A multidisciplinary approach to probing enthalpy-entropy compensation and the interfacial mobility model

Article abstract of DOI:10.1021/ja1098287

In recent years, interfacial mobility has gained popularity as a model with which to rationalize both affinity in ligand binding and the often observed phenomenon of enthalpy-entropy compensation. While protein contraction and reduced mobility, as demonst

Full text of DOI:10.1021/ja1098287

ARTICLE
pubs.acs.org/JACS
A Multidisciplinary Approach to Probing EnthalpyꢀEntropy
Compensation and the Interfacial Mobility Model
Erin M. Wilfong,† Yuri Kogiso,‡ Sivaramakrishnan Muthukrishnan,‡ Thomas Kowatz,§ Yu Du,†
Amber Bowie,† James H. Naismith,§ Christopher M. Hadad,‡ Eric J. Toone,† and Terry L. Gustafson*,‡
†Department of Chemistry, Duke University, Durham, North Carolina 27708, United States
‡Department of Chemistry, The Ohio State University, 100 West 18th Avenue, Columbus, Ohio 43210, United States
§Biomedical Sciences Research Complex, The University, St. Andrews, Fife KY16 9ST, Scotland, United Kingdom
S Supporting Information
b
ABSTRACT: In recent years, interfacial mobility has gained popu-
larity as a model with which to rationalize both affinity in ligand
binding and the often observed phenomenon of enthalpyꢀentropy
compensation. While protein contraction and reduced mobility, as
demonstrated by computational and NMR techniques respectively,
have been correlated to entropies of binding for a variety of systems,
to our knowledge, Raman difference spectroscopy has never been
included in these analyses. Here, nonresonance Raman difference spectroscopy, isothermal titration calorimetry, and X-ray
crystallography were utilized to correlate protein contraction, as demonstrated by an increase in protein interior packing and
decreased residual protein movement, with trends of enthalpyꢀentropy compensation. These results are in accord with the
interfacial mobility model and lend additional credence to this view of protein activity.
’ INTRODUCTION
interaction). From the perspective of free energy, as ligand size
increases, diminished favorable enthalpy (due to failure to maxi-
mize interactions) is compensated by a diminished entropic
penalty (less rigidification).
Molecular association lies at the center of virtually all bio-
logical processes. Enzymes bind their substrates to catalyze im-
portant biochemical transformations; ribosomes bind both
tRNAs and mRNA to facilitate protein translation. Despite the
ubiquity of biological binding, the underlying principles govern-
ing these interactions remain obscure. An understanding of
molecular associations is further complicated by several poorly
understood binding phenomena such as additivity, enthalpyꢀ
entropy compensation, and the molecular nature of hydrophobic
dissolution.
Among the most frequently observed and poorly understood
phenomena is enthalpyꢀentropy compensation, or large off-
setting changes in enthalpy and entropy in response to modifica-
tion in protein or ligand structure. In an effort to rationalize
enthalpyꢀentropy compensation in carbonic anhydrase, White-
sides and co-workers suggested that ligand binding might be
driven by tightening of the proteinꢀligand interface, a concept
termed the interfacial mobility.1 The basis of this theory is that
tightening of the ligandꢀprotein interface maximizes enthalpic
intermolecular forces (ionic, van der Waals, and dipoleꢀdipole
interactions), all of which vary inversely as the intermolecular
distance.2ꢀ4 These favorable enthalpic interactions are, in part,
offset by entropic penalties that arise from protein contraction
with resultant rigidification. As ligand size increases, the degree of
tightening of the interface required to maximize enthalpic inter-
actions would require too high an entropic penalty, and enthalpy
falls short of the maximum available. A diminished tightening
produces less rigidification (and a diminished entropic penalty
relative to that which would accompany maximum enthalpic
Because there was no evidence of protein contraction or
ligand-induced protein conformational change for carbonic
anhydrase, Whitesides and co-workers attributed the entropic
penalty to the ligand, that, when small, bound tightly to the
protein interface and, upon elongation, associated more loosely
with the protein binding site.1 However, there is significant evi-
dence that matrix metalloproteinases, particularly matrix metal-
loproteinase-3 (stromelysin-1), do undergo significant confor-
mational change upon binding.5ꢀ7 Such conformational changes
raise the possibility that the protein itself could contract and
rigidify around the ligand thereby tightening the proteinꢀligand
interface. The thermodynamics would be the same as described
by Whitesides and co-workers in the interfacial mobility model.1
While compelling in theory, experimental evidence for ligand-
induced protein contraction as an underlying basis for enthalpyꢀ
entropy compensation is lacking. Calorimetric studies of the
interaction of stromelysin-1 (matrix metalloproteinase 3) with
the CGS 270238,9 series of ligands revealed significant enthalpyꢀ
entropy compensation as a function of ligand complexity
(Figure 1). As with most instances of enthalpyꢀentropy com-
pensation, the significance and origin of this observation were
not readily apparent. A challenge in studying complex biological
phenomena is that any single biochemical technique provides
Received: November 1, 2010
Published: June 21, 2011
r
2011 American Chemical Society
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ARTICLE
’ EXPERIMENTAL SECTION
Raman Microscopy. Raman spectra were acquired using a Raman
microscope (Horiba JY, HR 800) operated by LabSpec5. The 632.8 nm
output from a HeꢀNe laser (20 mW) is passed through an interference
filter to eliminate the plasma lines of the laser. Subsequently, the output
is reflected by a notch filter and directed toward the sample. The sample
position and the laser focal point are adjusted by viewing a real-time
video. The laser power at the sample is 6 mW, and the laser spot size is
2 μm. Raman signal is collected at 180° backscattering geometry by a
50ꢁ objective lens (NA, 0.75), passed through a notch filter to reject the
Rayleigh line and directed through a 200 μm confocal hole. The signal
passes through a 200 μm slit of the spectrograph (800 mm focal length)
and is then then dispersed by an 1800 grooves/mm grating and detected
by a CCD. The grating is calibrated by setting it to 0 nm using zeroth
order white light and then to 520.7 cmꢀ1 using the silicon band. The
spectral resolution and wavenumber position repeatability are reported
as 0.3 cmꢀ1 at 680 nm and 1 pixel, which corresponds to 0.5 cmꢀ1
respectively.
,
Expression and Purification of Stromelysin-1. Stromelysin-1
catalytic domain (SCD) was expressed and purified as previously
described.20 Briefly, the nucleic acid sequence encoding amino acid
residues 83ꢀ256 was cloned into the pET28b vector (Novagen); a stop
codon was introduced in the reverse primer. The construct was
expressed as inclusion bodies in E. coli BL-21(DE3)Gold cells and was
purified using metal affinity chromatography. The purified protein was
then dialyzed into the appropriate buffer (50 mM TrisHCl, pH 7.5,
10 mM CaCl2, 1 μM Zn(OAc)2).
Figure 1. Structures and thermodynamics of ligand binding for CGS
27023 series ligands.
only a partial picture of the studied event. Although calorimetry
provides accurate measures of Keq and enthalpy, which can then
be used to calculate free energy of binding and entropy, thermo-
dynamic studies provide no structural insights. Crystallography
provides a static view of molecular associations, but lacks insight
into dynamic processes, that is, fast side chain motions, residual
protein motion, periodic secondary structure changes, and global
changes in physical properties upon binding. While dynamic
NMR studies can provide insight into the former,10ꢀ13 Raman
spectroscopy can provide insight into global changes in the
protein structure and microenvironment. As such, Raman spec-
troscopy represents a valuable, but underutilized, tool with which
to probe key facets of protein ligand association.
Raman spectroscopy reports on vibrational modes of a mole-
cule, and each vibration has a unique, characteristic signature.
Thus, Raman spectroscopy can provide unique insights into
changes in protein structure and, crucially for consideration of
the interfacial mobility model, contraction. Unfortunately, Raman
spectra are exceedingly complex, as each bond contributes to
several vibrational modes. The challenge then becomes one of
simplifying spectra such that useful information can be extracted;
this simplification is most frequently accomplished by focusing
on only a portion of the spectrum. Alternatively in Raman dif-
ference spectroscopy, the spectra of unbound protein and buffer
are subtracted from that of the bound complex.14ꢀ16 While the
difference spectrum is dominated by ligand bands, protein bands
are also observed, and these bands correspond to vibrational
modes of the protein that are altered by ligand binding.17,18
These Raman signatures yield insight into changes in secondary
structure, side chain hydrogen bonding, side chain conformation,
and, in the case of tryptophan, side chain environment.14,19 Here,
we utilize Raman difference spectroscopy in conjunction with
both crystallographic and thermodynamic data to assess the
validity of the interfacial mobility model in the case of strome-
lysin-1 ligand binding.
Ligand Synthesis. All ligands utilized in these investigations were
previously described using known synthetic techniques.8,9,21 Full details
can be found in the Supporting Information.
X-ray Crystallography. SCD was purified as described above,
dialyzed into 2 mM TrisHCl, pH 7.5, 10 mM CaCl2, and 1 μM Zn
(OAc)2, and lyophilized. After resuspension, a final purification step on a
HiPrep 16/60 Sephacryl S-200 size exclusion column (Amersham
Biosciences) was carried out. About 14 mg of the lyophilized enzyme
powder was dissolved in 5 mL of buffer containing 2 mM TrisHCl,
pH 7.5, 10 mM CaCl2, and 1 μM Zn (OAc)2 and loaded onto the
column. SCD fractions eluted in 2 mM TrisHCl, pH 7.5, 10 mM CaCl2,
and 1 μM Zn (OAc)2 were then checked for their purity by SDS-
PAGE. Fractions containing pure SCD were pooled and concentrated
(VIVASPIN 20, 3000 MWCO PES, Sartorius Stedim Biotech GmbH
(SSB)) to 11.5 mg/mL. Prior to crystallization, SCD at a concentration
of 11.5 mg/mL was centrifuged for 5 min at 4 °C. Either inhibitor 3 or 4
(dissolved in 100% dimethyl sulfoxide (DMSO) (Sigma)) was added
to the protein solution to a final inhibitor concentration of 4 mM. The
two samples were then incubated for 1ꢀ2 h at room temperature and
centrifuged again. Crystallization experiments employed sitting-drop vapor
diffusion at room temperature. Protein and mother liquor were mixed in a
ratio of 1:1 (2.5 μL + 2.5 μL, reservoir volume 100 μL). Sealed plates were
then incubated at 20 °C. Crystals large enough for diffraction experiments
appeared after 1ꢀ4 weeks under the previously published conditions.14
Before data collection, crystals were cryoprotected in a solution
containing 0.15 M ammonium sulfate, 0.1 M Na-cacodylate, pH 6.5,
30% PEG 8K, 4 mM inhibitor, and 10% PEG 400. X-ray data sets of SCD
cocrystals, one in complex with inhibitor 3 and a second one complexed
with compound 4, were collected on the in-house RA-Micro 7 HFM
Table Top Rotating Anode X-ray Generator (Rigaku) to 2.4 and 2.5 Å,
respectively. The diffraction data were processed with HKL2000.22 The
catalytic domain of MMP-3 (pdb accession code 1b8y)23 was used as a
model for Molecular Replacement using PHASER.24,25 The structures
were refined with REFMAC526 with manual intervention with COOT27
and validated using MOLPROBITY.28 Dictionaries for the compounds
were created by PRODRG.15
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ARTICLE
Protein Superimposition. The PDB files for SCD bound to nine
ligands (1BM6, 2JT5, 1BQO, 2JNP, 1B3D, 2USN, 1B8Y), including
SCD bound to 3 and 4, were superimposed to apo SCD (1cqr) using
Superpose29 to minimize the global rmsd of C(R) atoms. The super-
impositions were compared in PyMol,30 and the distance between
corresponding C(R) was measured for each bound structure relative
to apo SCD. These values were entered into Microsoft Excel for
additional data processing and graphical representations.
Table 1. Thermodynamics of Binding for Compounds 1ꢀ4a
ΔG
ΔHbind
TΔSbind
ΔCP
1
2
3
4
ꢀ11.1 ( 0.1
ꢀ10.2 ( 0.1
ꢀ10.6 ( 0.2
ꢀ8.9 ( 0.1
ꢀ9.0 ( 0.3
ꢀ7.5 ( 0.4
ꢀ10.7 ( 0.4
ꢀ13.6 ( 0.3
2.1 ( 0.3
2.7 ( 0.4
ꢀ73 ( 7
ꢀ74 ( 2
ꢀ69 ( 10
ꢀ60 ( 12
ꢀ0.1 ( 0.4
ꢀ4.7 ( 0.3
a ΔG, ΔH, and TΔS are described in kcal molꢀ1. ΔCP is reported as cal
Raman Spectroscopy Samples. Compounds 1ꢀ4 were pre-
pared as previously described.8,9,21 12.5 mM stocks of compounds 1, 2,
3, and 4 were made in MeOH. The stock concentration was validated by
titration against previously standardized protein. Protein was expressed
and purified as described above and concentrated to 200ꢀ250 μM;
concentrations were confirmed by the method of Edelhoch,31 using
ε280 = 27 630 Mꢀ1 cmꢀ1. Ligand was added in a 1:1 protein to ligand
ratio, and additional MeOH was added to a final concentration of 2%.
Samples were prepared within 24 h prior to obtaining Raman spectra.
Drop Coating Deposition Raman (DCDR). All experiments
were performed at room temperature. For protein samples, 4 μL of a
protein solution was deposited on a SpectRIM substrate, and water was
evaporated without further treatment. The resulting deposit had a
diameter of less than 2 mm. Raman spectra of protein samples were
obtained by focusing the laser on the protein ring, which was approxi-
mately 50 μm away from the outer edge of the ring. The exposure time
was 180 s for each spectrum, and six spectra were averaged for each
sample. A buffer sample was prepared in the same manner. The buffer
spectrum was obtained from the film-like portion of the deposit rather
than the crystals formed around the center of the sample to achieve
higher S/N. The exposure time was 10, and five spectra were averaged.
Solid ligands were prepared by depositing 3 μL of methanol stock
solutions on the SpectRIM substrate and evaporating the solvent. The
exposure time was 15, and five spectra were averaged for each ligand.
Collection of Stromelysin-1 Complex Spectra. A difference
spectrum that contains information of a bound ligand and changes in the
protein conformation was obtained by subtraction of an apo protein
spectrum from a spectrum of the complex (protein + ligand). The spec-
trum of the buffer was also subtracted or added because the contribution
of buffer signals in each spectrum varies slightly (e.g., [difference] =
[complex] ꢀ [apo protein]*f1 ꢀ [buffer]*f2, where f1 and f2 are scaling
factors). Scaling factors for the apo protein and buffer spectra were
chosen to achieve flat baselines. Positive bands can result from bound
ligands, newly formed bonds, and changes in protein upon complexa-
tion. Negative bands can be due to loss or decrease in band intensity of
existing protein bands upon ligand binding.
molꢀ1 Kꢀ1
.
Because stromelysin-1 undergoes protonation of H224 upon
binding,5 it is critical to ensure that analyses are conducted with
the actual thermodynamics of binding. In all cases, the calculated
n was between 0.3 and 0.35 protons molꢀ1, which is within the
error of measurement. The derived thermodynamic parameters
for binding are shown in Table 1.
Previously, enthalpyꢀentropy compensation has been ratio-
nalized by changes in protein or host hydration upon bind-
ing.33,34 Fortunately, hydrophobic hydration affects ΔCP in a
predictable fashion. While changes in electrostatics or vibrational
states of a protein during binding could, in principle, impact
ΔCP,35 ΔCP is the best available measurement of changes in
solvent-exposed hydrophobic suface area and is therefore con-
sidered a hallmark of the hydrophobic effect.36ꢀ38 The values of
ΔCP observed here, ranging from ꢀ60 to ꢀ74 cal molꢀ1 Kꢀ1
,
show no evidence of changes in protein solvation upon binding.
Additionally, there is no evidence that the presence of polar
ligand functional groups accounts for the observed enthalpyꢀ
entropy compensation. Clearly, transfer of water from the
protein to the ligand upon binding would contribute to the
observed ΔCP. However, if the transfer of water molecules from
the protein surface to polar ligand functionalities contributed
significantly to enthalpyꢀentropy compensation, such trends
should correlate the presence or absence of a polar moiety.
Compounds 1 and 3 contain a pyridyl group absent in com-
pounds 2 and 4. Nonetheless, compounds 1/2 and 3/4 have
similar heat capacities and thermodynamic parameters. We
therefore reject changes in protein solvation upon binding as
an explanation for the observed thermodynamic parameters and
will investigate a possible structural basis for enthalpyꢀentropy
compensation in this system.
Determination of ProteinꢀLigand Crystal Structures.
Both SCD complex structures for compounds 3 and 4 crystal-
lized in the same space group with identical cell dimensions (see
the Supporting Information). Unsurprisingly, both structures
of SCD share the same fold as other previously reported
SCD.5,23,39ꢀ46 Superposition of the SCD structures in complex
with inhibitors 3 and 4 gives an rmsd of 0.130 Å for 129 C(R)
positions (Figure 2). Difference electron density for the inhibi-
tors was visible in the first unbiased maps. The asymmetric unit of
both structures is composed of one monomer, both of which are
essentially complete apart from portions of the “long, flexible
loop” (amino acids 210ꢀ234).5 In the SCD structure complexed
with inhibitor 3, the electron density for amino acids 126, 213,
214, 218, 224, 231, 232, 237, 241, and 248 is weak; amino acids
215, 216, and 225ꢀ230 are absent. In the structure bound with
compound 4, the electron density for amino acids 213, 224, 231,
233, 236, 243, and 246ꢀ248 is poorly defined, while residues
215, 216, and 225ꢀ231 are completely disordered. Disordered
amino acids within this flexible loop have been previously
reported.5,23,42 The positions and interactions of the catalytic
Difference spectra between spectra of complexes were obtained
to determine the dependence of protein conformation changes on
ligand structure (e.g., [difference] = [complex 1] ꢀ [complex 2]*f1 ꢀ
[buffer]*f2), where f1 and f2 are scaling factors. The difference spectra
were smoothed using a smoothing function in the Igor software pack-
age (Wavemetrics). A binomial algorithm and smoothing factor of 5
were used.
’ RESULTS AND DISCUSSION
Thermodynamics of Ligand Binding. While all ligands in the
CGS 27023 series have been previously described,8 the thermo-
dynamics of binding were unknown. The free energy, enthalpy,
entropy, and heat capacity of binding were determined by
isothermal titration calorimetry. Heat capacity experiments were
conducted at 15, 25, and 37 °C. Ligand binding was characterized
in three buffers, MOPS, HEPES, and TRIS, to account for proton
transfer events.32 Observed enthalpies of binding were plotted
against enthalpies of ionization. Linear regression was then used
to determine the number of protons, n, transferred upon binding.
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Figure 2. Stereo view of the C(R) tracing of SCDs complexed with
inhibitors 3 (cyan) and 4 (magenta). This figure has been produced
using PyMol.30
Figure 3. Stacked difference spectra for compounds 1ꢀ4. Bands
assigned to protein are labeled with the appropriate Raman shift.
Unlabeled bands correspond to ligand shifts (unpublished results).
and structural Zn2+ and three Ca2+ ions are essentially identical in
both complexes and consistent with previous reports.5,23,39ꢀ46
One sulfate ion is also present in the SCD structures complexed
with compounds 3 and 4 at the same position observed for SCD
complexed with other nonpeptide inhibitors.23
complex spectra are compared (Supporting Information). Addi-
tional experiments would be required to assign these bands
unequivocally.
Collection of Difference Spectra. Solution-phase Raman
spectroscopy was not feasible because protein fluorescence
following excitation at 568 nm obscured the Raman spectrum.
Fortunately, drop coating deposition Raman (DCDR) generated
useful spectra. Difference spectra containing information on
protein vibrational changes upon binding were generated by
([complex] ꢀ [apo protein]*f). The scaling factor, f, is the
multiplication factor required to match the intensity of the
1010 cmꢀ1 phenylalanine band in the two subtracting spectra.
A total of five spectra were collected for all bound complexes and
unbound stromelysin-1 and averaged to yield the final spectra.
To avoid complicating the spectra with unbound ligand, a 1:1
ratio of protein:ligand was used. On the basis of the previously
determined KD, this ratio resulted in a protein saturation greater
than 99%. DCDR was also used to collect both pure ligand and
buffer spectra to ensure only protein bands were included in the
subsequent analysis (see the Supporting Information). The
details of ligand band assignment will be discussed in a future
publication. Table 2 summarizes the major protein peaks ob-
served in all Raman spectra.
Ligand Binding Induces Changes in Secondary Structure.
Raman spectroscopy is exquisitely sensitive to changes in protein
secondary conformation. The main-chain backbone has 12 normal
vibrational modes of which the amide I (carbonyl stretch) region is
most commonly used to analyze secondary structure changes.19,47
A positive band seen at 1660 cmꢀ1 appears in all difference spectra
for all protein complexes; a second positive feature at 1688 cmꢀ1 is
present in the difference spectra of proteinꢀligand complexes 1, 2,
and 4. While less obvious, the same feature near 1688 cmꢀ1 is also
seen in the difference spectrum of 3. Both bands are assigned to
correspond to random coil, which has multiple peaks across the
entire amide I region. This conclusion is supported by crystal-
lographic data, which show a universal decrease in R-helix and
β-sheet structure upon binding and a compensatory increase in
random coil features (Table 3).
Ligand Binding Changes the Local Environment of Tryp-
tophan. Tryptophan is one amino acid that is particularly
conducive to study by Raman spectroscopy as specific bands
exist that characterize hydrogen bonding, side chain orienta-
tion, and hydrophobic environment.19 In resonance Raman
spectroscopy, UV excitation amplifies the tryptophan spectrum
making them even more apparent. The ratio of the W7 fermi
doublet of tryptophan (1340/1360 cmꢀ1) has been repeatedly
validated as a marker of hydrophobicity in the tryptophan local
Newly Formed Bands. Three positive bands at 1386, 1072,
and 894 cmꢀ1 appear in all difference spectra between the apo
protein and bound complexes (Figure 3). These bands do not
correspond to any ligand modes or to bands observed in the apo
protein spectrum. Further, these bands disappear when two
Table 2. Observed Difference Spectra Raman Shifts Assigned to Protein19
compound 1
compound 2
compound 3
compound 4
amide I, random coil
tryptophan (W3)
unassigned protein
tryptophan or phenylalanine
unassigned protein
tryptophan
1689/1659
1557
1688/1660
1557
1660
1558/1550 (d)
1386 (br)
1207
1688/1660
1556
1386 (br)
1206
1385 (br)
1205
1386 (br)
1208
1071
1072
1072
1072
1013
1013
1013
1013
phenylalanine
1003
1004
1003
1004
unassigned protein
tyrosine
894
894
894
893
829
829
828
828
tryptophan (W18)
763
762
765
763
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Table 3. Changes in Secondary Structure for Compounds 1,
3, and 4 As Evidenced by X-ray Crystallography
R-helix residues
β-sheet residues
random coil residues
gained
compound
lost
lost
1
3
4
ꢀ4
ꢀ6
ꢀ7
ꢀ2
ꢀ2
ꢀ2
+6
+8
+9
environment.48ꢀ50 Unfortunately, attempts at solution-phase
Raman spectroscopy were limited by sample fluorescence, and the
Raman microscope available for DCDR did not have the option of
UV excitation. Thus, the studies described here were limited to
nonresonance Raman spectroscopy. In their initial report on the
W7 band, Miuri and co-workers note that the ratio of the W7
doublet cannot be used with visible light excitation as CH ali-
phatic stretches overlap the 1340 cmꢀ1 band.50 Additionally, the
1360 cmꢀ1 is not always discernible in nonresonance spectra,51
even if readily apparent in the resonance Raman spectra.48,49
One structurally sensitive vibrational band in the nonreso-
nance Raman spectroscopy of proteins is the W18 stretch
(760 cmꢀ1), corresponding to the indole ring of tryptophan.
Because the W18 stretch is among the most intense in the Raman
spectrum, changes in intensity are readily detected, and the
intensity of this band has been previously correlated to the
number of hydrophobic contacts made by the indole ring. This
deduction is based on work using the model compound,
3-methylindole. By measuring the W18 band intensity of
3-methylindole in solvents including H2O, 1:1 H2O:EtOH,
EtOH, and pentane, Miura et al. demonstrated that the band
intensity varied inversely with solvent hydrophobicity.52 Because
band intensity was comperable in vapor and aqueous spectra,
Miura et al. concluded that an increased number of hydrophobic
contacts on the indole ring resulted in a relatively decreased
band intensity.52 When considering the W18 band in difference
Raman spectra where the spectrum of unbound protein is sub-
tracted from that of bound protein, increased hydrophobic con-
tacts on tryptophan in the bound state lead to the W18 band ap-
pearing as a negative feature. Figure 3 shows negative features in
the difference spectra at ∼760 cmꢀ1 corresponding to the W18
stretch, indicating the presence of an increased number of
hydrophobic contacts in the bound state.
There is a significant temptation to analyze the relative
intensities of the various W18 ligand bands through the genera-
tion of a double difference spectrum. One must, however,
exercise caution as any error in the initial Raman spectra has
been significantly propagated as two difference spectra were
generated. Assuming a 5% error in the initial measurements
(inclusive of buffer subtraction), propogation of errors indicates
that the error in the difference spectra is 7%. Generation of a
double difference spectra would further increase the error to 10%.
To determine if any reliable trends in relative W18 band inten-
sities were present, a series of double displacement spectra were
generated (Supporting Information Figures S6 and S7). Unfortu-
nately, these spectra yielded unreliable and, at times, contradictory
results, likely due to the small variations in peak intensity and the
substantial error associated with generating double difference
spectra. To ascertain trends among ligands, nondifference Raman
spectroscopy will have to be combined with other high resolution
techniques, such as isothermal titration calorimtery X-ray crystal-
lography or NMR protein structure determinations.
Figure 4. Conformation of the three internal tryptophans of unbound
SCD (pink), SCD bound to 1 (blue), and SCD bound to 4 (purple).
Table 4. Conformations of Stromelysin-1 Catalytic Domainsa
W92
C1
W124
C1
W186
C1
C(R)
X1,2 C(R)
X1,2 C(R)
X1,2
1
3
4
1.58 Å 1.18 Å 17.3° 1.17 Å 1.1 Å 1.1° 1.22 Å 1.27 Å 2.3°
0.20 Å 0.20 Å 6.1° 0.21 Å 0.19 Å 2.7° 0.08 Å 0.15 Å 0.1°
0.28 Å 0.28 Å 8.7° 0.18 Å 0.18 Å 4.5° 0.22 Å 0.22 Å 0.3°
a Distances for C(R) and C(1) are the measured distance between the
unbound crystal structure 1CQR with the superimposed bound com-
plex. RMS of C(R) for superimposition of apo with compounds 3 and 4
was 0.63; RMS of C(R) for superimposition of apo with compound 1
was 3.75. Value for χ1,2 dihedral is the change in the dihedral upon ligand
binding.
Despite this limitation, nondifference Raman spectroscopy does
provide clear evidence that the number of hydrophobic contacts
within the protein interior of stromelysin-1 increases upon ligand
binding. Additionally, this technique is reasonably accessible and
does not require protein crystals, which can be difficult to obtain.
There are two ways in which interior tryptophans could experi-
ence increased hydrophobic contacts, residue translocation or
protein contraction with increased packing of the protein
interior.
We first consider the possibility of tryptophan translocation.
X-ray crystallographic data show no major change of tryptophan
conformation upon ligand binding (Figure 4). For tryptophan
translocation to account for the observed W18 band, the
translocation must differ significantly between compounds 1
and 4 because the W18 band is much stronger for both com-
pounds 3 and 4 than for compound 1. Table 4 shows the change
in position of C(R) and C1 and the χ1,2 from apo protein. While
the χ1,2 angle for W92 does change significantly upon binding,
the change is consistent across the series. Also, compound 1
exhibits the largest change in W92 χ1,2 dihedral angle but the
smallest change in W18 intensity upon ligand binding. These
small changes in χ1,2 dihedral are also evidenced by the W3
tryptophan stretch in all difference spectra. Thus, the structural
data show no evidence for translocation.
Protein contraction would also increase the number of
hydrophobic contacts on tryptophan. Because proteins contain
intraprotein voids53ꢀ57 and have average interior surface com-
plementarities of only 60ꢀ80%,58,59 contraction is feasible. An
analysis of 50 protein structures demonstrated that most buried
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tryptophans only have ∼60% surface complementarity (surface
area contacted by another amino acid residue); 40% of the
tryptophan surface lacks any hydrophobic contacts.58,59 Lyso-
zyme is a protein for which ligand binding and protein contrac-
tion were convincingly linked using both volume and intrinsic
compressibility measurements.60 Ligand binding by lysozyme
also decreases the W18 mode intensity and led to negative
features at ∼760 cmꢀ1 52
,
which could be contributed to by
Figure 5. Average variation of C(R) upon ligand binding. Based on
protein superimposition of PDB files 1BM6, 2JT5, 1BQO, 2JNP, 1B3D,
2USN, 1B8Y, and the crystal structures described herein. Solid line
depicts ligands binding only the S10 hydrophobic pocket and catalytic
zinc. Dashed line depicts larger ligands binding additional subpockets.
protein contraction. While direct correlation of the W18 bandwith
protein contraction has not, to our knowledge, been reported in
the literature, supporting data have, in fact, been published by several
groups independently. In his 2004 review of protein contraction/
expansion as a driving force for binding and enthlapyꢀentropy com-
pensation, Dudley Williams and co-workers described two model
systems, avidinꢀbiotin and hemoglobinꢀoxygen.61 Avidinꢀbiotin
binding is a representative system of protein contraction. Using H/D
exchange of amide backbone proteins characterized by MALDI,
binding of biotin by avadin led to a marked decrease of H/D
exchange of the protein backbone, which was interpreted as
increased protein packing and therefore decreased access of D2O
to the amide protons.62 Thermodynamically, binding of biotin
and avadin demonstrates a marked enthalpic benefit and entropic
penalty of binding.63,64 This system has also been studied by
difference resonance Raman spectroscopy to analyze changes in
the local tryptophan environments upon binding. Both the W7
and the W18 bands experienced an increase in intensity indica-
tive of an increase in local hydrophobicity about the indole ring.65
In conglomeration, these results support the use of the W18 band
as a marker for protein contraction.
Consideration of the Interfacial Mobility Model. The
interfacial mobility model postulates that high affinity ligand
binding results from tightening of the proteinꢀligand interface,
which in the case of stromelysin-1, we propose arises from
protein contraction about the ligand. Consequences of protein
contraction would include increased internal protein packing,
rigidification, and decreased residual movement. Thermodyna-
mically, the interfacial mobility model manifests with an enthal-
pic benefit and entropic penalty of binding that is inversely
proportional to ligand complexity. While it is currently impos-
sible to deconvolute accurately and reliably the contribution of
van der Waals interactions and hydrogen bonding to binding
enthalpy, there has been success in correlating entropies of
binding to either residual protein motion or residual entropy.70,71
As both decreased residual protein movement and increased
internal packing are consequences of protein contraction, we
correlate our Raman observations of internal packing (Figure 3),
and therefore contraction and decreased residual protein motion,
with the experimental entropies of binding (Figure 1).
In the case of compounds 1 and 2, both enthalpy and entropy
contribute to ligand binding, and the magnitude of entropic
contribution is comparable (2.1 and 2.7 kcal molꢀ1, respec-
tively). Entropy does not contribute significantly for binding of
compound 3 (ꢀ0.1 kcal molꢀ1), but a significant entropic
penalty of binding occurs for compound 4 (ꢀ4.7 kcal molꢀ1).
On the basis of these thermodynamic observations, the interfacial
mobility model predicts that compounds 1 and 2 would experi-
ence relatively small and comparable contractions as they have
comparable entropies of binding. Binding of compound 3 should
cause a protein contraction greater than that for compounds 1
and 2 but less than for compound 4.
While internal packing, as demonstrated by the W18 trypto-
phan band, is one proxy for protein contraction, a second proxy
marker for this phenomenon is decreased residual movement.
Additionally, X-ray crystallography is high resolution and likely
to yield insights into minor differences between ligands. To this
end, an analysis of crystal structures was undertaken.5,6,23,39,45,72
Ten solved bound structures, including those for compounds 1,
3, and 4, were superimposed with apo SCD using SuperPose,29
and the difference between C(R) positions for apo- and holo-
protein was measured for all residues. These results are plotted
graphically in Figure 5. The average difference in C(R) position
across all residues and structures is approximately 1 Å. There are,
however, certain protein regions where the difference C(R) is
much greater, and these regions correspond to random coil
secondary structure. All structures demonstrate increased variability
of C(R) position for the loop encompassing residues 221ꢀ233.
Similarly, when investigating protein expansion as demon-
strated by hemoglobin binding to oxygen, the converse finding is
obtained.61 Hb exists in two forms in the blood, a “tense” rigid
state having a low affinity for O2 and a high affinity “relaxed” state
that avidly binds both O2.66 Using H/D exchange and ESI
experiments, Williams and co-workers demonstrated that tran-
sitioning from a tense to relaxed state upon binding O2 resulted
in increased H/D exchange and protein expansion.62 Subsequent
binding events lead to relaxation of additional subunits until the
protein is fully relaxed after binding of the fourth O2 molecule.
Thermodynamically, this phenomena was entropically driven
and entropically opposed, with each successive binding event re-
sulting in a smaller entropic benefit and enthalpic penalty of bind-
ing.67,68 The concept of protein expansion is also supported spe-
ctroscopically by analyzing work by Nagatomo et al.,69 which
showed decreased hydrophobicity of tryptophans using the W18
band upon carbon monoxide binding. Because this was a non-
resonance study, the tryptophan spectra are not enhanced, and
the 1360 cmꢀ1 peak is not readily visible for analysis. Carbon
monoxide is more facile to use for spectroscopic experiments as
its higher affinity results in a more stable complex. Nonetheless,
the binding site and binding mode are conserved, and protein
dynamics should be conserved in both cases.
Ideally, our Raman spectroscopy work would be supplemen-
ted by physical measurements of intrinsic compressibility, but
these require crystals that we were unable to acquire. None-
theless, either tryptophan translocation or protein contraction
must occur to produce the W18 observations in these studies,
and data supporting translocation were lacking. This lack of
evidence for translocation combined with the previous correla-
tion of decreased W18 band and protein volume for lysozyme
leads us to the conclusion that protein contraction is responsible
for the increased hydrophobic contacts on tryptophan.
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Interestingly, two loops (150ꢀ160 and 170ꢀ175) demonstrate an
increased difference of C(R) position variability for large ligands, as
defined by binding more than just the S10 and Zn2+ subpockets,
versus small ligands that bind only the S10 and Zn2+ subpockets. The
average difference in C(R) for large as compared to small ligands is
0.9 ( 0.2 versus 2.0 ( 0.5 Å for residues 148ꢀ161, respectively.
Similarly, the average difference in C(R) for residues 169ꢀ175 is
1.1 ( 0.2 versus 1.8 ( 0.3 Å, respectively, for large and small ligands.
It is intriguing that these regions in relatively close proximity to the
active site (Figure 6) demonstrate increased variability in C(R) large
relative to small ligands as this could reflect decreased residual
movement of these residues. If C(R) variability between super-
imposed protein structures truly reflects residual protein motion,
this would further support the interfacial mobility model.
We have presented data that protein contraction, as judged by
the W18 Raman band, varies inversely with ligand complexity.
Additionally, crystallographic analysis reveals reduced motions in
two loops and possibly an R-helix near the active site upon
binding of high affinity small ligands. The W18 Raman band
indicates that for compounds 1 and 2, the protein contracts
identically even though 2 lacks the pyridyl substituent. Similarly,
compounds 3 and 4 behave similarly despite the difference of
pyridyl moiety. Figure 7 shows the subpockets of the SCD active
site. The majority of subpockets are composed of multiple
secondary features. The S10 hydrophobic pocket is made up of
a loop (residues 218ꢀ222) and an R helix (residues 195ꢀ201).
The S1 groove is formed by a loop (residue 163) and β sheet
(residue 165ꢀ166); that same loop also forms the P1 groove
(residues 162, 164). Thus, while lacking the pyridyl moiety ring,
compound 2 still contacts the P1 loop through binding of its
isopropyl group. Although compound 3 contains the pyridine
moiety, the structure reveals that this substituent is rotated away
from the P1 groove. Thus, compounds 1 and 2 contact the same
secondary features; the same observation is true for compounds
Figure 6. Regions of variable mobility for SCD upon ligand binding.
Regions depicted in red show a clear increase in mobility when SCD is
bound to large ligands. Regions depicted in orange show a trend toward
increased mobility when bound to large ligands.
Figure 7. Meshwork structures of SCD subpockets. (A) Empty meshwork of SCD. S10 pocket is in blue, P1 groove is in red, and S1 groove is in purple.
(B) Meshwork of bound compound 1. (C) Meshwork of bound compound 3. (D) Meshwork of bound 4.
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3 and 4. The p-methoxyphenyl group for compounds 3 and 4
interacts more extensively with the protein than seen in com-
pound 1. This could result in tightening of the S10ꢀligand
interface around the smaller ligands.
proteomics. E.J.T. acknowledges financial support for his re-
search efforts from the NIH (1R01GM57179).
When Krishnamurthy et al. first described the interfacial
mobility model, they defined ligand size as the number of “distal
residues” contacted.1 In the context of human carbonic anhy-
drase, the distal residues were amino acids at an increased
distance from the catalytic zinc in the conical active site. Because
SCD has an active site groove, we originally thought to define
ligand size as a function of subpockets occupied. This assumption
is flawed because small changes in ligand structure can drama-
tically alter the binding mode; that is, the absence of the isopropyl
moiety allowed reorientation of the pyridyl moiety in the active
site. While a priori we had expected to see a linear trend in protein
contraction, this ligand rearrangement in the active site caused
compound pairs 1/2 and 3/4 to give similar results despite
possessing moieties that could have bound additional subpock-
ets. The small ligands 3/4 bind loop 218ꢀ222 and R-helix
195ꢀ201 in addition to the catalytic zinc. The large ligands 1/2
interact with these residues as well as loop 162ꢀ164 and β-sheet
165ꢀ166. Once “ligand size” was defined by the secondary
structural elements involved in ligand binding, which correlates
closely with residues contacted, the degree of protein contraction
correlates well with “ligand size”. Thus, trends in protein con-
traction and thermodynamics are in good accord with those
predicted by the interfacial mobility model.1
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