 |
INTRODUCTION |
Protein-protein interactions control many critical functions in
biology, ranging from tight binding antibody-antigen recognition events
to transient interactions between enzymes in a signaling pathway. These interactions can be complex; there are sometimes a
number of diverse proteins that can interact with a particular target
molecule (1, 2). Elucidation of key intermolecular contacts between
protein partners can aid in the development of small molecule
inhibitors and/or promoters of these important interactions, which in
turn control function. A particularly interesting area in biology today
is the investigation of the molecular mechanisms of the
assembly/disassembly of signaling networks in response to a specific
cellular signal (3). Indeed, the spatio-temporal compartmentalization
of signaling molecules affords biological control by poising
interacting partners in close proximity to substrate(s) and/or
regulatory elements (4).
Targeting of the cyclic AMP-dependent protein kinase
(PKA)1 holoenzyme through
interactions with A kinase
anchoring proteins (AKAPs) has
emerged as an important modulator of PKA activity in diverse tissues
(5). The PKA holoenzyme consists of a regulatory subunit
(R2) dimer and two catalytic (C) subunits (6).
Phosphorylation of target proteins is carried out by the C subunit,
whereas the N-terminal 45 residues of the R subunit mediates both
dimerization and subcellular localization via AKAP recognition (7, 8). Hence, the N-terminal functional domain is termed a
D/D motif because it dimerizes and
docks to anchoring partners. Solution structural studies
revealed that the type II
D/D of PKA packs into an antiparallel,
dimeric X-type four-helix bundle, with a surface-exposed hydrophobic
groove that is the site of anchoring interactions (9, 10).
PKA interacts with a diverse family of proteins. Sequence alignment of
the identified AKAPs, to date, reveals no specific recognition sequence
for the D/D. However, a conserved structure consistent with an
amphipathic helix was predicted, and has been demonstrated in recent
solution structural studies of a peptide derivative of the prototypic
AKAP human thyroid anchoring protein Ht31 (residues 493-515 and
designated Ht31pep) (7, 11, 12). This peptide derivative of
Ht31 exhibits a nanomolar binding affinity for the type II D/D (12, 13)
via hydrophobic-hydrophobic interactions between the surface-accessible
hydrophobic groove on the D/D and the hydrophobic face of the
AKAP-derived amphipathic helix (10, 14).
In an effort to gain a better understanding of the physicochemical
basis for PKA-AKAP interactions, we initiated hydrogen/deuterium (H/D)
exchange and backbone relaxation studies of the D/D free and in complex
with Ht31pep. In contrast to recent work described by
Powell et al. (15) using H/D exchange to measure
ligand-binding affinities, we observe only modest changes in the H/D
protection factors upon complex formation, despite the nanomolar
binding affinity of Ht31 for the D/D (11). Unexpectedly, we also find that backbone flexibility in the binding interface of the D/D increases in the Ht31pep complex. We propose
that the increase in backbone mobility and display of modest changes in
H/D exchange protection factors upon high affinity ligand binding may
be a general effect observed for proteins that use solvent accessible hydrophobic surfaces to recognize diverse binding partners.
 |
EXPERIMENTAL PROCEDURES |
Sample Preparation--
The D/D·Ht31pep
peptide complex was prepared as described previously (16). The
Ht31pep peptide was obtained from PeptidoGenic Research and
Co. (Livermore, CA). The stoichiometry of binding for classical AKAPs
is one AKAP per R subunit dimer (11). The apo-D/D was prepared as a
0.25 mM (0.5 mM monomer) sample in 20 mM sodium phosphate buffer, 90% H2O, 10%
D2O, pH 4.0. Relaxation experiments were collected on
either a 0.25 mM (0.50 mM monomer apo) or a
0.50 mM (1.0 mM monomer complex) sample.
Samples for hydrogen/deuterium exchange studies were collected on
0.25-1.0 mM protein solutions.
Hydrogen/Deuterium Exchange
Studies--
Hydrogen/deuterium exchange experiments were initiated by
introducing the protein samples into deuterated buffer via a QuikChange gel chromatography step (Roche Molecular Biochemicals). A series of two-dimensional 1H-15N heteronuclear single
quantum coherence spectra (17) were collected on a Bruker DMX500
spectrometer at 0, 16, 30, 60, 100, 300, 1080, 1560, and 2880 min after
the introduction of the sample into D2O buffer. Calibration
of the individual spectra to correct for protein concentration
differences between samples was achieved by normalizing the data to the
intensity of the non-exchanging aliphatic resonance of
Val20 H
2* in a one-dimensional 1H spectrum
taken directly after the completion of the heteronuclear single quantum
coherence experiments.
Analysis of Kinetic Data--
The time-dependent
change in the cross-peak intensity (volume) of each amide proton
resonance was found to be exponential. Fitting of each of the observed
decay curves to an exponential decay function allowed the extraction of
the residue specific experimental exchange rate,
kobs, according to,
|
(Eq. 1)
|
where I is the observed cross-peak
intensity (volume) at time t. Fitting of data, whether
normalized or non-normalized, volume or intensity, from 600 or 500 MHz
spectrometer gave identical rates (data not shown).
Protection Factors--
Residue-specific protection factors,
P, for individual amide protons were calculated from the
following relationship,
|
(Eq. 2)
|
where P is the protection factor,
kint the intrinsic rate corrected for local
sequence variations (18), and kobs the observed rate for the solvent exchange of the amide proton.
Thermodynamic Analysis--
A general form of the
hydrogen-deuterium exchange mechanism can be described as (19),
|
(Eq. 3)
|
where H and D denote protonated and
deuterated backbone amides, C the "closed" form and O the
"open" form, and kcl and
kop are their corresponding rate constants. The
intrinsic rate constant for the chemical exchange reaction,
kint, for a specific amide proton depends upon
the local primary sequence, pH, and temperature under which exchange
takes place (18, 20). Given this model, the rate constant for the
exchange, kex, is given by the following relationship.
|
(Eq. 4)
|
When the chemical exchange step is much faster than
the rate constant for reprotection, we approach the EX1 limit,
kex = kop. When the
closing step, kcl, is faster than the exchange
rate, kint, then kex
reduces to kex = Kop × kint, where Kop
is the equilibrium constant for the opening reaction. This is known as
the EX2 limit. Under these conditions and assuming a well defined
native-state ensemble, an apparent free energy of exchange,
G
, can be estimated from the
calculated protection factors according to the following
relationship,
|
(Eq. 5)
|
where R is the gas constant, T the
temperature in Kelvin, and P is the residue-specific
protection factor (P = kex/kint) (20).
NMR Relaxation--
Relaxation experiments were collected at
25 °C on Bruker DMX500 and DRX600 spectrometers using a
triple-resonance gradient probe. The 15N T1,
15N T2, and NOE measurements were acquired with
established methods that use pulsed-field gradients for coherence
transfer pathway selection combined with sensitivity enhancement
(21-23). The 15N T1 and 15N
T2 relaxation experiments were collected as a time series
of two dimensional 1H-15N correlation spectra,
with variable delay times (40, 100*, 200, 300, 400*, 500, 600, 800, 100, and 1280 ms for T1 and 10, 18, 26, 38, 50*, 62, 78, 98, and 122 ms for T2), where asterisks indicate duplicate
points to estimate the error in the measured intensities.
Fitting of the Relaxation Data--
Data were processed using
the program FELIX 97.0 (Molecular Simulations Inc.) and the intensities
for the amide 1H-15N cross-peaks were assessed
with relax_scripts (62). The
R1 and R2 relaxation
rates values were determined by fitting the time series to a single
exponential decay function. The errors in the rates are reported as the
95% confidence limits in the kinetic fits.
1H-15N steady-state NOE values were determined
from the ratio of the intensities of the respective cross-peaks with
and without proton saturation. Errors were assessed both from replicate
experiments and from measurements of the root mean square values of the
noise in the spectra (21).
Model Free Analysis--
Relaxation of an amide 15N
nucleus is dominated by dipolar coupling with the attached proton, and
anisotropy of the 15N chemical shift tensor. Dynamics of
the NH bond axis are characterized by the spectral density function,
J(
), which is related to the three relaxation parameters
R1, R2, and NOE (24). The
model-free formalism (25, 26) allows the assessment of the amplitudes and time scales of the intramolecular motions by modeling the spectral
density function, J(
), in terms of the order parameter S2 (characterizing the amplitude of internal
motions of each NH bond),
e (the effective correlation
time for internal motions), and
m (the isotropic
rotational correlation time of the protein). For an axially symmetric
rotational diffusion tensor (27, 28), the spectral density
J(
) can be expressed as a function of the angle between
the N-H bond vector and the unique axis of the principal frame of the
diffusion tensor. We have followed standard protocols (with fitting
data acquired at 500 and 600 MHz simultaneously) for selection of a
dynamical model describing internal motions for each residue (26).
Once the model selection was completed, the parameters characterizing
overall molecular tumbling and the internal motional parameters were
optimized simultaneously. All optimizations involved minimization of
the
2 function (29). The model free calculations were
performed using the program modelfree (version 4.1), (also
r2r1_diffusion and pdbinertia) kindly provided by
Dr. Arthur G. Palmer. Protomer-specific assignments were used for the
D/D in the Ht31pep complex except for residues 33, 35, and
36, where it was not possible to obtain unambiguous protomer-specific
assignments (10). In the latter cases, Model Free analysis was
performed by assigning all possible protomer-specific assignments to
resonances (residues 33, 35, and 36). No significant differences were
observed in the resulting parameters for residues 33, 35, and 36.
 |
RESULTS |
Hydrogen/Deuterium Exchange--
Of a total of 44 residues in the RII
D/D protomer, five (Thr10,
Gln23, Phe31, Thr37, and
Glu41) in the apo and three (Phe31,
Thr37, and Glu41) residues in the Ht31 complex
were excluded from analysis because of weak signal intensity and/or
resonance overlap. Because the non-palindromic peptide
Ht31pep binds to RII
in a one peptide to one dimer
stoichiometry, it induces asymmetry into the complex resulting in
protomer-specific chemical shift changes in the D/D dimer upon peptide
binding (10, 16). Residues on both the exterior and buried faces of helices I/I' and II/II' experience reduction in hydrogen exchange kinetics upon peptide binding (Table I
and Fig. 1b). The largest
changes in amide exchange rate upon AKAP binding occur in helix I/I'
(Leu12/12' through Arg22/22'), which make up
the hydrophobic cleft that is the site of direct AKAP interaction.
Mutagenesis studies indicated that residues Ile3/3' and
Ile5/5' contribute important determinants for
Ht31pep binding (11), but interestingly these residues are
freely exchanging in both the apo and the
D/D·Ht31pep complex. Helices II/II'
(Ala32/32'-Leu39/39') make up the bottom
surface of the molecule and are removed from the AKAP binding surface.
Nonetheless, increases in amide-proton exchange protection factors are
also observed in this region upon complex formation (Fig.
1b).

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Fig. 1.
a, differences in order
parameters observed between the free and the complexed forms of
RII -(1-44). The data between the dashed lines
correspond to changes that are not statistically significant (smaller
than the root mean square error). Values for protomers A and B are
indicated by filled triangles and filled circles,
respectively. Contact residues are indicated by a cross in
the plot. b, differences in the protection factors
(solid line) and surface accessible area (red dotted
line), observed between the free and the complexed forms of
RII -(1-44). Contact residues are indicated by a cross in
the plot. The surface accessible area was calculated with MolMol
(61).
|
|
Relaxation--
Backbone 1H-15N resonance
assignments of the apo- and Ht31pep-bound D/D were
determined as described previously (16). Relaxation rate parameters
R1, R2, and NOE were
obtained from the analysis of proton detected 15N and
1H correlation spectra of the free and the
D/D·Ht31pep complex. Data were collected at
two magnetic fields, 500 and 600 MHz. 52 of 56 amide proton resonance
cross-peaks were of sufficient quality for the reliable quantitation of
the cross-peak intensities in the individual spectra. The relaxation
parameters are given in Table II. We
observe a decrease in the NOE value upon complex formation
for most of the residues located in the hydrophobic binding groove, and
a concomitant increase in the NOE value for the biologically important
residues Ile3 and Ile5 (13, 14), among others.
These compensatory increases/decreases upon complex formation leads to
a system where the domain average NOE values are the same
for the free and Ht31pep complex, as determined at 500 and
600 MHz (Table II and Fig. 2). Comparison
of the R1 relaxation rates for the apo and
Ht31pep complex shows a field dependence to the observed
domain average rates. The average value for the
free is larger than that observed for the complex, when measured at 500 MHz, but is within error as assessed at 600 MHz. The
average R2, relaxation rates for the
free and the complex (at 500 and 600 MHz) are within
experimental error, but individual residue rates show variable changes
upon complex formation, suggesting the presence of low frequency
motions, including possible conformation exchange.

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Fig. 2.
Plots of the measured 15N
relaxation parameters (heteronuclear NOEs, longitudinal and transverse
relaxation rate constants, R1, and
R2, respectively) and their uncertainties
as a function of residue number for free (open
triangles) and Ht31pep-bound
RII -(1-44) (filled triangles
and circles, corresponding to protomer A and B,
respectively). Panels a-c and d-f
correspond to data measured at 500 and 600 MHz, respectively.
|
|
Model Free Analysis of Relaxation Parameters (R1,
R2, and NOE)--
We have performed a Model Free analysis
(25) of the data in an effort to interpret relaxation parameters in
terms of dynamical variables. Chemical shift splitting is observed in
the D/D·Ht31pep complex and protomer-specific
assignments were used where available (see "Experimental
Procedures") (16). The observed residue-specific relaxation
parameters (Fig. 2) show an overall (protomer A versus protomer B) similar behavior, thus we included a total of 52 residues in our analysis of the complex.
The experimental data (500 and 600 MHz) were examined assuming either
an isotropic or an anisotropic axially symmetric molecular tumbling
model. The diffusion tensors of overall reorientation were calculated
from the R2/R1 ratios
(those within one standard deviation of the average (30)) using the
program r2r1_diffusion (provided by Dr. Arthur G. Palmer) and the atomic coordinates of the free or
Ht31pep·D/D complex, respectively (16). The
calculated ratio of diffusion tensor components were
2Dzz/(Dxx + Dyy) = D||/D
= 0.97 ± 0.02 (free) and D||/D
= 1.16 ± 0.02 (Ht31pep complex). Thus, the isotropic
model adequately describes the overall reorientation and was used
for further analysis.
The overall rotational times,
m, calculated from
R2/R1 were used as the
initial input values and the final optimization yielded the isotropic
correlation times,
m, of 7.94 ± 0.04 and 8.25 ± 0.04 ns for the free and D/D·Ht31pep complex,
respectively. A simultaneous fit of the data acquired at 500 and 600 MHz allowed the description of a dynamic model for 36 spins (70% of
the total). Residues that could not be fit by a simultaneous protocol
are located in the extended and/or
disordered regions of the protein. The model free parameters are
plotted in Fig. 3, and a table listing these values is supplied (Table
III).

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Fig. 3.
Plots of optimized Model Free parameters and
their uncertainties as a function of residue number for
RII -(1-44) in the free state (open
triangles) and in the AKAP bound state (filled
triangles and filled circles, corresponding
to protomers A and B, respectively). a, the generalized
order parameter S2; b, the effective
internal correlation time e; c, the exchange
broadening contribution Rex.
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|
 |
DISCUSSION |
Structural Overview--
The protomers in the type II
D/D of
PKA pack together to form an X-type four-helix bundle with an
alternating pattern of (nearly) antiparallel and (nearly)
orthogonal helix-helix interactions around the bundle (Fig.
4) (9, 10, 31). The protein core is
maintained by strong hydrophobic interactions between side chains that
form the dimer interface. In addition, the D/D possesses a
hydrophobic groove along the solvent exposed part of the interface of
helices I, I' (14). The hydrophobic side chains of this groove cluster
against each other and are well defined in the solution structure,
despite being solvent exposed. This is an unusual, but significant
characteristic of the D/D, which promotes participation in
protein-protein interactions with a diverse family of anchoring proteins (32).

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Fig. 4.
Color coding of the observed changes in
backbone dynamics of RII D/D upon
Ht31pep binding. Different views of the dimmer.
a, Connolly surface representation of bound RII -(1-44).
Residues colored in magenta make contact to the AKAP.
b, color coding of changes in order parameter with the same
orientation view as in a. Residues that showed changes in
order parameter ( S2 = S S )
upon binding are colored as red,
S2 < 0, and blue,
S2 > 0. Residues that did not show
significant change upon binding or for which data are not available are
shown in gray. c, ribbon diagram of the
RII -(1-44)·Ht31pep complex. The RII -(1-44)
protomers are colored in orange and yellow,
whereas the AKAP peptide is in red.
d-f are as a, b, and
c, respectively, but with different views.
|
|
Mapping studies on a growing family of anchoring proteins have helped
define a primary sequence of 20 amino acids that exhibit a high
probability of amphipathic helix formation that is the likely site for
D/D binding (33). Ht31pep has emerged as the prototypic
AKAP and structural studies confirmed the role of hydrophobic groups
from the amphipathic helix of Ht31 for high-affinity D/D anchoring
interactions (11). Indeed Ht31pep is a powerful reagent for
the disruption of PKA anchoring inside cells (11). As
Ht31pep displaces a wide variety of AKAP partners in
vivo, it emerges as the ideal system for understanding AKAP
recognition by PKA (14, 32).
Hydrogen Exchange in the Ht31pep·D/D
Complex--
In principle, the changes in the amide-exchange
protection factors of the slowest exchanging protons upon ligand
binding can be used to extract thermodynamic binding constants, given
that the protein is undergoing amide exchange in the EX(2) limit (see "Experimental Procedures") (15, 34). EX(2) is generally the dominating exchange mechanism for backbone amide protons in proteins under conditions where the native state is stable and the intrinsic exchange rate is slow. A pH rate study to test for the possibility of
EX(1) exchange in the D/D was not possible due the fact that the
protein tends to aggregate under pH conditions different from 4.0 (16).
However, mass spectrometric analysis of the exchange upon peptide
binding showed no evidence for EX(1)
exchange.2 Thus, we initially
interpreted our results assuming an EX(2) model.
Unlike the observation of Powell et al. (15) where the
observed changes in amide proton exchange rates are in good agreement with those predicted from the measured ligand binding affinities, we
have found that the calculation of the thermodynamic binding constant
from the observed changes in protection factors greatly underestimates
the Ht31pep binding affinity (Table I and Fig. 1b). Instead, our data suggests that the D/D exchanges
either through local breathing motions (35) or behaves as a highly dynamic conformational ensemble composed of nearly isoenergetic states,
which differ slightly in their exchange properties (36). In the latter
case, binding energy is used for the redistribution of the ensemble
without the necessity of restricting the backbone motions that allow
for H/D exchange (37). Interestingly, members of the calmodulin family
and protein L9, which also bind a diverse family of targets, show
minimal exchange protection upon target binding (38, 39).
The Distribution of Order Parameters--
Whereas the values of
S2 for the entire D/D domain in the
free and Ht31-bound RII
(Fig. 3a) are within error, there
are differences observed between the two species when compared at the
residue level (Fig. 1a). Interestingly, both
increases (10 residues) and decreases (21 residues) in order parameter were observed upon complex
formation (Fig. 4, b and e). Many residues in the
hydrophobic peptide-binding groove (Fig. 4b) showed
decreases (
S2 < 0) in order
parameter, indicating that the residues within this binding
cleft are more flexible in the Ht31pep·D/D
complex. These changes are mapped onto the structure in Fig. 4,
b and e, and include Leu9,
Glu11, Gly15 (only protomer A),
Tyr16, Thr17, Glu19,
Val20 (which demonstrates a very large decrease), and
Arg22. Other residues with decreases in the observed order
parameter are Leu28, Asp30, and
Arg38. Increases in order parameter are observed
(reflecting restricted motions) in residues Ile3
(only protomer B), Gln4 (both located in the extended N
terminus region), Leu12 (only protomer A),
Leu13 (only protomer A), Gln14 (only protomer
B), Leu21 (in the first helix), and Phe36 (in
the second helix).
Internal Motion
e and Exchange Broadening Factor
Rex--
Most residues in helix I and II are well
characterized by the original Lipari-Szabo (25) formalism in which the
internal motions are described by the order parameter and the
effective internal correlation time,
e. Residues that
are disordered in the NMR structures (including the first five residues
in the N terminus and the last six of the C terminus) were better
described with three parameters (S
,
S
, and
e) for slow and fast
internal modes (see Fig. 3b and Table III). Inclusion of an
exchange-broadening factor (Table III and Fig. 3c) to
account for µs-ms motions was necessary for a few residues located in
the first helix (Glu11, Leu13,
Gln14, and Arg22 for the free D/D and
Glu11, Tyr16, and Glu19 for the
complex), and for one residue, Phe36, located in the second
helix in both forms. This region encompasses the hydrophobic binding
groove, and may reflect conformational exchange processes, consistent
with the ensemble view of protein dynamics (40). A similar effect was
also observed in studies on the C-terminal domain of Escherichia
coli topoisomerase I bound to a single-stranded DNA (41).
Conformational Entropy Changes Because of Complex
Formation--
Protein-protein interactions control a diverse set of
biological functions, yet we still do not have a full understanding of
target recognition. Clearly, the molecular basis for protein target
binding is controlled by a variety of factors including favorable
binding enthalpy as well as changes in solvent, side chain, and
backbone entropies of the interacting partners (42, 43). As we observed
unexpected increases in backbone motions as a result of
complex formation, we were interested in estimating the contribution of
these motions to the overall Gibbs free energy of binding. The
energetic benefit associated with increases in backbone flexibility
upon binding can be estimated from the experimental relaxation data,
using the experimentally measured order parameters, S2 (44, 45). This model assumes that the bond
motions of all NH vectors are independent and provides an upper limit
to the true value, as the model is simplified with the assumption of complete independence of motions (44). Nonetheless, the correlation between observed changes in order parameters (and the derived entropy
values) and ligand binding/activity supports the examination of these
parameters (46, 47). The entropic contribution to the free energy of
binding,
G, was determined as described previously (44),
|
(Eq. 6)
|
where S2 is the order parameter,
R is the molar gas constant, and G is the free
energy of Gibbs. Complex formation leads to a small, but favorable,
entropic contribution to the Gibbs free energy of binding
(
G =
H
T
S)
at T = 25 °C of
T
S =
3.7 ± 1 kcal/mol. The total binding free energy change
calculated from the dissociation constant (KD = 16 nM) for the Ht31pep-RII
interaction (16)
yields a value of
10.53 kcal/mol. The corresponding value calculated
(see "Experimental Procedures") from the changes in protection
factors of the core residues upon binding is
G =
0.62 kcal/mol, which is much smaller than calculated from the dissociation constant, and may be explained with the idea of the protein being a highly dynamic collection of states (37).
Adaptive Sites in Protein Target Recognition--
In the majority
of studies on the backbone and/or side chain dynamics of molecular
complexes, decreased motions upon complex formation are observed, as is
expected from an "induced-fit" mechanism (48-50). In these cases,
binding is associated with a loss of conformational entropy that is
necessarily offset by increases in solvent entropy and/or the formation
of favorable enthalpic interactions. Decreases/increases in backbone
dynamics that are compensated for by increases/decreases (respectively)
in distal regions in the backbone, as we report here, have also been
found for protein hydrophobic target interactions (21, 41, 45, 51-55).
It may be that the observed increase in flexibility in the
binding pocket accompanies the release of structured water molecules
from the solvent accessible hydrophobic surfaces. This solvent release
could induce a concurrent disordering of the protein structure (52, 56)
and remains an area of significant interest. Interestingly, in all
cases of increased dynamics upon binding, reported to date, occur in
proteins that recognize diverse targets through hydrophobic
interactions. Thus, a reasonable hypothesis would be that these
proteins use this increased plasticity (either in backbone and/or side
chain domain) to accommodate the differences between target molecules.
Interestingly, structural analysis of various complexes of
bacterial phosphotransferase pathway, involving the protein HPr (57,
58), shows that this protein can interact with proteins of drastically
different folds (e.g. EI, IIAglucose, IIAmannitol) yet uses
the same recognition surface (58). Relaxation studies on Crh, a
structural homolog of Hpr, indicate that it also experiences increased
flexibility upon target binding (59). Whereas side chain plasticity is
obviously important in protein/protein recognition in the HPr system
(58), it would be interesting to determine whether the relaxation
properties for the Hpr complexes described above were consistent with
its structural homolog and was a characteristic of proteins that
interact with many partners. Like the RII
D/D, the consensus binding
site on HPr is an adaptive, highly exposed and energetically important
region that is primed for interaction with diverse molecules (58).
However, in the case of the RII
D/D, although the sequences of AKAPs
are diverse, until recently the structural motifs in recognition
appeared to be conserved (14). Recent evidence indicates that the
centrosomal anchoring protein, pericentrin, appears to present a novel
interaction motif for the D/D (60). Further structural studies of this
anchoring partner will highlight the range of structural motifs capable of participating in anchoring interactions. Clearly, conservation of
hydrophobicity will remain an important recognition mechanism for
tethering PKA through its D/D to diverse anchoring partners.