NTPase activity, for which the
presence of magnesium is an essential cofactor, with optimal
concentrations between 1 and 5 mM MgCl2 (17),
we next studied the protein in 5 mM MgCl2. It
can be seen from Fig. 3 that under these
conditions NSP2 exists in a heterogeneous mixture of species, including
populations that sediment significantly slower than the octamer. This
effect disappeared when overnight dialysis in buffer without
MgCl2 followed the incubation with magnesium, which
indicates the reversibility of the observed magnesium effect. As is
described below, this change in the oligomeric state by magnesium is
also accompanied by small changes in secondary structure.
To unravel the distribution of species, the sedimentation coefficient
distribution was determined. The commonly used approach for a
model-free analysis is an apparent sedimentation coefficient distribution g*(s), which operates in the
approximation of D = 0 (no diffusion). A much more
realistic treatment for diffusion has recently been developed, where an
approximate average frictional ratio is used to estimate the diffusion
coefficients D(s) as a function of the
sedimentation coefficient (30), and initial applications have
demonstrated the high resolution achieved (35).2 The
resulting distributions c(s) are not
diffusionally broadened, and the residual width of the peaks obtained
for discrete protein species is dependent on the correctness of the
shape assumption and (due to the maximum entropy regularization
employed) on the signal-to-noise ratio of the data. We have used this
method in the present study with the estimate 1.3 for
f/f0 of all species, using the
property of c(s) that it is not very sensitive to
this value and assuming that the frictional ratio of the octameric NSP2
may be a reasonable approximation for the other unknown species. The
resulting c(s) distribution obtained from the
data presented in Fig. 3A shows two main peaks at 6.4 S
(24%) and 11.9 S (57%) as well as very small populations at 3.5 S
(2%), between 8 and 10 S (7.6%), and at 14.5 S (3.1%).
A discrete Lamm equation analysis was also performed to analyze the
boundary spreading from the ~12 S species. This resulted in a molar
mass estimate of 306 ± 10 kDa, suggesting that this species is
still octameric. To further improve the size distribution analysis by
utilizing this knowledge of the 12 S species being octameric with a
theoretical molar mass of 301 kDa, we have modified the
c(s) method such that the diffusion coefficients
of the species between 10 and 13 S are calculated by the Svedberg
equation. This combines the "model-free" analysis of the
sedimentation coefficient distribution with the assignment of the ~12
S peak as the octamer, and instead of assuming shape similarity, it
constitutes a model more appropriate for protein complexes with
possible conformational changes (termed
CM(s)). The results are shown in Fig.
3D, clearly reflecting the multimodal boundary shape, with
populations of protein at ~3.5, 6.44, ~9, 11.90, and ~15 S.
In both the continuous c(s) and the discrete Lamm
equation analysis, it is apparent that the sedimentation coefficient of the octamer appears at a slightly lower value of 11.80 S. Although this
difference in the octameric sedimentation coefficient induced by
magnesium is very small, it was reproducible and significant when
results from the same protein preparation and from the same velocity
run were compared. One possible explanation of this smaller s value could be a shift in the apparent peak s
value due to the chemical reaction of the protein coupled to the
sedimentation. However, dissociation of the octamer was slow compared
with the sedimentation. When interpreted in the context of hydrodynamic shape, this small increase in the hydrodynamic friction would correspond to a slightly more asymmetric shape of the octamer (Table
I).
The identification of the nature of the second most abundant species at
6.4 S is more difficult, although this s value can theoretically only be assumed by oligomers larger than the dimer. Sedimentation equilibrium profiles and dynamic light scattering data
were acquired (data not shown), but they lead to ill conditioned analyses. From dynamic light scattering, the distribution of Stokes radii showed a single peak at 5.9 nm. In sedimentation equilibrium, the
weight-average molar mass corresponded to 6.9 subunits, and a
two-component model of the sedimentation equilibrium with an octameric
species and an unknown oligomer resulted in the average subunit number
of 5.5 for the smaller species, although good fits with a
tetramer/octamer model could be obtained. This suggests that the second
major peak in the sedimentation coefficient distribution may be a
tetramer, with the species between 8 and 10 S being intermediates. Results from the interpretation of the gross hydrodynamic shapes are
summarized in Table I.
ADP and ATP
This indicates that the oligomeric state of NSP2 is not affected by the
binding of ADP. Unfortunately, we were unable to unambiguously clarify
the origin of the increased sedimentation coefficient. However, the
sedimentation coefficient of NSP2 is increased by 5%, which is a
significantly larger increase than one would expect on the basis of the
additional mass of eight nucleotides (assuming one binding site per
protomer; to account for the increased sedimentation rate through the
additional mass of bound nucleotides alone, this would require >30
nucleotides if the frictional coefficient of the complex would not
change, or >45 nucleotides with the usual
To study in more detail the nature of this faster sedimenting
protein-nucleotide complex, we performed sedimentation velocity experiments in the presence of 50 mM NaCl (so as to
diminish the potential electrostatic interactions of the nucleic acid
binding sites of the basic protein with the nucleotides) and with
different nucleotide concentrations. First, at an ADP concentration of
50 µM, we applied multidetection to observe
simultaneously the sedimentable absorbance at 260 and 280 nm, as well
as the fringe shift data for the total protein distribution (data not
shown). From the ratio of 260- to 280-nm absorbance, we found that at
this concentration ~0.3 nucleotides are bound per protein protomer.
Surprisingly, no shift in the sedimentation coefficient was observed in
comparison with a control sample without nucleotides. When the ADP
concentration was increased, no shift in the sedimentation coefficient
was observed below 0.4 mM, while above this concentration
the sedimentation coefficients were consistently higher by 0.6-0.7 S. No significant difference in the s value between 1 and 2 mM ADP was observed. (Unfortunately, due to the high
extinction of the nucleotides, absorbance detection for the measurement
of protein/nucleotide ratio is not possible at nucleotide concentration
higher than ~50 µM.) Although the limited precision of
the small changes in the s values does not permit the
interpretation of an isotherm s(c), this steep
concentration dependence in the range of 0.5 mM would be
consistent with a cooperative transition of the octamer conformation.
In experiments with the nonhydrolyzable analogue ATP
An additional aspect of nucleotide binding can be observed in the
absence of a slower sedimenting component in Fig. 4, which shows that
nucleotide binding prohibits the MgCl2-induced dissociation of NSP2. To see if binding of ADP can lead to reassembly of NSP2 that
was already dissociated by 5 mM MgCl2, we
performed a sequential sedimentation velocity experiment. After
measuring the sedimentation profiles shown in Fig. 3A, we
mixed the solution in the ultracentrifuge cell to resuspend the
sedimented protein. (In control experiments, we observed that mixing
the solution column and restarting a second velocity experiment did not
change the sedimentation parameters, and full resuspension of the
material was achieved.) We then added 1 mM ADP to the
solution and incubated for 3 h, before starting a new velocity
run. The resulting sedimentation coefficient distribution is shown by
the dotted line in Fig. 3D. After the
addition of ADP, the population was shifted from the dissociated state
back to an octamer in the more compact form, indicating that the
MgCl2-induced dissociation is reversed by binding of ADP.
However, no complete reversibility was observed, which may indicate a
slow kinetic process of reassembly.
NSP2 Octamer Binds RNA and DNA--
It has been shown previously
that NSP2 binds both RNA and DNA with high affinity, which may be
central to its function
(17).3 Therefore, we were
interested in the effects of nucleic acid binding to the protein. For
the present study, we have chosen short oligonucleotides (23-mers) to
simplify the protein-RNA/DNA interaction and to avoid the possible
cooperative binding of more than one functional unit of NSP2 to a
nucleic acid strand, forming larger complexes such as those observed
previously by electrophoretic mobility shift assays (17).
First, we wanted to demonstrate that RNA23 binds NSP2 under our
experimental conditions. Therefore, we performed sedimentation equilibrium experiments, with absorbance scans acquired at the absorbance maxima of both the protein and the nucleic acid (Fig. 5). The equilibrium distribution could be
well modeled with a single species of 301 kDa. (It can be expected that
the buoyant molar mass of the oligonucleotides is negligible compared
with the octameric protein.) Since the sedimenting absorbance at 260 nm
is significantly larger than would be expected based on the protein
extinction at this wavelength (dashed line in
Fig. 5A), it is apparent that the RNA23 is bound to the NSP2
octamer. The quantitative analysis of the increased absorbance signal
at 260 nm leads to a molar ratio of 2.0 ± 0.4 RNA23 molecules per
NSP2 octamer. Similar results were obtained for the binding of
DNA23.
As an independent measurement of the oligomeric state of NSP2 when
bound to nucleic acid, we performed sedimentation velocity experiments
(Fig. 6). The sedimentation coefficient
distribution of the NSP2-nucleic acid complex shows a peak at
s20 = 12.11 ± 0.03 S (Fig. 6D)
(12.06 ± 0.06 S for DNA23), which is consistent with the value of
s20 = 11.96 S in the absence of
oligonucleotides, considering the small increase in mass. To study the
effect of other ligands, we performed sedimentation velocity
experiments in the presence of ADP or MgCl2. When 1 mM ADP was present, the protein-RNA complex
sedimented at a slightly higher rate of 12.38 (0.04) S (12.35 S for the
protein-DNA23 complex). In the presence of MgCl2, a value
of 12.04 S was measured, accompanied by some dissociation of the
octamer (Fig. 6D). These shifts in the sedimentation coefficient distributions are consistent with the effects of
nucleotides and magnesium on the protein alone (Fig. 3D).
However, the magnesium-induced dissociation seemed to be less in the
protein/RNA mixture. Additionally, a sequential sedimentation velocity
experiment performed with 9.7 µM NSP2 and 5 mM MgCl2, followed by resuspension of the
solution, incubation with 1.5 µM RNA23, and a repeat
sedimentation velocity experiment indicated a 15% increase in the
population of octamer after the addition of RNA (data not shown). In a
separate sedimentation velocity experiment with DNA23 in 5 mM MgCl2 after longer incubation, no
dissociation was observed, and as described in the following, the
secondary structure changes induced by magnesium are absent in the
presence of RNA or DNA.
Circular Dichroism Studies of the Secondary Structure of
NSP2--
Fig. 7A shows the
CD spectrum of NSP2, typical of an In the present paper, we have applied hydrodynamic and
thermodynamic techniques and CD spectroscopy to the study of structural aspects of NSP2 of rotavirus in solution, with the goal to better understand the function of NSP2 in the life cycle of the virus. It was
shown previously that NSP2 assembles into multimers (12, 17), which
were also detected in infected cells (12). Our results indicate that
these multimers are highly stable octamers, formed by very specific and
strong self-assembly, and that they are the oligomeric state relevant
for RNA binding of the protein as well as NTPase activity. Furthermore,
our data show that NSP2 can undergo conformational changes that are
controlled by its ligands. Several structural and functional
consequences will be discussed in the following.
From circular dichroism, we found that the secondary structure of the
protein contains a high amount of It has been demonstrated that the helicase Dmc1, a eukaryotic homologue
of the E. coli RecA, forms an octameric ring with a central
channel for binding DNA (41). The Dmc1 octamer appeared in electron
micrographs as a symmetrical side-by-side arrangement of eight
monomers, similar to the hexameric ring-structure of other helicases
(42). Despite the same number of subunits of Dmc1 and NSP2 involved in
polynucleotide binding, the ability of the NSP2 to dissociate in
relative distinct tetrameric subunits does not seem to support a strong
structural relationship between these proteins. (Interestingly, initial
sedimentation velocity experiments with a protein related to NSP2, the
NS2 protein from bluetongue virus, could be modeled on the basis of a
rapidly reversible monomer-tetramer-octamer self-association
equilibrium.)4 SSB proteins
from E. coli and several other species have been found to be
tetrameric (43-46), with DNA wrapped around the protein complex (44,
47). EcoSSB can cooperatively multimerize to higher oligomers when
bound to the nucleic acid, and it has been proposed that this includes
the formation of octamers through the dimerization of tetramers (48);
similar octameric forms were also suggested for the vaccinia virus I3L
gene product (49). However, generally little sequence similarity of
EcoSSB and related proteins with the single-stranded DNA-binding
proteins of animal viruses are found (50), and it is not clear if NSP2
and EcoSSB share more structural motifs than a high fraction of
From the data described in the present paper, we found that the octamer
has a Stokes radius of 5.8 nm and frictional ratio of 1.3, which
indicates that it is spatially not very compact. Binding of magnesium
is accompanied by small conformational changes in the secondary
structure and by significantly weaker self-association of the protein,
which allows additional insight into the structure of the protein. Our
data suggest that in the presence of magnesium the octamer may adopt a
slightly more "loose" conformation and partially dissociates, with
the main smaller species probably being a tetramer. When using the
gross hydrodynamic models of equivalent ellipsoids, in the prolate
model the tetramer would appear more extended than the octamer, while
in the oblate model it would appear to be approximately half the
thickness of the octamer. Because a possible mechanism of octamer
formation would consist of the dimerization of two tetramers, it is
noteworthy that the oblate model would suggest an arrangement with two
tetramers stacked on top of each other. Unfortunately, because of the
poor ability of hydrodynamic modeling to predict precise molecular shapes, the clarification of the spatial structure has to await the
results from methods with higher resolution.
A second type of conformational change of NSP2 was observed when
nucleotide diphosphates or triphosphates were bound. Under these
conditions, a small but significant increase of the rate of
sedimentation was observed, corresponding to a more compact octameric
conformation with a frictional ratio of 1.23. One alternative explanation for this may be the binding of a large number of
nucleotides to the protein, possibly through electrostatic
interactions. However, our studies with buffers of different ionic
strength and at different nucleotide concentrations make this appear
unlikely. Furthermore, the steep concentration dependence of the shift
in sedimentation rate rather suggests the hypothesis of an allosteric,
cooperative transition in the octamer conformation. Interestingly,
nucleotides also seem to reverse the effect of magnesium to weaken the
intersubunit interactions of the octamer. As judged by the oblate
ellipsoid model, the total spatial amplitudes of the conformational
changes corresponding to the increased sedimentation rate are 1.3 and 2 nm. Despite the lack of precise three-dimensional structures at the
current stage, these changes are highly significant. Since we did not
observe large effects of ADP or ATP The role of these conformational changes of NSP2 is not clear.
Although, in principle, it could provide for an allosteric regulatory
mechanism, the conformational changes may also have a more specific
role as mechanical transducers. Because NSP2 associates with core RIs
and cross-linking of infected cell-lysates showed it to interact with
the RNA polymerase VP1 (12), it has been implicated in the packaging of
mRNA into cores (9, 36). Based on the finding of NTPase activity of
NSP2, it has recently been suggested that the protein may function as a
molecular motor (17), possibly utilizing the energy obtained from
hydrolysis for the transport of mRNA through the channels of core
RIs. Since a molecular motor requires at least two distinct
conformational states with mechanically different shapes, the changes
of hydrodynamic shape from a loose to a more compact or tight
conformation upon nucleotide binding is consistent with such a
hypothesis. However, we have insufficient knowledge of how the reverse
change from the compact to the more loose conformation would take
place, and how this could be exploited for packaging. So far, only few
findings seem to provide partial insights. Both ADP and ATP On the other hand, this mechanical NSP2 function could also require the
interaction with other viral proteins, such as NSP5. NSP5 and NSP2
coimmunoprecipitate in the presence of RNA after UV cross-linking (29),
and both proteins can be cross-linked with the RNA polymerase VP1 (13),
suggesting that NSP5 also is a component of the multiprotein complex
that possesses replicase activity (13, 29, 36). Also, NSP2 up-regulates
the phosphorylation of NSP5 (13), possibly by transfer of the phosphate
group obtained from the NTPase activity of NSP2 (17). The
conformational changes of NSP2 observed here after binding of
nucleotides seem to add an additional perspective of the NSP2-NSP5
interaction with regard to a possible role of NSP5 in assisting the
conformational transitions in a multiprotein complex.
In the present study, we have focused on the oligomeric state of the
NSP2 protein. Combining the findings from ultracentrifugation and CD
spectroscopy that the octameric NSP2 binds RNA and that RNA binding
stabilizes the octamer suggests that this is the functional unit for
RNA binding. In a recent study, it was observed that NSP2 exhibits a
nucleotide- and magnesium-independent strand displacement activity,3 which could play a role in the unwinding of the
mRNA template used for double-stranded RNA synthesis.
Interestingly, this activity was inhibited in a
concentration-dependent way by magnesium, which parallels
our observation that magnesium causes partial dissociation of the NSP2
octamer into tetramers. Further studies will be required that
investigate the stoichiometry of the protein-mRNA complexes, the
possibility of cooperative binding of multiple NSP2 octamers to longer
RNA strands (17), and the affinity of mRNA for NSP2 in the
different conformations.
-sheet,
with small changes induced by magnesium that were reversed in the
presence of RNA. That NSP2 can exist in different conformations lends
support to the previously proposed hypothesis (Taraporewala, Z., Chen,
D., and Patton, J. T. (1999) J. Virol. 73, 9934-9943) of its function as a molecular motor involved in the
packaging of viral mRNA.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
S, additional scans at 260 and 285 nm
were taken. Data analysis was performed by global analysis of several
data sets obtained at different loading concentrations and rotor speeds
using the mathematical modeling software MLAB (Civilized
Software, Silver Spring, MD). No thermodynamic nonidealities were
observed, and least-squares fits were based on Boltzmann distributions
of ideal species in the centrifugal field,
where a
(Eq. 1)
denotes the experimentally
measured absorbance at a wavelength
and at the distance
r from the center of rotation, c and
the concentration and extinction coefficient of the protein, d the optical path length of 1.2 cm, M
and
the solvent density,
the angular velocity of
the rotor, T the absolute temperature, R the gas
constant, r0 an arbitrary reference radius, and
a base-line absorbance (19), and their well known combination for
models involving multiple species (20-22). Reported errors in the
equilibrium analyses are calculated confidence intervals from
F statistics (23).
(Eq. 2)
where a(r, t) denotes the experimentally measured
absorbance,
and
the systematic and random signal offset,
s and D the sedimentation and diffusion
coefficients of the protein, and
its concentration at position
r and time t (24). This was combined with the
algebraic calculation of systematic time-invariant and radial-invariant
noise components
(25). Because these systematic signals are
arbitrary offsets introduced from the detection system, calculated
systematic offsets can be subtracted from the raw data without
introduction of bias if the degrees of freedom in the analysis
are not reduced. Therefore, the final calculated best-fit offsets are subtracted from the raw data for presentation.
2t and
the t entry of the data files, and the rotor
speed-dependent matrix elements in the finite element
algorithm were dynamically updated in intervals of 10 s for the
calculated duration of the acceleration phase. In contrast to the
previous strategy of calculating effective sedimentation times from the
elapsed
2t, which provides the
correct boundary positions, the simulated acceleration phase
additionally calculates the extent of diffusion during the acceleration
phase and thereby improves the accuracy of the boundary shapes and
leads to slightly improved quality of the fits. Using a model for
multiple discrete species, the location of the meniscus was treated as
a floating parameter and optimized in the nonlinear regression of the
data. The resulting meniscus position was then used in the continuous
size distribution analysis (see below).
0.05 and +0.05
to grayscale values from 0 to 256. This transformation should result in
a neutral gray picture without structure for a perfect fit, and any
visible structure represents systematic deviations of the model from
the data.
where c0(s)ds is
the loading concentration of species with a sedimentation coefficient
between s and s + ds). Following the procedure outlined in detail in Ref. 30, the integral was solved by
discretization into 250-300 s values between 4 and
18 S (unless noted otherwise), and a value of 1.3 for the frictional
ratio f/f0 was used for estimating
the average diffusional broadening of the sedimentation boundaries
D(s). Maximum entropy regularization was used to
calculate the simplest size distribution within a confidence level of
0.68 of the best-fit distribution (30). While the use of the same
frictional ratio for all species estimates the average degree of
diffusion (corrected for the different macromolecular sizes via
Stokes-Einstein relationship), a more refined calculation was performed
once the ~12 S peak was identified as an octamer. The known molar
mass of the oligomer was applied to all species in the sedimentation
coefficient distribution within the range from 10 to 13.5 S, and
diffusion coefficients were calculated directly by insertion of
s and M into the Svedberg equation (19),
(Eq. 3)
For all species outside this predefined range of s
values, the approximation by using the frictional ratio was maintained. This method allows analyzing more rigorously the distribution of
sedimentation coefficients for a macromolecule with known molar mass
that can be present in different conformations, and its application leads to slightly sharper sedimentation coefficient distributions. To
distinguish this distribution and indicate the use of a molar mass
value, it will be referred to as cM(s) in
the following.
(Eq. 4)
where
(Eq. 5)
represents the decay time and q = (4
n/
)sin(
/2), with the solvent refractive index
n, the wavelength of the incident light
, and the
scattering angle
(32). The size distribution analysis was performed
analogous to the sedimentation coefficient distribution analysis, by
replacing the Lamm equation solutions in the kernel by autocorrelation
functions of the form in Equation 5.
as follows,
where MRW is the mean residue weight (115.9),
l is the path length in cm, and c is the protein
concentration in mg/ml. Secondary structure was estimated from averaged
spectra using the CONTIN program (33), and reported errors are from the
variation of results for fits of a quality close to that of the best fit.
(Eq. 6)
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
7
cm2/s (with very minor impurities visible outside the size
range of the protein). This value for D is consistent with
the boundary spreading of the sedimentation velocity data, as indicated
by the excellent fit to the Lamm equation model using the predetermined D (Fig. 1, B and C) with a root mean
square (r.m.s.) deviation in the order of the instrument noise, and
only minor systematic deviation visible in the residuals bitmap. Taking
the values of s and D together, a molar mass
estimate of 312 kDa is obtained. This strongly suggests that NSP2 is
octameric in solution.
View larger version (54K):
[in a new window]
Fig. 1.
Hydrodynamic analysis indicates that NSP2
forms an octameric species. A, sedimentation velocity
data of 9.7 µM NSP2 in 10 mM Tris-HCl, pH.
7.2, 10 mM NaCl, 0.5 mM DTT, 0.5 mM
EDTA, sedimenting at 50,000 rpm, 22 °C. B and
C, bitmap and overlay representation of the residuals from a
single-species model with D20 = 3.81 × 10 7 cm2/s (predetermined from
dynamic light scattering) and with s20 = 11.97 S, which leads to a r.m.s. deviation of 0.0059 fringes.
D, autocorrelation coefficients from a dynamic light
scattering experiment (circles) and best-fit autocorrelation
function (solid line) based on a model for the
species D20 = 3.81 × 10
7 cm2/s, in the presence of
signal contributions of two species with D20 = 27.1 × 10
7 cm2/s (1.2%
total signal contribution) (dotted line) and
D20 = 0.057 × 10
7 cm2/s (2.1%)
(dashed line). These contaminating species are
not visible in the sedimentation velocity profiles. The
inset shows the distribution of hydrodynamic radii (in nm),
with a peak at 5.8 nm. E, residuals of the fit, which has a
r.m.s. deviation of 1.8 × 10
4. Assuming
a partial specific volume of 0.74 ml/g, a molar mass of 312 kDa is
obtained when combining the values of s and
D.
View larger version (40K):
[in a new window]
Fig. 2.
NSP2 in sedimentation equilibrium can also be
modeled as an octameric species. NSP2 in 10 mM
Tris-HCl, pH 7.2, 100 mM NaCl, 0.5 mM DTT, 0.5 mM EDTA in sedimentation equilibrium at 4 °C.
A, experimental absorbance profiles at 280 nm at a loading
concentration of 17 µM (circles), 8 µM (triangles), and 4 µM
(squares) at 8,000 rpm (all offset by 0.2 OD and 0.1 cm),
and 17 µM (+), 8 µM (×), and 4 µM (inverted triangles) at 12,000 rpm. Solid lines are the best-fit single species
distribution with a molar mass of 301.4 kDa. B, residuals of
the fit, which lead to an r.m.s. deviation of 0.011 OD.
Estimated dimensions of equivalent hydrodynamic ellipsoids for NSP2
View larger version (46K):
[in a new window]
Fig. 3.
NSP2 exhibits a mixture of fast and slow
sedimenting species in the presence of Mg. A,
sedimentation velocity data of 6.6 µM NSP2 in 10 mM Tris-HCl, pH 7.2, 10 mM NaCl, 5 mM MgCl2, 0.5 mM DTT, 0.5 mM EDTA, sedimenting at 50,000 rpm, 20 °C. For clarity,
every fifth scan is shown. B and C, bitmap (all
scans) and overlay representation (every fifth scan) of the residuals
from the continuous cM(s) model when
constraining the molar mass of all species between 10 and 13 S to 301 kDa, as a model for the sedimentation coefficient distribution of
octameric NSP2 with different conformational states. The boundary
spreading of all other species (slower than 10 S and faster than 13 S)
was estimated via a frictional ratio of 1.3. The r.m.s. deviation is
0.0062 fringes. The resulting cM(s)
distribution is shown in D (solid
line). Also shown in D is the corresponding
cM(s). distribution of the same NSP2
sample in 5 mM Mg after the addition of 1 mM
ADP (dotted line), and for reference the
cM(s) distribution in the absence of Mg
and ADP (dashed line).
S Bind to Octameric NSP2 and Increase the
Sedimentation Coefficient--
As the next step, we examined effects
of nucleotide diphosphate and a triphosphate analogue on the
conformation of NSP2. Fig. 4 shows the
sedimentation coefficient distributions
cM(s) of NSP2 in the presence of 1 mM ADP obtained from velocity centrifugation. It is clearly
visible that NSP2 is sedimenting faster, with a sedimentation
coefficient of s20 = 12.61 S. To rule out the
possibility that larger oligomers may have formed, we calculated the
apparent molar mass from boundary spreading to be 296-306 kDa. The
analysis of the autocorrelation coefficients from dynamic light
scattering in the presence of ADP leads to a D20
value of 3.71× 10
7 cm2/s (data
not shown), which, taken together with the sedimentation coefficient of
12.61 S, corresponds to a molar mass of 318 kDa. Therefore, within
experimental uncertainty, this is consistent with the molar mass of the
NSP2 octamer.
View larger version (15K):
[in a new window]
Fig. 4.
ADP and ATP S induce
conformational change in NSP2. Sedimentation coefficient
distributions cM(s) from the
conformational change model for species between 10 and 13.5 S. NSP2 in
10 mM Tris-HCl, pH 7.2, 10 mM NaCl, 0.5 mM DTT, 0.5 mM EDTA is shown for reference
(solid line) and after the addition of 1 mM ADP (dashed line), 1 mM ADP plus 5 mM MgCl2
(dashed and dotted line),
or 1 mM ATP
S plus 5 mM MgCl2
(dotted line).
S is added in the
presence of 5 mM MgCl2. Furthermore, a similar
increase of the sedimentation coefficient was observed for
phosphorylated NSP2 (data not shown).
S, we found
already a significant
s at concentrations of 0.1 mM, suggesting a higher affinity of this nucleotide.
View larger version (27K):
[in a new window]
Fig. 5.
RNA binds to octameric NSP2.
A, absorbance scans of 9.7 µM NSP2 in
sedimentation equilibrium with 1.5 µM RNA23 are shown at
wavelengths of 280 nm (circles) and 260 nm
(squares). As calculated from the absorbance ratio, two
RNA23 molecules bind per NSP2 octamer. Solid
lines are calculated best-fit distribution for octameric
NSP2. For comparison, the dashed line shows the
signal at 260 nm that would be expected for the protein alone.
Experimental conditions were as follows: 10 mM Tris-HCl, pH
7.2, 100 mM NaCl, 0.5 mM DTT, 0.5 mM EDTA in sedimentation equilibrium at 8,000 rpm and at
4 °C. B, residuals of the fit.
View larger version (50K):
[in a new window]
Fig. 6.
NSP2 is octameric in the presence of
RNA. A, sedimentation velocity data of 9.7 µM NSP2 and 1.5 µM RNA23 in 10 mM Tris-HCl, pH 7.2, 10 mM NaCl, 0.5 mM DTT, 0.5 mM EDTA, sedimenting at 50,000 rpm,
22 °C. B and C, bitmap and overlay
representation of the residuals from the continuous
c(s) model. The r.m.s. deviation is 0.0044 fringes. The resulting c(s) distribution is shown
in D (solid line). For comparison,
D also shows the c(s) distribution
obtained in the presence of 1 mM ADP (dashed
line) and 5 mM MgCl2
(dotted line).
/
protein. Analysis with the
CONTIN program gave a best fit with estimates of 24%
-helix and
45%
-sheet. The addition of 5 mM MgCl2
seemed to be accompanied by a small change in secondary structure (best fit 32%
-helix, 35%
-sheet), apparent from Fig. 7A
in the decreased ellipticity at 210 nm. This indicates that the
observed dissociation is accompanied by or the result of changes in the
secondary protein structure. After incubation of the protein-magnesium
complex for 24 h, magnesium-induced change seemed to be reversed
with 1.5 µM RNA23 or 1.5 µM DNA23 (Fig.
7B). This supports the reversibility of the magnesium effect
on the protein by binding of oligonucleotides as observed by
sedimentation velocity (above). Because their spectra were of such low
intensities, it was impossible to detect any conformational changes
induced in the nucleic acids after binding to the protein.
Interestingly, the addition of 1 mM ADP did not appear to
restore protein secondary structure to its state in the absence of
magnesium and nucleotide (Fig. 7A, best fit 23%
-helix,
55%
-sheet), suggesting that the conformational change that leads
to a more compact octamer is also accompanied by small changes in the
secondary structure.
View larger version (22K):
[in a new window]
Fig. 7.
CD spectra of NSP2. A, CD
spectrum of NSP2 is shown (circles) with best fit
corresponding to 24% -helix (24-28%) and 45%
-sheet
(29-50%) (solid line). Also shown are CD data
in the presence of 5 mM MgCl2
(squares) with a best fit of 32%
-helix (30-36%), 35%
-sheet (27-36%) (solid line), and in the
presence of 5 mM MgCl2 and 1 mM ADP
(+) with a best fit of 23%
-helix (14-32%), 55%
-sheet
(26-58%) (solid line). B shows the
spectra of NSP2 in the presence of 1.5 µM RNA23 (×)
(best fit leads to 24%
-helix (23-29%), 35%
-sheet
(28-41%)) or 1.5 µM DNA23 (
) (24%
-helix
(24-28%), 22%
-sheet (22-41%)). As a reference, the CD spectrum
of NSP2 is also shown again (solid
circles).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
-sheet. This correlates well with
the secondary structure found for single-stranded DNA-binding proteins
of several other organisms (37-40). With respect to the tertiary and
quaternary structure, as has been pointed out previously, single-stranded DNA-binding proteins exhibit a great variety of oligomeric states in different species (37), and they also differ in
the mode of DNA binding. While some partial similarities of NSP2 with
other single-stranded DNA-binding proteins can be observed, it may be
that NSP2 adopts a novel quaternary structure.
-sheet. Supporting this view are preliminary pictures from cryo-EM
reconstructions,5 which show
a barrel-shaped particle, for which one could speculate that the
central cavity in the NSP2 may be the binding site for mRNA (36),
either accommodating a loop of mRNA or with NSP2 threaded on the
nucleic acid strand.
S on the secondary structure,
this shape change may involve rearrangement of protein domains.
S bind
NSP2 in the tighter conformation (Fig. 4), although ATP
S may have a
higher affinity. We did not find a difference in the hydrodynamic
behavior as a result of phosphorylation, and it has been shown that
phosphorylated NSP2 retains mRNA binding activity (17). In the
absence of nucleotides, binding of mRNA occurs to the loose
conformation. Clearly, a more detailed study of mRNA interaction
with the protein under the different conditions is required (see below).
![]() |
FOOTNOTES |
---|
* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed: Laboratory of
Infectious Diseases, National Institutes of Health, 7 Center Dr., MSC
0720, Rm. 117, Bethesda, MD 20892. Tel.: 301-496-3372; Fax: 301-496-8312; E-mail: jpatton@niaid.nih.gov.
Published, JBC Papers in Press, December 19, 2000, DOI 10.1074/jbc.M009398200
2 P. Schuck, D. Burkwall, W. Newcomb, D. Schubert, and J. Brown, submitted for publication.
3 Z. Taraporewala and J. T. Patton, (2001) J. Virol., in press.
4 P. Schuck, Z. Taraporewala, P. McPhie, and J. T. Patton, unpublished data.
5 H. Jayaram, Z. Taraporewala, J. T. Patton, and B. V. V. Prasad, unpublished data.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
RI, replication
intermediate;
DTT, dithiothreitol;
ATPS, adenosine
5'-O-(thiotriphosphate);
r.m.s., root mean square.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Chen, D., Luongo, C. L., Nibert, M. L., and Patton, J. T. (1999) Virology 265, 120-130[CrossRef][Medline] [Order article via Infotrieve] |
2. | Estes, M. K. (1996) in Fundamental Virology (Fields, B. N. , Knipe, D. M. , and Howley, P. M., eds) , pp. 731-761, Lippincott Williams & Wilkins, Philadelphia |
3. | Prasad, B. V., and Chiu, W. (1994) Curr. Top. Microbiol. Immunol. 185, 9-29[Medline] [Order article via Infotrieve] |
4. | Prasad, B. V., Burns, J. W., Marietta, E., Estes, M. K., and Chiu, W. (1990) Nature 343, 476-479[CrossRef][Medline] [Order article via Infotrieve] |
5. | Chen, D., Zeng, C. Q., Wentz, M. J., Gorziglia, M., Estes, M. K., and Ramig, R. F. (1994) J. Virol. 68, 7030-7039[Abstract] |
6. | Tian, P., Ball, J. M., Zeng, C. Q., and Estes, M. K. (1996) J. Virol. 70, 6973-6981[Abstract] |
7. | Tian, P., Ball, J. M., Zeng, C. Q., and Estes, M. K. (1996) Arch. Virol. Suppl. 12, 69-77[Medline] [Order article via Infotrieve] |
8. |
Vende, P.,
Piron, M.,
Castagne, N.,
and Poncet, D.
(2000)
J. Virol.
74,
7064-7071 |
9. | Patton, J. T., and Spencer, E. (2000) Virology 277, 217-225[CrossRef][Medline] [Order article via Infotrieve] |
10. | Fabbretti, E., Afrikanova, I., Vascotto, F., and Burrone, O. R. (1999) J. Gen. Virol. 80, 333-339[Abstract] |
11. | Chen, D., Gombold, J. L., and Ramig, R. F. (1990) Virology 178, 143-151[Medline] [Order article via Infotrieve] |
12. | Kattoura, M. D., Chen, X., and Patton, J. T. (1994) Virology 202, 803-813[CrossRef][Medline] [Order article via Infotrieve] |
13. | Afrikanova, I., Fabbretti, E., Miozzo, M. C., and Burrone, O. R. (1998) J. Gen. Virol. 79, 2679-2686[Abstract] |
14. |
Torres-Vega, M. A.,
Gonzalez, R. A.,
Duarte, M.,
Poncet, D.,
Lopez, S.,
and Arias, C. F.
(2000)
J. Gen. Virol.
81,
821-830 |
15. | Gonzalez, R. A., Torres-Vega, M. A., Lopez, S., and Arias, C. F. (1998) Arch. Virol. 143, 981-996[CrossRef][Medline] [Order article via Infotrieve] |
16. | Gallegos, C. O., and Patton, J. T. (1989) Virology 172, 616-627[Medline] [Order article via Infotrieve] |
17. |
Taraporewala, Z.,
Chen, D.,
and Patton, J. T.
(1999)
J. Virol.
73,
9934-9943 |
18. | Laue, T. M., Shah, B. D., Ridgeway, T. M., and Pelletier, S. L. (1992) in Analytical Ultracentrifugation in Biochemistry and Polymer Science (Harding, S. E. , Rowe, A. J. , and Horton, J. C., eds) , pp. 90-125, The Royal Society of Chemistry, Cambridge |
19. | Svedberg, T., and Pedersen, K. O. (1940) The ultracentrifuge , Oxford University Press, London |
20. | Laue, T. M., and Stafford, W. F. I. (1999) Annu. Rev. Biophys. Biomol. Struct. 28, 75-100[CrossRef][Medline] [Order article via Infotrieve] |
21. | Rivas, G., Stafford, W., and Minton, A. P. (1999) Methods Companion Methods Enzymol. 19, 194-212[CrossRef] |
22. | Schuck, P., and Braswell, E. H. (2000) in Current Protocols in Immunology (Coligan, J. E. , Kruisbeek, A. M. , Margulies, D. H. , Shevach, E. M. , and Strober, W., eds) , pp. 18.8.1-18.8.22, John Wiley & Sons, Inc., New York, in press |
23. | Johnson, M. L., and Straume, M. (1994) in Modern Analytical Ultracentrifugation (Schuster, T. M. , and Laue, T. M., eds) , pp. 37-65, Birkhäuser, Boston |
24. | Lamm, O. (1929) Ark. Mat. Astr. Fys. 21B, 1-4 |
25. |
Schuck, P.,
and Demeler, B.
(1999)
Biophys. J.
76,
2288-2296 |
26. | Claverie, J.-M., Dreux, H., and Cohen, R. (1975) Biopolymers 14, 1685-1700[Medline] [Order article via Infotrieve] |
27. |
Schuck, P.
(1998)
Biophys. J.
75,
1503-1512 |
28. |
Schuck, P.,
MacPhee, C. E.,
and Howlett, G. J.
(1998)
Biophys. J.
74,
466-474 |
29. | Poncet, D., Lindenbaum, P., L'Haridon, R., and Cohen, J. (1997) J. Virology 71, 34-41[Abstract] |
30. |
Schuck, P.
(2000)
Biophys. J.
78,
1606-1619 |
31. | Darawshe, S., Rivas, G., and Minton, A. P. (1993) Anal. Biochem. 209, 130-135[CrossRef][Medline] [Order article via Infotrieve] |
32. | Murphy, R. M. (1997) Curr. Opin. Biotechnol. 8, 25-30[CrossRef][Medline] [Order article via Infotrieve] |
33. | Provencher, S. W., and Glockner, J. (1981) Biochemistry 20, 33-37[Medline] [Order article via Infotrieve] |
34. | Kuntz, I. D. (1971) J. Am. Chem. Soc. 93, 516-518[Medline] [Order article via Infotrieve] |
35. |
Perugini, M. A.,
Schuck, P.,
and Howlett, G. J.
(2000)
J. Biol. Chem.
275,
36758-36765 |
36. | Patton, J. T. (2001) Gastroenteritis Viruses , pp. 64-77, John Wiley & Sons, Chichester, United Kingdom |
37. |
Soengas, M. S.,
Mateo, C. R.,
Rivas, G.,
Salas, M.,
Acuna, A. U.,
and Gutierrez, C.
(1997)
J. Biol. Chem.
272,
303-310 |
38. | Misselwitz, R., Welfle, K., Curth, U., Urbanke, C., and Welfle, H. (1995) J. Biomol. Struct. Dyn. 12, 1041-1054[Medline] [Order article via Infotrieve] |
39. | Tucker, P. A., Tsernoglou, D., Tucker, A. D., Coenjaerts, F. E., Leenders, H., and van der Vliet, P. C. (1994) EMBO J. 13, 2994-3002[Abstract] |
40. | Folmer, R. H., Nilges, M., Konings, R. N., and Hilbers, C. W. (1995) EMBO J. 14, 4132-4142[Abstract] |
41. |
Passy, S. I., Yu, X.,
Li, Z.,
Radding, C. M.,
Masson, J.-Y.,
West, S. C.,
and Egelman, E. H.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
10684-10688 |
42. |
Egelman, E. H., Yu, X.,
Wild, R.,
Hingorani, M. M.,
and Patel, S. S.
(1995)
Proc. Natl. Acad. Sci. U. S. A.
92,
3869-3873 |
43. | Curth, U., Urbanke, C., Greipel, J., Gerberding, H., Tiranti, V., and Zeviani, M. (1994) Eur. J. Biochem. 221, 435-443[Abstract] |
44. | Raghunathan, S., Kozlov, A. G., Lohman, T. M., and Waksman, G. (2000) Nat. Struct. Biol. 7, 648-652[CrossRef][Medline] [Order article via Infotrieve] |
45. |
Li, K.,
and Williams, R. S.
(1997)
J. Biol. Chem.
272,
8686-8694 |
46. | Genschel, J., Litz, L., Thole, H., Roemling, U., and Urbanke, C. (1996) Gene (Amst.) 182, 137-143[CrossRef][Medline] [Order article via Infotrieve] |
47. | Yang, C., Curth, U., Urbanke, C., and Kang, C. (1997) Nat. Struct. Biol. 4, 153-157[Medline] [Order article via Infotrieve] |
48. | Ferrari, M. E., Fang, J., and Lohman, T. M. (1997) Biophys. Chem. 64, 235-251[CrossRef][Medline] [Order article via Infotrieve] |
49. |
Tseng, M.,
Palaniyar, N.,
Zhang, W.,
and Evans, D. H.
(1999)
J. Biol. Chem.
274,
21637-21644 |
50. | Lohman, T. M., and Ferrari, M. E. (1994) Annu. Rev. Biochem. 63, 527-570[CrossRef][Medline] [Order article via Infotrieve] |