Structural mobility of the monomeric C-terminal domain
of the HIV-1 capsid protein
Luis A. Alcaraz
1,
*, Marta del A
´
lamo
2
, Mauricio G. Mateu
2
and Jose
´
L. Neira
1,3,
*
1 Instituto de Biologı
´
a Molecular y Celular, Universidad Miguel Herna
´
ndez, Elche (Alicante), Spain
2 Centro de Biologı
´
a Molecular ‘Severo Ochoa’ (CSIC-UAM), Universidad Auto
´
noma de Madrid, Spain
3 Biocomputation and Complex Systems Physics Institute, Zaragoza, Spain
Dynamic processes in proteins contribute toward defin-
ing their structure and function, including protein fold-
ing, association and ligand binding [1]. The main
challenge in all structural and dynamic studies is to
find a relationship between the structural and mobility

a Molecular y
Celular, Edificio Torregaita
´
n, Universidad
Miguel Herna
´
ndez, Avenida del Ferrocarril
s ⁄ n, 03202 Elche (Alicante), Spain
Fax: +34 966 658 758
Tel: +34 966 658 459
E-mail:
*These authors contributed equally to this
work
(Received 4 February 2008, revised 22 April
2008, accepted 24 April 2008)
doi:10.1111/j.1742-4658.2008.06478.x
The capsid protein of HIV-1 (p24) (CA) forms the mature capsid of the
human immunodeficiency virus. Capsid assembly involves hexamerization
of the N-terminal domain and dimerization of the C-terminal domain of
CA (CAC), and both domains constitute potential targets for anti-HIV
therapy. CAC homodimerization occurs mainly through its second helix,
and it is abolished when its sole tryptophan is mutated to alanine. This
mutant, CACW40A, resembles a transient monomeric intermediate formed
during dimerization. Its tertiary structure is similar to that of the subunits
in the dimeric, non-mutated CAC, but the segment corresponding to the
second helix samples different conformations. The present study comprises
a comprehensive examination of the CACW40A internal dynamics. The
results obtained, with movements sampling a wide time regime (from pico-
to milliseconds), demonstrate the high flexibility of the whole monomeric
protein. The conformational exchange phenomena on the micro-to-milli-

the monomeric CAC mutant Trp184Ala, CACW40A,
resembles a transient monomeric intermediate formed
during dimerization [24,25]. In the present study, for
sake of clarity, the mutant is referred to as CACW40A
to denote the position of the mutation in the C-termi-
nal domain; in addition, the amino acids of
CACW40A are numbered from its first residue (i.e. the
added N-terminal methionine is Met1, and the second
residue is Ser2, which corresponds to Ser146 in the
numbering of the intact CA). The CACW40A protein
is monomeric, and its structure is similar to that of the
subunits in the dimeric, non-mutated CAC, but, in
the monomeric form, the segment corresponding to the
second helix samples different conformations [26]
(Fig. 1). At the end of this region, several hydrophobic
residues are buried and, as a consequence, the last two
helices are rotated compared to their position in
dimeric non-mutated CAC. Thus, from a structural
point of view, only the dimerization interface has
substantially changed.
To determine whether the apparent dynamic charac-
ter of this region is shown by other polypeptide
patches, we have studied the dynamics of monomeric
CACW40A. Flexibility is often associated with inter-
faces, and it is well known that complex formation
(either in an oligomer or in a more simple substrate–
enzyme reaction) can lead to conformational and
dynamic changes at some, if not all, of the residues
involved [27]. In our previous description of the struc-
ture of CACW40A, we observed a high flexibility in

mentary Table S1). Residues in the first a -helix pre-
sented a mean of 2.90 s
)1
(range 2.56–3.69); the second
a-helix presented a mean of 3.06 s
)1
(range 2.79–3.15);
the third a-helix presented a mean of 2.78 s
)1
(range
2.26–3.05); and, finally, amino acids in the loop region
presented a mean of 3.18 s
)1
(range 2.91–3.47). There
Fig. 1. Structure of CACW40A. UCSF CHIMERA software was used to
render the model from the 2JO0 Protein Data Bank deposited
structure: the first a-helix is in blue; the second one in green; and
the last a-helix is shown in yellow. The single turn of a 3
10
-helix at
the N-terminus of the protein is shown in red.
Dynamics of monomeric CAC L. A. Alcaraz et al.
3300 FEBS Journal 275 (2008) 3299–3311 ª 2008 The Authors Journal compilation ª 2008 FEBS
was no clear correlation between the elements of sec-
ondary structure and the values of R
1
. Similar findings
have been found in proteins of similar size at the same
magnetic field, such as eglin c [32,33], CI2 [34], and the
GAL4 domain [35,36].

in
CACW40A were clearly higher than those of other pro-
teins of similar size measured at the same magnetic field
(eglin c, CI2 or GAL4 with average values of 5.6, 6 and
8s
)1
, respectively [32–36]; GAL4 is the most disordered
protein, and thus shows the highest values of R
2
).
The mean of the nuclear Overhauser effect (NOE) in
CACW40A was 0.60 (range 0.28–0.87) (Fig. 2C; see
also supplementary Table S1). This mean is lower than
the value of 0.79 expected from theoretical consider-
ations at a field strength of 11.7 T [37]. These results
(together with those of the R
2
described above) suggest
a high flexibility of the whole backbone of CACW40A;
interestingly, a study of dynamics of the C-terminal
region of dimeric CAC also shows low NOE values
[38], and extensive signal broadening has been observed
in the assignment of dimeric non-mutated CAC [31].
The residues with low NOE values (< 0.65) in
CACW40A were Ile9 (at the C-cap of the 3
10
-helix);
Tyr20 (at the beginning of the first helix); Lys26 and
Ala30 (at the C-cap of the first helix); Val37 (in the mid-
dle of the long disordered loop); Thr44, Val47 and

to avoid any potential error in the
determination of the model-free parameters.
Fig. 2. Relaxation rates of CACW40A. The relaxation rates are
shown for (A) R
1
, (B) R
2
and (C)
15
N-
1
H NOE for CACW40A at
11.7 T. Sample conditions were 293 K, pH 7.0 in 0.1
M phosphate
buffer. The cylinders at the top of each panel indicate the three
a-helices.
L. A. Alcaraz et al. Dynamics of monomeric CAC
FEBS Journal 275 (2008) 3299–3311 ª 2008 The Authors Journal compilation ª 2008 FEBS 3301
We first estimated the s
m
with tensor2, by using the
subset of rigid residues (see Experimental procedures),
yielding a value of 6.4 ± 0.1 ns.
The s
m
was also determined by using the approach
developed by Wagner et al. [35,36]. Briefly, this
method assumes that, if the re-orientation of an inter-
nuclear
15

J(0) (i.e. the spectral density function at 0 MHz)
(Fig. 3). In CACW40A, the positive solutions to the
cubic equation lead to 1.28 ± 0.03 ns and
7.6 ± 0.6 ns. The first root is assigned to an internal
motion of the protein, and the second is the overall
tumbling of the molecule, which is close to the value
obtained previously. As can be observed, only a small
number of the experimental points in CACW40A are
close to the crossing point, demarcating the s
m
bound-
ary of the theoretical Lorentzian curve for the spectral
density function. Experimental points close to the
boundary imposed by the theoretical curve correspond
to residues with fast internal dynamic contributions,
whereas those undergoing slower dynamics are located
at J(0) values above the limit of the correlation time,
as occurs in CACW40A (Fig. 3).
We also used different theoretical approaches to
estimate the s
m
[44,45], and the results are similar to
those described above (data not shown). The value
used in the model-free formalism (see below) was
6.4 ± 0.1 ns. It is important to indicate that relaxation
measurements of the dimeric, non-mutated CAC have
been carried out, and the s
m
obtained is much higher
than that reported here [46].

0.56 ± 0.29 (see supplementary Table S2). This num-
ber is significantly lower than the average value of 0.86
found in other proteins [47], probably due to the long
loop in CACW40A, which is not very well hydrogen-
bonded to the rest of the structure [48].
None of the residues in CACW40A, except Ala65,
could be fitted to the simplest model of tensor2 (see
supplementary Table S2). Residues Glu15, Lys26,
Gly62, Ala73 and Gly81 could be fitted to the second
model. Amino acids Phe17, Asp19, Arg23 and Gly79
could be analysed with the third one, where an
exchange contribution, R
ex,
is included. Residues
Gln11, Thr42 and Thr72 were fitted to the fifth model;
and the remaining residues could be analysed accord-
ing to the fourth model, where R
ex
contributions and
fast movements are included. A large number of resi-
dues (i.e. those fitted to models three and four) did
experience conformational exchange on a micro-to-
millisecond time scale (Fig. 4B).
In conclusion, most of the residues in CACW40A,
and not only those in the loop region, have a fast
internal mobility. Furthermore, the fast internal corre-
Fig. 3. Relationship between J(x
N
) and J(0). The theoretical varia-
tion between both parameters assuming a simple Lorentzian curve

2
, s
e
and R
ex
to those reported in the supplementary
(Table S2) were observed when a fully anisotropic
model was used (data not shown). All these findings
suggest that the assumptions of the model-free
approach are no longer valid in CACW40A (i.e. it is
not possible to separate the overall tumbling of the
molecule and the local fast movements of each
15
N-
1
H
bond). Thus, although the model-free approach is very
intuitive, we decided to use the reduced spectral inten-
sity formalism to test whether our results (i.e. large
mobility through all the elements of structure) were
not an artifact of the model-free approach.
Reduced spectral density approach
This approach provides insights into the motion of the
N–H bond vector at three selected frequencies, x
0
(= 0), x
N
and 0.87x
H
(Fig. 5).

comparison of Tables S2 and S3 in the supplementary
material shows that all residues with J(0) values higher
than 3.2 ns did show a R
ex
contribution in the model-
free approach. These residues were Gly12, Lys14,
Phe17, Asp19, Tyr20, Val21, Arg23, Tyr24, Thr27,
Glu31, Val37, Met41, Thr44, Gln48, Asn49, Ala50,
Asp53 to Leu58, Leu67, Met70, Met71 and Gln75.
Because J(x
N
) (i.e. the spectral density function at
the Larmor frequency of the
15
N) and J(0.87x
H
) (i.e.
the spectral density function at the 0.87 times the Lar-
mor frequency of the
1
H) are independent of R
2
[see
Eqns (3,4) in Experimental procedures] and less sensi-
tive than J(0) to the distribution of correlation times,
they can provide insights into protein dynamics. The
mean value of J(x
N
) was 0.58 nsÆrad
)1

are shown on the structure of the protein:
10 < R
ex
<16s
)1
(red); 5 < R
ex
<10s
)1
(orange) and 0 < R
ex
<5s
)1
(blue).
L. A. Alcaraz et al. Dynamics of monomeric CAC
FEBS Journal 275 (2008) 3299–3311 ª 2008 The Authors Journal compilation ª 2008 FEBS 3303
for a 1% of J(0) (Fig. 5B). The mean value was
0.0138 nsÆrad
)1
(range 0.00138–0.0245) (see supplemen-
tary Table S3). The tendency in J(0.87x
H
) was the
opposite to that observed in J(0): the highest values in
J(0.87x
H
) correspond to the termini of the helices and
the regions in between, indicating efficient picosecond
averaging.
In conclusion, using the reduced spectral density

N backbone dynamics is
to predict regions of a protein with sufficient potential
flexibility to allow functional events to occur (binding,
conformational changes or catalysis). However, experi-
ments with several dozens of proteins [27] demonstrate
that there is no easy and general correspondence
between the order parameter (S
2
), the spectral density
function [J(x)] and the secondary structural elements
of a protein. Furthermore, there are no simple rules
for the interpretation of the exchange rates (R
ex
)or
the different correlation times (s
m
, s
s
or s
f
).
In CACW40A, although the helical elements have
the highest order parameters, there is no relationship
between S
2
and the location of structural elements
(Fig. 4). Furthermore, the R
ex
terms are distributed
throughout the 3D structure of the protein, and most

3304 FEBS Journal 275 (2008) 3299–3311 ª 2008 The Authors Journal compilation ª 2008 FEBS
a much slower time regime than the other helices.
These equilibria also occur in the other two helices, as
shown by the exchange pattern [26], although they are
less well-ordered, as judged by the lower S
2
.
The types of movements and the residues involved
are described below.
The pico-to-nanosecond dynamics
Residue Ala65 (at the N-terminus of the third helix) is
the sole residue that has restricted internal dynamics
(model-free formalism). Fast internal dynamics (i.e.
residues with at least another tumbling time) occurs at
the N (Gln11 and Glu15) and at the C-termini of the
first a-helix (Lys26); in the long disordered loop
(Thr42); and at the N- (Gly62, Ala64), and C-termini
of the third helix (Thr72, Ala73). However, it is not
possible to establish any correlation between any
structural parameter of those residues and the fast
dynamics observed.
The micro-to-millisecond dynamics
Most of the residues in CACW40A required an R
ex
term (model-free formalism) or had long J(0) values
(reduced-spectral approach); furthermore, most of the
residues in the loop (which forms the second helix in
the dimeric non-mutated CAC protein [22,23]) were
broad beyond detection in the HSQC experiments [26].
Although the arguments could be considered as specu-

function (the function describing the movement) of each
bond vector is decomposed as the product of the corre-
lation function for overall (global) and internal (local)
motions (i.e. the internal motions of the bond vectors
are independent of the overall rotational movement of
the molecule). Furthermore, the internal motions of
each bond vector are independent of each other, but the
rotational diffusion of the molecule affects each of those
bond vectors identically [42,43]. On the other hand,
spectral density mapping makes no assumptions about
the nature of the rotational diffusion (i.e. the informa-
tion on which oscillations for a particular bond vector
are associated with global molecular rotation or segmen-
tal molecular motions is lost). Thus, based on the spec-
tral density formalism results, we are unable to discern
whether the movement of each NH bond is due to local
internal or overall tumbling, but we can conclude that
the CACW40A has an intrinsically high structural mobi-
lity (Figs 4 and 5). To support this conclusion, the s
e
s
obtained from the model-free approach for most of the
residues are similar (i.e. they are not faster) than the
overall molecular tumbling of the protein; this means
that we cannot strictly separate the overall tumbling of
the molecule from the internal motions of each bond
vector and, thus, the model-free formalism cannot be rig-
orously applied. This is not the sole example where the
use of the model-free formalism has been unsuccessful:
this approach cannot be applied on natively unfolded

The detection of slow dynamics not only at the dimer-
ization interface (residues Glu31 to Ala40), but also in
the rest of the protein implies the presence of a small
population of pre-existing conformers within the native-
state ensemble. This population interacts with other
CACW40A monomers forming the dimeric CAC, prob-
ably through the side chains of the hydrophobic residues
of the long disordered loop, buried to avoid nonspecific
hydrophobic interactions [26]. There are several exam-
ples of proteins in which binding residues are involved
in slow-exchange processes [27,58], most likely to facili-
tate rapid partner-binding, and the recognition of
several ligands. Internal motions allow amino acids to
explore large regions of the conformational space at a
very low energetic cost, increasing the chances of
successful binding. However, are those slow-exchange
processes responsible, from a thermodynamic point of
view, for the binding of the monomeric species of CAC?
We have previously discussed the variation in the free
energy of binding as a function of the changes in buried
surface area upon dimer formation [59]. On the other
hand, there are no clear correlations between the
enthalpy of binding and the changes in buried surface
area [60]; thus, the only thermodynamic magnitude that
has not been estimated in CAC is the binding entropy
change, DS
b
. The binding entropy, DS
b
, can be divided

ln(T ⁄ 385), where DC
p
is the heat capacity
change of the binding reaction. We have previously
determined the DC
p
()211 ± 10 calÆmol
)1
ÆK
)1
per
monomer) and DS
b
()230 ± 10 calÆmol
)1
ÆK
)1
per
monomer) [59], and then, the contribution from the
conformational flexibility to the entire entropy of
binding will be: DS
con
= )234 calÆmol
)1
ÆK
)1
per mono-
mer. Because, on average, the entropy cost per amino
acid for a folding transition is approximately
5.6 calÆmol

] propionate was obtained from Sigma (Madrid,
Spain). Dialysis tubing was obtained from Spectrapore
(Breda, the Netherlands), with a molecular mass cut-off
of 3500 Da. Standard suppliers were used for all other
chemicals. Water was deionized and purified on a Millipore
(Barcelona, Spain) system.
Protein expression and purification
The
15
N-labelled CACW40A protein was expressed in
Escherichia coli BL21(DE3) in LB and purified as previ-
ously described [26]; the DNA segment used for the mutant
protein encoded for residues 146–231 of CA from HIV-1
(strain BH10) and was cloned as described [24]. The protein
concentration was calculated from A
240
by using the extinc-
tion coefficients of amino acids [64]. Samples were concen-
trated at the desired final NMR concentration by using
Centriprep Amicon devices (Millipore), with a molecular
mass cut-off of 3500 Da.
Protein structure calculations
The determination of the solvent-accessible surface area
was obtained using the VADAR web server [65].
NMR samples
All NMR experiments were acquired on an Avance Bruker
DRX-500 spectrometer (Bruker, Karlsruhe, Germany)
Dynamics of monomeric CAC L. A. Alcaraz et al.
3306 FEBS Journal 275 (2008) 3299–3311 ª 2008 The Authors Journal compilation ª 2008 FEBS
equipped with a triple resonance probe and pulse field

1
rates
using relaxation delays of 50, 100 (· 2), 200, 300, 400, 500,
600, 700 (· 2), 850 and 1000 ms, where the experiments at
100 and 700 ms were repeated twice. The
15
N-T
2
measure-
ments were made using delays of 15, 25 (· 2), 50, 100, 150,
175, 225 (· 2), 300 and 425 ms. For the T
1
and T
2
pulse
sequences, the delay between transients was 5 s. The
1
H-
15
N
NOEs were measured by recording interleaved spectra in the
presence and in the absence of proton saturation. The spec-
trum recorded in the presence of proton saturation was
acquired with a saturation time of 5 s. The spectrum
recorded without proton saturation incorporated a relaxa-
tion delay of 5 s. Each experiment was repeated twice.
Experiments were carried out at two protein concentra-
tions (1 mm and 400 lm) to rule out any possible concen-
tration-dependent effect on the measured relaxation rates,
as has been observed in dimeric non-mutated CAC [46].

Eqn (1) with kaleidagraph software (Abelbeck Software,
Reading, PA, USA).
The steady-state NOE values were determined from the
ratios of the peak intensities with and without proton satu-
ration (i.e. NOE = I
sat
⁄ I
nonsat
). The standard deviation of
the NOE value was determined on the basis of the measured
background noise levels by using the repeated experiments.
The T
1
and T
2
relaxation times (or, R
1
=1⁄ T
1
and
R
2
=1⁄ T
2
) and the NOE enhancement of an amide
15
N
nucleus are dominated by the dipolar interaction of the
15
N

Þ; ð3Þ
Jð0:87x
N
Þ¼ð4r
NH
Þ=ð5d
2
Þ; ð4Þ
and
r
NH
¼ R
1
ðNOE À 1Þðc
N
=c
H
Þ; ð5Þ
where c =(x
N
⁄Ö3)(r
||
– r
^
) and d = l
0
hc
N
c
H

15
N, x
H
is the Larmor frequency of the
1
H, <r> is the
length of the amide bond vector (1.02 A
˚
), and r
||
and r
^
are the parallel and perpendicular components of the CSA
tensor (r
||
)r
^
= )160 p.p.m for a backbone amide [70]).
The uncertainties in a particular J(x) are the quadrature-
weighted sum derived from Eqns (2–5), assuming that
errors in the relaxation rate constants are independent.
Rotational diffusion tensor
An initial estimation of s
m
and the rotational diffusion ten-
sors were obtained with tensor2 [71], from the subset of
residues which accomplished the following criteria [72]: (a)
all residues should have a NOE ‡ 0.65 and (b) the residues
should satisfy:
R

satisfying criteria (a), and r is the standard deviation of:
R
2;i
À
R
2
hi
R
2
hi
À
R
1;i
À
R
1
hi
R
1
hi








The residues which did not accomplish criterion (a) were
Ile9, Lys26, Ala30, Val37, Thr44, Val47, Gln48, Lys55,

defined in terms of: (a) the overall tumbling time, s
m
(in the
order of nanoseconds), and the diffusion anisotropy; (b) the
time scale of internal motions faster than s
m
, the so-called
effective internal correlation time, s
e
(in the pico-to-nano-
second time scale); and (c) the degree of restriction of these
fast internal motions (which is measured by the square of
the order parameter, S
2
). Thus, in residues where the relax-
ation mechanism is dominated by the internal motion (i.e.
residues highly mobile relative to the overall rotational
tumbling), S
2
would approach to zero; on the other hand,
in residues where relaxation is described only by the global
motion of the molecule, S
2
would approach to the unity.
Extensions of this formalism have been developed to incor-
porate two time scales of internal motions or to account
for the effects of slow (micro-to-millisecond time scale) con-
formational exchange; in these cases, the global order
parameter is defined as S
2

e
of each amide proton is very fast and not
relaxation-active; (b) in the second model, the s
e
is relaxa-
tion-active; (c) the third model is identical to the first,
except the conformational (or chemical) exchange on a
microsecond-to-millisecond time scale is taken into account
(by using the R
ex
parameter); (d) the fourth model is identi-
cal to (b), but also includes the R
ex
term; and (e) the fifth
model includes the extension of the formalism, with
two kinds of internal motions: one very fast and other
very slow.
Acknowledgements
We thank the two anonymous reviewers for their help-
ful suggestions and discussions. This work was sup-
ported by grants from Ministerio de Sanidad y
Consumo (MSC) (FIS 01 ⁄ 0004-02), Ministerio de Edu-
cacio
´
n y Ciencia (MEC) (CTQ2005-00360⁄ BQU) and
the private organization FIPSE (Exp: 36557⁄06) to
J. L. N.; grants from MSC (FIS 01 ⁄ 0004-01) and
MEC (BIO2006-00793) and the private organization
FIPSE (Exp: 36557 ⁄ 06) to M. G. M., and by institu-
tional grants from Fundacio

6 Palmer AG, Willimas J & McDermott A (1996) Nuclear
magnetic resonance of biopolymer dynamics. J Phys
Chem 100, 13293–13310.
7 Freed EO (1998) HIV-1 gag proteins: diverse functions
in the virus life cycle. Virology 251, 1–15.
8 Freed EO & Martin M (2001) HIVs and their replica-
tion. In Fields Virology (Knipe DM & Howley PM,
eds), pp. 1971–2041. Lippincott, Philadelphia, PA.
9 Vogt VM (1996) Proteolytic processing and
particle maturation. Curr Top Microbiol Immunol 214,
95–132.
10 Fuller TW, Wilk T, Gowen BE, Kra
¨
usslich H-G &
Vogt VM (1997) Cryo-electron microscopy reveals
ordered domains in the immature HIV-1 particle. Curr
Biol 7, 729–738.
Dynamics of monomeric CAC L. A. Alcaraz et al.
3308 FEBS Journal 275 (2008) 3299–3311 ª 2008 The Authors Journal compilation ª 2008 FEBS
11 Ehrlich LS, Agresta BE & Carter CA (1992) Assembly
of recombinant human immunodeficiency virus type 1
capsid protein in vitro. J Virol 66, 4874–4883.
12 Grob I, Hohenberg H & Kra
¨
usslich H-G (1997) In vitro
assembly properties of purified bacterially expressed
capsid proteins of human immunodeficiency virus. Eur
J Biochem 249, 592–600.
13 Grob I, Hohenberg H, Huckangel C & Kra
¨

HIV-1 capsid protein. Nat Struct Biol 3, 763–770.
20 Gitti RK, Lee BM, Walker J, Summers MF, Yoo S &
Sundquist WI (1996) Structure of the amino-terminal
core domain of the HIV-1 capsid protein. Science 273,
231–235.
21 Gamble TR, Vajdos FF, Yoo S, Worthylake DK, Hou-
seweart M, Sundquist WI & Hill CP (1996) Crystal
structure of human cyclophilin A bound to the amino-
terminal domain of HIV-1 capsid. Cell 87, 1285–1294.
22 Gamble TR, Yoo S, Vajdos FF, von Schwedler UK,
Worthylake DK, Wang H, McCutcheon JP, Sundquist
WI & Hill CP (1997) Structure of the carboxyl-terminal
dimerization domain of the HIV-1 capsid protein.
Science 278, 849–853.
23 Worthylake DK, Wang H, Yoo S, Sundquist WI & Hill
CP (1999) Structures of the HIV-1 capsid protein
dimerization domain at 2.6 A
˚
resolution. Acta Crystal-
logr D 55, 85–92.
24 Mateu MG (2002) Conformational stability of dimeric
and monomeric forms of the C-terminal domain of
human immunodeficiency virus-1 capsid protein. J Mol
Biol 318, 519–531.
25 del Alamo M, Neira JL & Mateu MG (2003) Thermo-
dynamic dissection of a low-affinity protein-protein
interface involved in human immunodeficiency virus
assembly. J Biol Chem 278, 27923–27929.
26 Alcaraz LA, del A
´

32274–32279.
32 Peng JW & Wagner G (1992) Mapping of the spectral
densities of N-H bond motions in eglin c using heter-
onuclar relaxation experiments. Biochemistry 31, 8571–
8586.
33 Peng JW & Wagner G (1995) Frequency spectrum of
NH bonds in eglin c from spectral density mapping at
multiple fields. Biochemistry 34, 16733–16752.
34 Shaw GL, Davis B, Keeler J & Fersht AR (1995) Back-
bone dynamics of chymotrypisn inhibitor 2: effect of
breaking the active site bond and its implications for
the mechanism of inhibition of serine proteases. Bio-
chemistry 34, 2225–2233.
35 Lefe
`
vre JF, Dayie KT, Peng JW & Wagner G (1996)
Internal mobility in the partially folded DNA binding
and dimerization domains of GAL4: NMR analysis of
the N-H spectral density functions. Biochemistry 35,
2674–2686.
36 Dayi KT, Wagner G & Lefe
`
vre JF (1996) Theory and
practice of nuclear spin relaxation in proteins. Annu
Rev Phys Chem 47, 243–282.
37 Perez-Can
˜
adillas JM, Guenneugues M, Campos-Olivas
R, Santoro J, Martı
´

44 Daragna VA & Mayo KH (1997) Motional analyses of
protein and peptide dynamics using
13
C and
15
N NMR
relaxation. Prog NMR Spectrosc 31, 63–105.
45 Krishnan VV & Cosman M (1998) An empirical rela-
tionship between rotational correlation time and solvent
accessible surface area. J Biomol NMR 12, 177–182.
46 Stitch J, Humbert M, Fidlow S, Bodem J, Mu
¨
ller B,
Dietrich U, Werner J & Kra
¨
usslich H-G (2005) A pep-
tide inhibitor of HIV-1 assembly in vitro. Nat Struct
Mol Biol 12, 671–677.
47 Goodman JL, Pagel MD & Stone MJ (2000) Relation-
ships between dynamics from a database of NMR-
derived backbone order parameters. J Mol Biol 295,
963–978.
48 Akerud T, Thulin E, van Etten RL & Akke M (2002)
Intramolecular dynamics of low molecular weight pro-
tein tyrosine phosphatase in monomer-dimer equilib-
rium studied by NMR: a model for changes in
dynamics upon target binding. J Mol Biol 322, 137–152.
49 Tjandra N, Feller SE, Pastor RW & Bax A (1995)
Rotational diffusion anisotropy of human ubiquitin
from

ics of ribonuclease H: temperature dependence of
motions on multiple time scales. Biochemistry 35,
16009–16023.
55 Bracken C, Carr PA, Cavanagh J & Palmer AG (1999)
Temperature dependence of intramolecular dynamics of
the basic leucine zipper of GCN4: implications for the
entropy of association with DNA. J Mol Biol 285,
2133–2146.
56 Gu
¨
mral D, Nadalim L, Corazza A, Fogolari F, Daman-
te G, Viglino P & Esposito G (2008) Helix mobility and
recognition function of the rat thyroid transcription fac-
tor 1 homeodomain-hints from
15
N-NMR relaxation
studies. FEBS J 275, 435–448.
57 Ivanov D, Stone JR, Maki JL, Collins T & Wagner G
(2005) Mammalian SCAN domain dimer is a domain
swapped homolog of the HIV capsid C-terminal
domain. Mol Cell 17, 137–143.
58 Botuyan MV, Mer G, Yi G-S, Koth CM, Case D,
Edwards AM, Chazin WJ & Arrowsmith CH (2001)
Solution structure and dynamics of yeast elongin C in
complex with a von-Hippel-Lindau peptide. J Mol Biol
312, 177–186.
59 Lido
´
n-Moya MC, Barrera FN, Bueno M, Pe
´

server for quantitative evaluation of protein structure
quality. Nucleic Acids Res 31, 3316–3319.
66 Cavanagh JF, Wayne J, Palmer AG & Skelton NJ
(1996) Protein NMR Spectroscopy: Principles and Prac-
tice, pp. 167–168. Academic Press, San Diego, CA.
67 Farrow NA, Muhandiram R, Singer AU, Pascal SM,
Kay CM, Gish G, Shoelson SE, Pawson T, Forman-
Kay JD & Kay LE (1994) Backbone dynamics of a free
and phosphopeptide-complexed src homology 2 domain
studied by
15
N NMR relaxation. Biochemistry 33, 5984–
6003.
68 Abragam A (1961) The Principles of Nuclear Magne-
tism. Clarendon Press, Oxford.
69 Farrow NA, Zhang O, Szabo A, Torchia DA & Kay
LE (1995) Spectral density function using
15
N relaxa-
tion exclusively. J Biomol NMR 6, 153–162.
70 Hiyama Y, Niu C, Silverton JV, Bavaso A & Torchia
DA (1988) Determination of the
15
N chemical shift ten-
sor via
15
N-
2
H dipolar coupling in Boc-glycylglycyl
[