Spin-labeled derivatives of cardiotonic steroids as tools for characterization of the extracellular entrance to the binding site on Na+,K+-ATPase

Article abstract of DOI:10.1111/febs.14480

The information obtained from crystallized complexes of the Na⁺,K⁺-ATPase with cardiotonic steroids (CTS) is not sufficient to explain differences in the inhibitory properties of CTS such as stereoselectivity of CTS binding or effect of glycosylation on the preference to enzyme isoforms. The uncertainty is related to the spatial organization of the hydrophilic cavity at the entrance of the CTS-binding site. Therefore, there is a need to supplement the crystallographic description with data obtained in aqueous solution, where molecules have significant degree of flexibility. This work addresses the applicability of the electron paramagnetic resonance (EPR) method for the purpose. We have designed and synthesized spin-labeled compounds based on the cinobufagin steroid core. The length of the spacer arms between the steroid core and the nitroxide group determines the position of the reporting group (N-O) confined to the binding site. High affinity to Na⁺,K⁺-ATPase is inferred from their ability to inhibit enzymatic activity. The differences between the EPR spectra in the absence and presence of high ouabain concentrations identify the signature peaks originating from the fraction of the spin labels bound within the ouabain site. The degree of perturbations of the EPR spectra depends on the length of the spacer arm. Docking of the compounds into the CTS site suggests which elements of the protein structure might be responsible for interference with the spin label (e.g., steric clashes or immobilization). Thus, the method is suitable for gathering information on the cavity leading to the CTS-binding site in Na⁺,K⁺-ATPase in all conformations with high affinity to CTS.

Full text of DOI:10.1111/febs.14480

Spin-labelled derivatives of cardiotonic steroids as tools for
characterization of the extracellular entrance to the Na+,K+-ATPase
binding site
Jin-Hua Guo1, Ren-Wang Jiang1, Jacob Lauwring Andersen2, Mikael Esmann2,
Natalya U. Fedosova2
1College of Pharmacy, Jinan University, Guangzhou City, Guangdong Province,
2
510632, P. R. China, Department of Biomedicine, Aarhus University, DK-8000,
Aarhus, Denmark
Corresponding authors:
Ren-Wang Jiang:
College of Pharmacy, Jinan University, Guangzhou City,
Guangdong Province, 510632, P. R. China;
rwjiang2008@126.com, +8620 85221016
Natalya U. Fedosova: Department of Biomedicine, Aarhus University, DK-8000,
Aarhus, Denmark;
nf@biomed.au.dk, +45 87167752
Article type : Regular Paper
Running title: Application of EPR for spatial description of the digitalis binding site
Keywords: Na-pump, EPR, structure, membrane protein, protein-ligand interactions
This article has been accepted for publication and undergone full peer review but has not
been through the copyediting, typesetting, pagination and proofreading process, which may
lead to differences between this version and the Version of Record. Please cite this article as
doi: 10.1111/febs.14480
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Abbreviations, Nomenclature, Symbols: CTS, cardiotonic steroids; EPR, electron
paramagnetic resonance; ESI-HR-MS, high resolution electrospray ionization mass
spectrometry; J, coupling constant; SL, spin-label;
1. Abstract
The information obtained from crystallized complexes of the Na+,K+-ATPase with
cardiotonic steroids (CTS) is not sufficient to explain differences in the inhibitory properties
of CTS such as stereoselectivity of CTS-binding or effect of glycosylation on the preference
to enzyme isoforms. The uncertainty is related to the spacial organization of the hydrophilic
cavity at the entrance of the CTS-binding site. Therefore, there is a need to supplement the
crystallographic description with data obtained in aqueous solution, where molecules have
significant degree of flexibility. This work addresses the applicability of the electron
paramagnetic resonance (EPR) method for the purpose.
We have designed and synthesized spin-labelled compounds based on the cinobufagin
steroid core. The length of the spacer arms between the steroid core and the nitroxide group
determines the position of the reporting group (N-O) confined to the binding site. High
affinity to Na+,K+-ATPase is inferred from their ability to inhibit enzymatic activity. The
differences between the EPR spectra in the absence and presence of high ouabain
concentrations identify the signature peaks originating from the fraction of the spin-labels
bound within the ouabain site. The degree of perturbations of the EPR spectra depends on the
length of the spacer arm. Docking of the compounds into the CTS-site suggests which
elements of the protein structure might be responsible for interference with the spin-label
(e.g. steric clashes or immobilization). Thus, the method is suitable for gathering information
on the cavity leading to the CTS-binding site in Na+,K+-ATPase in all conformations with
high affinity to CTS.
2. Introduction
Na+,K+-ATPase is an enzyme critical for the survival of animal cells. Its activity
maintains an uneven distribution of Na+ and K+ ions over the membrane, the basis for the
resting membrane potential and excitability, as well as the driving force for transport of
nutrients and other ions [1]. The central role of the Na+,K+-ATPase in metabolism is reflected
in the evolutionary development of poisons and venoms targeting this enzyme. Most of them
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belong to a large family of cardiotonic steroids (CTS, or cardiac glycosides), and the
sensitivity towards these compounds is considered as a hallmark of Na+,K+-ATPase activity.
We have recently solved the crystal structures of the high affinity complexes of the α1β1
Na+,K+-ATPase with three different CTS: bufalin (non-glycosylated with 6-membered
lactone ring), ouabain (one sugar unit, 5-membered lactone ring) and digoxin (heavily
glycosylated, 5-membered ring) [2,3]. As expected, the ligands plug the entrance to the ion
binding sites from the extracellular side. The transmembrane helices forming the binding
pocket are highly superimposable (with the exception of the loop of αM1-2), consequently,
the binding modes of the ligands also revealed similar tracks.
The available structural information, however, is not sufficient to explain certain kinetic
observations. Thus, comparison of inhibitory properties of different stereoisomers of
bufadienolides showed that all β-stereoisomers are more potent compared to α-stereoisomers
[4]. Even for the bufalin molecule, the anomerization of the OH-group shifted the inhibition
curve and decreased affinity of binding to the α1β1 Na+,K+-ATPase from nM (β-form) to
tenth µM (α-form). It should be stressed that, according to the crystal structure, β-bufalin is
placed almost perpendicular to the membrane surface, with the lactone located in a
hydrophobic funnel, leading to the cation binding sites. There are no anticipations of
interactions around OH-group or clashes with the side chains in the vicinity of the entrance to
the binding site.
A positive effect of glycosylation on CTS affinity towards the Na+,K+-ATPase and its
contribution to Na+,K+-ATPase isoform selectivity (at least in the case of digoxin and
digitoxin) [5] represents another riddle, since in the crystal structure the sugar moiety is
found near extracellular loops, with no indications of firm interactions with the side chains.
Thus, there is a need to supplement the crystallographic description of the Na+,K+-ATPase-
CTS complexes with the data obtained in aqueous solution, where the molecules have a
certain degree of flexibility. This paper addresses the applicability of the electron
paramagnetic resonance (EPR) method for this purpose.
We have designed and synthesized spin-labelled cinobufagins where different spacer arms
allow variation in the distance between the steroid core and the nitroxide group. Thus, the
reporting group (N-O) is placed in different depths within the binding site. In an earlier study
spin-labeled ouabain was used to characterize motion of the whole oligomeric complex [6,7].
In the present paper, the degree of perturbation of EPR spectra, possibly due to increased
motional restriction, characterizes the binding cavity at the chosen distance from the steroid
core and correlates with the length of the spacer arm. This information together with results
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of the docking studies affords a description of the cavity leading to the CTS-binding site in
α1β1 Na+,K+-ATPase under physiological conditions.
3. Results and Discussion
Many CTS have 1-4 carbohydrate residues attached at 3β-OH of the steroid core, e.g.
ouabain has one and digoxin three sugar moieties, and all of these are found in the wide
cavity on the extracellular side of the Na+,K+-ATPase. Therefore the 3β-OH site is an obvious
choice for derivatization and attachment of a nitroxide group. Cinobufagin is used as a CTS-
core for the design of EPR probes. This ensures binding within the specific site, while spacer
arms varying in length and structure place the spin-label at different depths in the cavity. The
structures of seven synthesized EPR probes (C-1 to C-7) are presented in Fig. 1 (see
Materials and Methods for details of chemical synthesis and characterization of the EPR
probes).
3.1. Inhibitory activities of cinobufagin spin-labels on Na+,K+-ATPase.
The performed derivatization should not prevent specific binding of the compound in the
CTS binding site, i.e. the ability to inhibit enzyme with high affinity. Inhibitory potencies
were determined from the residual Na+,K+-ATPase hydrolytic activities after incubation with
varying concentrations of the derivatives using our reported method [4] (Fig. 2). The results
were fitted with hyperbolic functions corresponding to a dominant high-affinity component
and a second smaller low-affinity component. The resulting kinetic parameters are
summarized in Table 1. C-1 revealed a high inhibitory potency with a KDiss value of 0.15
0.004 μM for the high affinity component. The labels C-4 to C-7 all had KDiss values between
0.24 and 0.84 μM, whereas labels C-2 and C-3 had lower inhibitory activity. The inhibitory
potency of all labels was large enough to allow for a high degree of spin-labelling of the
enzyme at the specific site. The conformational equilibrium constant KC derived from the
inactivation patterns varies between about 2 and 6.7 (see Table 1), suggesting that between
67% and 87% of the enzyme molecules cannot release inhibitor directly, see Materials and
Methods and references therein for details of the kinetic analysis. The kinetic parameters are
discussed below in relation to heterogeneity of the EPR signals from bound inhibitor.
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3.2. EPR spectroscopy results
3.2.1 Verification of the method.
Theoretically, the spin-labelled compound added to the native membrane preparation of
the Na+,K+-ATPase may be localized in four different environments. Fig. 3 depicts Na+,K+-
ATPase (PDB ID 4HYT) embedded in the membrane and cinobufagin spin-labels are shown
as orange rectangles.
There are 4 possible loci, where binding to sites A, B and C might diminish the mobility
of the spin-label, and locus D refers to solvent phase, where the spin-label moves freely. Only
locus A represents the CTS binding site of the Na+,K+-ATPase and is, therefore, saturable.
Loci B, C and D are nonspecific and for all practical purposes non-saturable. Fractions of the
spin-label located in each locus will be characterized by signature peaks in the EPR spectrum,
dependent on their mobility. Ouabain at a high concentration will prevent binding of the
cinobufagin spin-label to the specific binding site (A), i.e. the signal from the spin-label in the
CTS-site will disappear from the EPR spectrum, but will not affect interactions with other
sites.
The above theoretical considerations were fully confirmed by the EPR spectra. Fig. 4
represents data for C-1 - C-3 added to the membranes containing Na+,K+-ATPase in the
presence or absence of ouabain. The spin-label concentration of 4 µM in Fig. 4 is chosen to
give an EPR signal with contributions primarily from the specifically bound label (Locus A).
From the inhibitory potencies (Table 1) it can be calculated that between 85-98% of the
labels C-1 - C-3 are bound to the specific binding sites. In the absence of ouabain, spectra of
all three spin-labels have broad peaks in the low-field region and these peaks are very much
reduced by addition of ouabain (red lines in Fig. 4).
The EPR spectra on Fig. 4 have been normalized (to 100%) to allow for a quantitative
comparison of the line shapes with and without ouabain present. For C-1 we find a KDiss
=
0.12 µM (Table 1), and with the used concentrations of spin-label (4 µM) and enzyme (12
µM) the amount of specifically bound spin-label reaches 98.2% (see legend to Fig. 4 and
section 4.4). The residual 1.8% of the spin-label is found in the non-specific sites. The cyan
line in Fig. 4A illustrates the negligible contribution from the spin-label bound to the non-
specific sites to the total spectrum (black line) under these conditions.
With the overwhelming majority of the spin-label bound to the CTS site, the shapes of
spectra in the low-field region points to a heterogeneous environment of the spin-label. For
C-1 there is a marked sharp peak in addition to a broader low-field line, while for C-2 and C-
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3 the peaks show significant overlap. Ouabain induces changes in the low field region
confirming that the heterogeneity originates from the binding modes in the CTS-site. It is
therefore possible that this heterogeneity in the EPR spectrum is related to the equilibrium
between the two complexes of the enzyme with spin-label bound: E·I ↔ E·I* (see Materials
and Methods for details).
Fig. 5 demonstrates the changes in the normalized spectra in response to increasing probe
(C-1) concentrations. In the presence of ouabain (Fig. 5A) the spectra have the same shape
independently on concentration used (from 12 to 72 µM in the presence of 36 µM enzyme).
This implies that the probe is to be found only in the non-saturable non-specific sites (B, C,
D) (while the CTS-binding site is occupied by ouabain) and that the relative proportions of B,
C and D are independent of the total concentration of spin-label (up to 72 µM).
In the absence of ouabain (Fig. 5B) the low-field peak changes height. The saturation of
the specific site changes between relatively low, intermediate and full saturation as the spin-
label concentration is increased. It is also evident that the proportion between non-
specifically vs. specifically bound probes increases.
The outer hyperfine splitting 2Amax for C-1 of about 67.9 G is the same as we earlier
found for the maleimide spin-labels (6-MSL in Table 1 in [8]), about 67.7 G. The linewidth
(here measured as width at half height) of the low-field peak is about 2.5 G compared to 3.7
G for 6-MSL [8] and the corresponding value is about 2.9 G for the high-field line, compared
to about 4.9 G for 6-MSL [8]. The lower values for C-1 indicate a more immobilized type
spectrum for C-1 than for the maleimide spin-label bound to Class II groups of the Na+,K+-
ATPase ([8]).
Based on the above results and considerations, the EPR signals observed from the probes
bound to the membranous preparation of the Na+,K+-ATPase were assigned to loci A, B and
C, see Fig. 5. Binding in site (A) results in intensity in the low-field region of the spectrum in
the absence of ouabain, and it appears that specific binding of C-1 leads to two distinct types
of signal (marked with arrows in Fig. 5B). The fraction of the spin-label dissolved in the
bilayer is heterogeneous since they occupy locus C, with rapid rotational motion reflected by
the sharpness of the line, and locus B, where the spin-label interacts with the protein
boundary which restricts the motion (see Fig. 5A). There is virtually no signal from the spin-
labels in the aqueous phase (D) for this label. In Fig. 5B the position of spin-label added in
excess of saturation of the specific site is also indicated (C).
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3.2.2 Correlation between the length of the spacer arm and perturbation of the
spin-label bound within the CTS-binding site.
The spin-labelled derivatives C-4 to C-7 comprise a family of homologous structures,
differing only in length of the spacer arms. The comparison is straight forward, since the
distance between the steroid core and the nitroxide spin-label increases gradually due to
addition of CH2 groups. Spectra in Fig. 6 are normalized for convenience and stacked
according to the length of the spin-label.
In the presence of ouabain (Fig. 6A) the spectra are roughly similar, and the positions of
signature peaks for labels in loci B and C are similar to those in Fig. 5, while the very sharp
peaks in C-4 and C-5 spectra are presumably due to spin-labels in the aqueous phase (locus
D).
In an attempt to quantitate this similarity the lineheights at about 3299 Gauss
(corresponding to locus C) and that at 3290.5 Gauss (corresponding to locus B) were read
from the ouabain-spectra of Figs. 4 and 6, and the signal ratio "3299 G"/"3290.5 G" was
calculated and shown in Fig. 6C with an average of 8.8 0.9. The linewidth of the sharp low-
field line (at locus B, here measured as the peak-to-trough distance) is also similar for the 7
labels (average about 2.9 G 0.3 G). The peak-to-peak distance of the sharp lines in the outer
wings of the spectra for the "type C" labels is also similar (average about 36.6 0.7 G, data
not shown). Thus, the overall spectral characteristics of non-specifically bound spin-labels
appear to be independent of the length of the spacer arm.
In the absence of ouabain, spectra vary significantly from analog to analog (Fig. 6B). In
these cases, the contributions from the non-specifically bound spin-labels can be estimated
using the spectra in Fig. 6A and the KDiss values from Table 1, as described in Section 4.6.
These responses are shown as dotted lines in Fig. 6B to illustrate a very small contribution
from the spin-labels in loci B, C and D to the overall EPR spectra obtained in the absence of
ouabain.
Comparison of the responses in the presence or absence of ouabain allows assignment of
the broad component in the low-field part of the C-4 spectrum to the signal from the probe
bound within CTS-site (locus A). The size and the shape of the low-field component show a
correlation with the length of the spacer arm. C-4 appears more affected (possible motionally
restricted) in the EPR spectrum compared to the longer labels C-5 to C-7.
For the labels C-5, C-6 and C-7 the linewidth of the sharp low-field line (here measured
as the peak-to-trough distance) shows a clear decrease from about 5 G for C-5 to about 3 G
for C-7, see Fig. 6C, possibly indicating an increase in motional freedom at the longer spacer
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arms. The peak-to-peak distance for the major peaks in the outer wings of the spectrum also
show a decrease (from about 44 G to about 39 G), also indicating gradually increased
motional freedom. The gradual increase in intensity ratio "3299 G"/"3290.5 G" in the row C-
5, C-6 and C-7 in the absence of ouabain suggests that there is very little low-field absorption
as the distance between the steroid nucleus and the spin-label moiety increases. A very small
fraction of the spin-label C-7 bound at the A-locus absorbs at low fields compared to the
conditions when C-7 is found only in loci B and C in the presence of ouabain (see Fig. 6 and
Fig. 3).
It is possible that the broad peak for C-4 is the sum of two close peaks. It is tempting to
interpret the results of the inactivation kinetics as follows: the bound spin-label is distributed
between two protein conformations E·I ↔ E·I* which are present at a 1:3 ratio (Kc=3, see
Table 1). These two will have absorption maxima in the low-field part of the EPR spectrum
in the 3290 - 3296 G range (cf. Fig. 6B). We are currently undertaking a lineshape analysis of
this broad peak to interpret the results of the inactivation kinetics with the aim of describing
the difference between the structure of the initial binding state E·I and the fully bound form
E·I*.
In summary, the consistent changes in the spectral characteristics of the spin-labeled
derivatives C-4 to C-7 possibly indicate an increase in mobility as the length of the spacer
arm in increased. The shortest spacer arm (in C-4) correlates with the most motionally
restricted (broad) peak in the low-field part of the spectrum.
3.3 Docking of cinobufagin derivatives into the CTS-binding site of Na+,K+-
ATPase.
The crystal structure of pig kidney Na, K-ATPase in complex with bufalin (PDB ID
4RES) was used for the docking of spin-labeled derivatives. Fig. 7 shows that cinobufagins
have a binding mode similar to that of bufalin in the crystal complex. It seems, however, that
orientation of the steroid core of cinobufagin in the binding cavity might vary slightly and
that the substituent (linker and spin-label) has a tendency to pull the molecule upwards in the
cavity compared to bufalin.
Variations in the position of the spin-label in the upper part of the cavity were more
significant for C-7 (Fig. 7B) compared to C-4 (Fig. 7A). Potential ligand poses generated by
the docking program placed the spin-label group in C-4 in the proximity to the extracellular
loop M1-M2. The question, whether a spatial restriction or a specific interaction with the
protein is responsible for this relatively rigid binding mode, remains unresolved.
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For the C-7 derivative the long linker either brings the spin-label well out into the
extracellular matrix or bends and directs it towards the protein cavity. The former orientation
yields probably the very sharp EPR peak at 3299 G. The spin-labels interacting within the
protein cavity could give rise to low-field intensity in the EPR spectra suggesting lower
mobility. The very high ratio "3299 G"/"3290.5 G" for C-7 (cf. Fig. 6C), however, suggests
that only a small fraction of these spin-labels are actually located within the cavity. We have
attempted to illustrate this in Fig. 7C, where a set of C-7 spin-labels illustrate the more
extended forms of the C-7 spin-labels. Here the steroid moiety is docked as in panels A and
B, but the spin-label group is allowed to stretch into the aqueous phase. It is conceivable that
these spin-labels will experience a much larger degree of motional freedom than for example
C-4 (cf. Fig. 7A), giving rise to the dominant sharp lines shown for C-7 in Fig. 6B.
In conclusion, the results illustrate that the EPR approach is suitable for detailed
characterization of the extracellular cavity of the Na+,K+-ATPase CTS-binding site. It
supplements and verifies the existing description of crystallized enzyme-CTS complexes
since it is able to provide information valid under physiological conditions in aqueous
solutions. The degree of perturbation of the spin-label reflects the interactions within the
cavity at the chosen distance from the steroid core, that could e.g. limit its mobility. This
interaction, in turn, may be identified by docking studies. Compilation of the data will allow
mapping of the surface of the CTS binding site. Since both the chemical structure and the
length of the spacer affect the spectral characteristics of the nitroxide label we expect to
identify the specific amino acid residues as points of contact.
4. Materials and Methods
4.1. General synthetic procedures.
We used the nitroxyl free radical as reporting group, and attached it to the steroid core of
cinobufagin through different linkers. Cinobufagin was purchased from Shanxi Chenguang
Biological Technology Co. Ltd with purity ≥ 98%. 4-hydroxy-2,2,6,6-tetramethylpiperidine
(4-OH-TEMPO) was purchased from Aladdin (Shanghai, China) and used without further
purification. All other commercially available reagents were used as received unless stated
otherwise. NMR data were collected on a Bruker® 300 MHz spectrometer at ambient
temperature with the residual solvent peaks as internal standards. Chemical shifts were given
in ppm with respect to the TMS signal and coupling constants (J) were given in Hz. ESI-HR-
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MS were obtained using Agilent 6210 LC/MSD TOF mass spectrometer (Palo Alto, USA).
EPR spectra were obtained with JES-FA200 X-band EPR spectrometer at ambient
temperature. Optical rotations were measured using a JASCO P-2000 polarimeter (Tokyo,
Japan) at 22 to 24 °C in CH3OH. IR spectra were obtained using a JASCO FT/IR-4600
Fourier Transform Infrared spectrometer. The 200-300 and 300-400 mesh silica gel (Qingdao
Ocean Chemical Factory, P. R. China) was used for TLC and column chromatography,
respectively. All air- or moisture-sensitive reactions were operated using dry solvents under
argon atmosphere.
4.2. Synthesis of nitroxides with different linkers 3, 4 and 6a-d.
The synthesis routes are outlined in Fig. 8.
Synthesis of compounds 2a-d. A suspension of compound 1a-d (1 g) in methanol (20
mL) was added thionyl chloride at 0°C under argon atmosphere. Then, the mixture was
allowed to reach room temperature and stirred for 16h. After the reactant was consumed
completely, the solution was evaporated to dryness under reduced pressure, and the residue
was re-dissolved in 10 mL methanol and then concentrated. The above procedure was
repeated twice to remove thionyl chloride in excess. At last, the crude material was
recrystallized from ethyl acetate to give the white solid compound 2a-d.
Synthesis of compound 3. To a solution of 4-OH-TEMPO (100 mg, 0.58 mmol, 1.0
equiv.), Bromoacetic acid (121 mg, 0.87 mmol, 1.5 equiv.) and DMAP (14 mg, 0.12 mmol,
0.2 equiv.) in 5 mL anhydrous dichloromethane was added EDCI (222 mg, 1.2 mmol, 2.0
equiv.), then the solution was stirred at room temperature under atmosphere overnight. The
reaction mixture was diluted with 30 mL dichloromethane and extracted with water (50 mL).
The aqueous layer was extracted with dichloromethane (20 mL) twice. The combined organic
layer was washed with saturated NaHCO3 solution (3×10 mL) and saturated NaCl solution
(3×10 mL), then dried over anhydrous sodium sulfate and filtered. The resulting solution was
concentrated in vacuum to give the red liquid, which was purified by column chromatography
on silica gel (petroleum ether / ethyl acetate = 20:1) to afford compound 3 (158 mg, 0.54
mmol, 93%) as red solid powder.
Synthesis of compound 4. 4-OH-TEMPO (1 g, 5.8 mmol, 1.0 equiv.) was added slowly
to the suspension of NaH (697 mg, 60% in mineral oil, 11.6 mmol, 3.0 equiv.) in anhydrous
THF (200 mL) at 0°C. Subsequently, bromoacetic acid was added, then the solution was
heated under gentle reflux under argon for 18h. After the reaction mixture was cooled to
room temperature, water (20 mL) was added carefully, the resultant dark orange solution was
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concentrated to 50 mL and transferred to a separation funnel. The aqueous layer was washed
with diethyl ether (3×20 mL) and dichloromethane (3×20 mL) in sequence and then acidified
with 10% acetic acid to pH 6-7. The acidified aqueous layer was extracted with
dichloromethane (3×30 mL). The combined organic layer was dried with anhydrous sodium
sulfate, filtered and concentrated under reduce pressure. The residue was purified by flash
silica gel column chromatography (chloroform / methanol = 15:1) to achieve compound 4
(978 mg, 4.3 mmol, 73%) as red oil.
Synthesis of compounds 5a-d. Compound 1 (200 mg, 0.9 mmol, 1.0 equiv.) and the
corresponding compound 2a-d (1.2 equiv.) were dissolved in a solution consisting of 3 mL
anhydrous DMF and 1 mL anhydrous dichloromethane, then 0.2 mL 4-methylmorpholine
(NMM, 176 mg, 1.7 mmol, 2.0 equiv.) was added. After the mixture was cooled to 0 °C
using ice bath, 0.15 mL isobutyl chloroformate (IBCF, 1.1 mmol, 1.3 equiv.) was added
dropwise. The above solution was stirred overnight under argon after warming to room
temperature. The reaction mixture was extracted with dichloromethane (25 mL) twice, and
then the combined organic layer was washed with water (3×10 mL), saturated NaHCO3
solution (3×10 mL) and saturated NaCl solution (3×10 mL), dried over anhydrous sodium
sulfate, filtered and concentrated under reduced pressure. The residue was subjected to silica
gel column chromatography (cyclohexane / ethyl acetate = 3:1) to afford compound 5a-d as
red oil.
Synthesis of compounds 6a-d. This synthesis was done using the reported method [9].
To a solution of compound 5a-d (100 mg) in 2 mL methanol was added 60 mg LiOH in 1 mL
water, then the reaction mixture was stirred at room temperature for 3-4h. After the reaction
was completed as monitored by TLC, the resultant solution was concentrated to remove the
methanol. The residue was dissolved in dichloromethane and then extracted. The aqueous
layer was carefully acidified with 1N hydrochloric acid to pH 2-3 then extracted with
dichloromethane (20 mL) twice. The combined organic layer was washed with water (3×10
mL) and saturated NaCl solution (3×10 mL), dried with anhydrous sodium sulfate, filtered
and concentrated in vacuum to obtain the compound 6a-d as red powder. The physical and
spectroscopy characters of compounds 3, 4 and 6a-d were listed in Table S1.
4.3. Synthesis of spin-labeled cinobufagins C-1 to C-7.
The synthesis routes are outlined in Fig. 9.
Synthesis of compound 7. Succinic anhydride (23 mg, 0.23 mmol, 2.0 equiv.) was added
to a solution of cinobufagin (50 mg, 0.13 mmol, 1.0 equiv.) and DMAP (28 mg, 0.23 mmol,
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2.0 equiv.) in 5 mL anhydrous dichloromethane, the solution was refluxed gently under argon
for 8h. After the reactant was consumed completely, the solution was diluted with 20 mL
dichloromethane and extracted with water. The organic layer was washed with water (3×10
mL) and saturated NaCl solution (3×10 mL), dried with anhydrous sodium sulfate, filtered
and then concentrated under reduced pressure to give compound 7 as white solid (52 mg, 95
μmol, 85%). The NMR data of 7 was similar to the parent compound cinobufagin [10]
except for the side chain at C-3 [2.73-2.58 (4H, m) for the two methylene and downfiled shift
1
of H-3 in the 1H-NMR spectrum]. H NMR (300 MHz, CDCl3) δ: 7.91 (1H, d, J = 9.8 Hz,
H-22), 7.17 (1H, s, H-21), 6.22 (1H, d, J = 9.8 Hz, H-23), 5.51-5.42 (1H, m, H-16), 5.13 (1H,
s, H-3), 3.65 (1H, d, J = 1.3 Hz, H-15), 2.80 (1H, d, J = 9.3 Hz, H-17), 2.73-2.58 (4H, m, -
CH2CH2-), 1.89 (3H, s, 16-OAc), 0.99 (3H, s, H-19), 0.81 (3H, s, H-18). ESI-HR-MS calcd
for C30H37O9 [M-H]+: 541.2443, found 541.24449.
Synthesis of compound C-1. To a solution of cinobufagin (30 mg, 68 μmol, 1.0 equiv.)
and DMAP (2 mg, 14 μmol, 0.2 equiv.) in 3 mL anhydrous dichloromethane was added
compound 4 (19 mg, 81 μmol, 1.2 equiv.), the solution was cooled to 0 °C using ice bath and
then 20 μL DIC (17 mg, 136 μmol, 2.0 equiv.) was added dropwise. After the addition was
completed, the reaction mixture was allowed to reach room temperature and stirred under
argon for 24h. Then the solution was diluted with 15 mL dichloromethane and extracted with
water. The organic layer was washed with saturated NaHCO3 solution and saturated NaCl
solution in sequence, dried with anhydrous sodium sulfate, filtered and evaporated to dryness.
The resulting residue was subjected to flash silica gel column chromatography (petroleum /
ethyl acetate = 5:1) give the compound C-1 (33 mg, 50 μmol, 74%) as red powder.
Synthesis of compound C-2. Compound 7 (20 mg, 37 μmol, 1.0 equiv.), 4-OH-TEMPO
(7 mg, 40 μmol, 1.1 equiv.) and DMAP (2 mg, 18 μmol 0.5 equiv.) were dissolved in 3 mL
anhydrous dichloromethane, then EDCI (15 mg, 74 μmol, 2.0 equiv.) was added. The above
solution was stirred under argon at room temperature overnight. Then the reaction mixture
was diluted with 15 mL dichloromethane and extracted with water. The organic layer was
washed with saturated NaHCO3 solution and saturated NaCl solution, dried with anhydrous
sodium sulfate, filtered and concentrated in vacuum. The residue was purified by silica gel
column chromatography (petroleum / ethyl acetate = 3:1) to afford compound C-2 (22 mg, 32
μmol, 86%) as light red powder.
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Synthesis of compound C-3. To a solution of compound 7 (30 mg, 55μmol, 1.0 equiv.)
in 3 mL DMF was added anhydrous potassium carbonate (15 mg, 110μmol, 2.0 equiv.), the
solution was stirred at room temperature for 10 min. Subsequently, compound 3 (20 mg, 66
μmol, 1.2 equiv.) was added and the mixture was stirred for another 10h. After the reactant
was consumed, the reaction mixture was diluted with 20 mL ethyl acetate and extracted with
water. The organic layer was washed with water, saturated NaHCO3 solution and saturated
NaCl solution, dried with anhydrous sodium sulfate, filtered and concentrated in vacuum.
The residue was purified by silica gel column chromatography (cyclohexane / ethyl acetate =
5:1) to give compound C-3 (32 mg, 42 μmol, 77%) as red powder.
Synthesis of compounds C-4 to C-7. To a solution of cinobufagin (30 mg, 68 μmol, 1.0
equiv.) and DMAP (2 mg, 14 μmol, 0.2 equiv.) in 3 mL anhydrous dichloromethane was
added the corresponding compound 6a-d (1.2 equiv.), the solution was cooled to 0 °C using
ice bath and then 20 μL DIC (17 mg, 136 μmol, 2.0 equiv.) was added dropwise. After the
addition was completed, the reaction mixture was allowed to reach room temperature and
stirred under argon for 24h. Then the solution was diluted with 15 mL dichloromethane and
extracted with water. The organic layer was washed with saturated NaHCO3 solution and
saturated NaCl solution in sequence, dried with anhydrous sodium sulfate, filtered and
evaporated to dryness. The resulting residue was subjected to flash silica gel column
chromatography (petroleum / ethyl acetate = 2:1) give the compounds C-4 to C-7 as red
powders. Owing to the paramagnetic nature of the spin-labels, it is difficult to gain valuable
structural information by NMR spectroscopy. In order to prove the presence of nitroxide
groups, we collected the EPR spectra of the target compounds, which showed triplet with the
hyperfine coupling constants in the range of 16.00-16.45 Gauss due to the interaction
between the unpaired electron and nitrogen nucleus. The structures of all the target
compounds were further identified by ESI-HR-MS and IR spectral analysis. Optical rotations
were also determined. The physical and spectroscopy characters of compounds C-1 to C-7
are listed in Tables S1, S2 and S3.
4.4. Preparation and activity assay of Na+,K+-ATPase.
Na+,K+-ATPase from pig kidney microsomal membranes was purified by differential
centrifugation [11]. The stock solution has protein concentration of 5-6 mg/ml in 250 mM
sucrose, 20 mM histidine, 0.9 mM EDTA (pH 7.0). The specific Na+,K+-ATPase activity was
determined from the difference in the amount of phosphate released in the absence and
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presence of 1 mM ouabain in a solution containing 130 mM NaCl, 20 mM KCl, 4 mM MgCl2
and 3 mM ATP. The specific activity of the enzyme preparation was approximately 30 µmol
o
Pi/mg protein released per min at 37 C [11]. The phosphorylation capacity is about 2.9
nmol/mg protein and is equal to the high-affinity ouabain binding capacity [4,11].
4.5. Details of the cinobufagin spin-label inhibition experiments.
Stock solutions of the inhibitors, typically 10 mM, were prepared in DMSO.
The inhibitory effects of cinobufagin spin-labels on Na+,K+-ATPase were determined
essentially as previously reported [4]. In brief, Na+,K+-ATPase is preincubated at 37 oC for 3
hours in the presence of 3 mM MgCl2, 3 mM NMG-phosphate and 40 mM Tris (pH 7.0) with
increasing concentrations of inhibitor. For each inhibitor concentration the residual Na+,K+-
ATPase activity is subsequently determined by 40-fold dilution into a standard ATPase assay
medium. Measurements were performed in triplicate and expressed in percent of the Na+,K+-
ATPase activity in the absence of inhibitor (see legend to Fig. 2 for details).
4.6. Details of the kinetic analysis of inhibition.
The interaction between cinobufagins (I) and Na+,K+-ATPase (E) is interpreted as a two-
step process:
E + I ↔ E·I ↔ E·I*
where both complexes E·I and E·I* are enzymatically inactive and in relatively slow
equilibrium (details of this model can be found in refs. [3, 4]). The binding of I to E is
described by a dissociation constant Ki = [E] · [I]/[E·I]. The conformational change E·I ↔
E·I* is described by an is described constant KC= [E·I*]/[E·I].
The data are fitted by a non-linear least squares function relating the Na+,K+-ATPase
activity (Activity(I)) at a given concentration of inhibitor [I] to a sum of two hyperbolic terms
containing the parameters for the high-affinity as well as the low-affinity components of
inhibition:
Activity(I) = 100% - Fhigh [I]/( KDiss,high + [I]) - Flow [I]/( KDiss,low + [I])
·
·
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The two components have magnitudes of Fhigh and Flow (in %). The dissociation constant
derived from the curve fitting (KDiss,high) is related to Ki and KC = Fhigh/Flow through the
equation KDiss,high = Ki/(Kc + 1).
Following the above analysis, the relative magnitude of the high- and low-affinity
component is a measure of KC.
The 7 inhibitors have widely different kinetic properties, and it is illustrative to consider
one of them in detail. Compound C-4 inactivates the Na+,K+-ATPase with about 75% (=Fhigh
)
of the activity being very sensitive towards the inhibitor and the remaining activity is not
affected by further increase in inhibitor concentration (there is virtually no low-affinity
inactivation in the concentration ranges studied here, KDiss,low is thus too large to be
determined in the present assay, see Fig. 2 and Table 1). The simplest interpretation of the
above finding is that the equilibrium between E·I and E·I* is displaced 3.00 fold (= Kc)
towards E·I*, which represents 75% of the enzyme. Ki = KDiss,high · (Kc + 1) = 0,952 µM,
indicating a much stronger initial binding of this compound than for compounds C-2 to C-7
(see Table 1).
4.7. Binding of cinobufagin spin-labels to Na+,K+-ATPase.
In a typical experiment Na+,K+-ATPase stock solution is diluted to 4 mg/ml (12 µM) with
Tris-phosphate buffer and spin-label to give final concentrations of 40 mM Tris, 3 mM
MgCl2, 3 mM NMG-Pi and 4 µM spin-label. For ouabain-samples the solution also contains
0.5 mM ouabain. In some experiments higher enzyme and spin-label concentrations were
used as indicated in the figures.
o
Cinobufagin spin-label binding is carried out at 37 C for 3 hours and the reaction is
started by addition of spin-label. Ouabain-containing samples were preincubated with 0.5
mM ouabain at 37 oC for 30 min before addition of spin-label.
o
After incubation the samples were cooled and centrifuged 26 500 g at 10 C for 30 min.
o
Pellets were transferred to quartz capillaries with inner diameter 1 mm and stored at -20 C.
The samples used for EPR were all subjected to centrifugation in order to increase the
concentration of enzyme in the EPR tubes (Figs. 4-6). The effect of centrifugation is shown
in Fig. 10. The overall shapes of the spectra are very similar, suggesting that centrifugation
does not alter the EPR characteristics. In the spectra of the sample before centrifugation (red
lines) a small sharp component is visible especially in the high-field section section (marked
with D in Fig. 10A and Fig. 10B). After centrifugation this peak is almost absent (black
lines). This suggests that spin-label in the aqueous phase gives rise to this sharp peak. The
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enzyme concentration in the pellet is 5-10 times larger than before centrifugation, and the
relative contribution from free spin-label in the pellet will thus be negligible (in these
normalized spectra) compared to the situation before centrifugation.
In Fig. 4A a comparison is made between the spectrum in the absence of ouabain and a
spectrum representing the spin-label not bound to the specific sites. The amount of spin-label
bound to the specific site is calculated using the equation
2
ꢁ
E
+I +K
tot tot
- (E +I +K
) -4E
I
Diss
tot tot
Diss
tot tot
EI ꢀ
, where Etot and Itot are the total
2
concentrations of the enzyme and the spin-label, respectively. For Etot = 12 µM, Itot = 4 µM
and KDiss=0.18 µM (see Table 1) we find EI/Itot = 0.982 (=98.2%) and the concentration of
spin-label not bound to the specific sites thus amounts to 1.8% (indicated by the cyan line in
Fig. 4A). The contribution of non-specifically bound spin-label to the full spectrum is
similarly indicated with cyan lines in Figs. 4B and 4C, and by dotted lines in Fig. 6B.
4.8. EPR spectroscopy.
EPR measurements were carried out on a 9-GHz Bruker EMX plus EPR spectrometer
(Bruker Biospin, Rheinstetten, Germany). All spectra were recorded at 25 °C and the
temperature was set by the internal temperature controller of the spectrometer. The samples
were placed into thin glass capillaries (~1ꢀmm inner diameter, BLAUBRAND micropipettes),
these were placed in Wilmad Lab Glass standard 4 mm OD quartz EPR tube (Rotec Spintec
GmbH, Griesheim, Germany) which were then mounted in the spectrometer. EPR spectra
were recorded at X-band frequency (9.4 GHz) with a microwave power of 20 mW and a
modulation frequency 100 kHz. The sweep width was 100 G, modulation amplitude 1.25 G,
time constant 10 ms. The typical sweep time was 60 s, and 16 scans were summed in order to
improve the signal-to-noise ratio. Spectra were characterized using the approaches in ref.
[12].
4.9. Docking procedures.
The crystal structure of Na+,K+-ATPase in complex with bufalin (PDB ID 4RES) was
used for docking of spin-labeled cinobufagins [3]. The docking model was prepared using the
Protein Preparation Wizard in the Schrödinger Suite 2012. The protonation states of the
titratable groups in the structure were deduced from pKa values calculated in PROPKA.
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Bufalin and spin-labeled cinobufagins were built in Maestro version 11.2 and the
protonation state of each compound was predicted in Epik. A conformational search was
performed by the Advanced Search procedure using the OPLS3 force field. Each step in the
conformational search was followed by a 2500 step minimization or until the minimization
reached convergence using a conjugate gradient algorithm. The obtained lowest-energy
conformation of each compound was used for docking.
A receptor grid centered on the bufalin binding site was generated and docking was
performed in Glide. The docking included flexible ligand sampling and was set up to retrieve
a maximum of 100 poses. The final scoring was achieved using the XP protocol, all other
settings were default. Docking of bufalin and cinobufagin placed both ligands in the position
of bufalin in the crystal structure, thus verifying the docking approach. Docking of bufalin,
cinobufagin and spin-labeled cinobufagins resulted in scores around -7.
5. Acknowledgements
This work was supported by grants from the National Natural Science Foundation of
China (81573315, to R. W. J.), Guangdong Natural Science Fund (2015A030313313, to R.
W. J.), Guangzhou Industry-University Collaborative Innovation Major Projects (to R. W. J.),
Danish Council for Independent Research (DFF-7016-00125, to N.U.F.) and The A.P. Møller
Foundation for the Advancement of Medical Science (to N.U.F.). We are thankful to B.
Bjerring Jensen for excellent technical assistance and to Dr. Derek Marsh for many helpful
comments on the manuscript. Lars Duelund (University of Southern Denmark) is gratefully
acknowledged for assistance with the EPR spectrometer, and the Department of Chemistry
and Pharmacology at University of Southern Denmark is thanked for making EPR data
collection possible.
6. Author Contributions
J-HG synthesized and characterized the spin-labels. ME and NUF performed inactivation
and EPR experiments and data analysis, and JLA performed the docking experiments. R-WJ,
ME and NUF designed and supervised the experiments. The paper was written and proofread
by all authors.
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7. References
1. Glynn IM (2002) A hundred years of sodium pumping. Annu. Rev. Physiol. 64, 1-18.
2. Laursen M, Yatime L, Nissen P & Fedosova NU (2013) Crystal structure of the high-
affinity Na+,K+-ATPase–ouabain complex with Mg2+ bound in the cation binding site. Proc.
Natl. Acad. Sci. 110, 27 10958-10963.
3. Laursen M, Gregersen JL, Yatime L, Nissen P & Fedosova NU (2015) Structures and
characterization of digoxin- and bufalin-bound Na+,K+-ATPase compared with the ouabain-
bound complex. Proc. Natl. Acad. Sci. 112(6), 1755-1760.
4. Tang H, Ruan L, Tian H, Liang G-P, Ye W-C, Hughes E, Esmann M, Fedosova NU,
Chung T-Y, Tzen JTC, Jiang R-W & Middleton DA (2016) Novel stereoselective
bufadienolides reveal new insights into the requirements for Na+,K+-ATPase inhibition by
cardiotonic steroids. Sci.Rep. DOI:10.1038/srep29155.
5. Katz A, Lifshitz Y, Bab-Dinitz E, Kapri-Pardes E, Goldshleger R, Tal DM & Karlish SJ
(2010) Selectivity of digitalis glycosides for isoforms of human Na+,K+-ATPase. J Biol
Chem. 285:19582-92.
6. Solomonson LP & Barber MJ (1984) Spin-labeled ouabain as a probe for cardiac glycoside
receptor/Na+,K+-ATPase. Biochem. Biophys. Res. Commun. 124:210-216.
7. Mahaney JE & Grisham CM (1992) Effect of ouabain on the rotational dynamics of renal
Na+,K+-ATPase studied by saturation-transfer EPR. Biochemistry 31:2025-2034.
8. Esmann M, Hideg K & Marsh D (1992) Conventional and saturation transfer EPR
spectroscopy of Na+,K+-ATPase modified with different maleimide nitroxide derivatives.
Biochim. Biophys. Acta 1159: 51-59.
9. Zakrzewski J & Krawczyk M (2014) Reactions of nitroxides XIV. Analogs of phenoxy
carboxylic herbicides based on the piperidine scaffold; unexpected fungicidal activity of the
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2-[(1-oxyl-2,2,6,6-tetramethylpiperidin-4-yl)oxy]butanoic
acid.
Heterocyclic
Communications 20: 89-91.
10. He X, Tang J, Qiao A, Wang G, Jiang M, Liu R & Yiao X (2006) Cytotoxic
biotransformed products from cinobufagin by Mucor spinosus and Aspergillus Niger.
Steroids 71: 392-402.
11. Klodos I, Esmann M & Post RL (2002) Large-scale preparation of sodium-potassium
ATPase from kidney outer medulla. Kidney Int. 62:2097-2100.
12. Marsh D (1981) Electron Spin Resonance: Spin Labels. Membrane Spectroscopy (ed.
Ernst Grell) Springer-Verlag Heidelberg.
8. Supporting Information
Supplementary Material Tables
Table S1 : Physical and spectroscopy characters of compounds 3, 4 and 6a-d.
Table S2 : Physical and spectroscopy characters of compounds C-1 to C-7.
Table S3 : IR spectroscopy characters of compounds C-1 to C-7.
Supplementary Material Figures
Figure S1 : ESI-HR-MS of compound 3.
Figure S2 : ESI-HR-MS of compound 4.
Figure S3 : ESI-HR-MS of compound 6a.
Figure S4 : ESI-HR-MS of compound 6b.
Figure S5 : ESI-HR-MS of compound 6c.
Figure S6 : ESI-HR-MS of compound 6d.
Figure S7 : ESI-HR-MS of compound 7.
Figure S8 : 1H NMR spectrum of compound 7.
Figure S9 : ESI-HR-MS of compound C-1.
Figure S10 : IR spectrum of compound C-1.
Figure S11 : ESI-HR-MS of compound C-2.
Figure S12 : IR spectrum of compound C-2.
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Figure S13 : ESI-HR-MS of compound C-3.
Figure S14 : IR spectrum of compound C-3.
Figure S15 : ESI-HR-MS of compound C-4.
Figure S16 : IR spectrum of compound C-4.
Figure S17 : ESI-HR-MS of compound C-5.
Figure S18 : IR spectrum of compound C-5.
Figure S19 : ESI-HR-MS of compound C-6.
Figure S20 : IR spectrum of compound C-6.
Figure S21 : ESI-HR-MS of compound C-7.
Figure S22 : IR spectrum of compound C-7.
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9. Tables
Table 1. Kinetic parameters of cinobufagin spin-label interactions with Na+,K+-
ATPase extracted from the inhibition curves (see section 4.6 for details).
Number
of linker
atomsa
Residual
activityb
(%)
Spin-
label
KDiss,high SE
(µM)
Ki
KC
(µM)
C-1
C-2
C-3
C-4
C-5
C-6
C-7
4
6
0.147 0.004
1.24 0.414
1.55 0.092
0.238 0.024
0.366 0.078
0.356 0.056
0.842 0.278
18
13
28
25
33
30
30
4.56 0.817
6.69 9.59
2.57 5.55
3.0 0.952
2.03 1.11
2.33 1.19
2.33 2.80
9
7
8
9
11
a The numbers give the atoms linking the steroid skeleton to the nitroxide. See also
structures in Fig. 1.
b The percentages of low-affinity components of the inhibition curves is represented
as residual activity.
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Fig. 1. Structures of seven cinobufagin spin-labels.
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Fig. 2. The inhibitory potencies of spin-labelled cinobufagins against pig kidney
Na+,K+-ATPase at 37 °C. (A) Dose-response curves showing the ATPase activity
remaining at the specified concentrations of C-1 to C-3. (B) Dose-response curves
for C-4 to C-7. The solid lines represent hyperbolic fits to the inactivation measured
after preincubation of the inhibitors with Na+,K+-ATPase for 3 hours at 37 °C in the
o
presence of Mg2+ and Pi. Vmax (=100%) equals 29 units/mg at 37 C. Each point
represents the average value from three independent experiments SE.
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Fig. 3. Sites for interaction of cinobufagin spin-labels with Na+,K+-ATPase.
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Fig. 4. EPR spectra of spin-labelled pig kidney Na+,K+-ATPase. (A) The normalized
spectra for a sample with enzyme (12 µM) in buffer and 4 µM C-1 in the absence
(black line) and in the presence of 0.5 mM ouabain (redline). In addition, the
spectrum with ouabain is also shown scaled to 1.8% (cyan line). (B) The normalized
spectra for a sample with enzyme (12 µM) in buffer and 4 µM C-2 in the absence
(black line) and in the presence of 0.5 mM ouabain (red line). In addition, the
spectrum with ouabain is also shown scaled to 12.8% (cyan line). (C) The
normalized spectra for a sample with enzyme (12 µM) in buffer and labelled with 4
µM C-3 in the absence (black line) and in the presence of 0.5 mM ouabain (red line).
In addition, the spectrum with ouabain is also shown scaled to 15.3% (cyan line).
Samples were centrifuged before EPR. Temperature 26 oC. Total scan width 100 G.
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Fig. 5. EPR spectra of spin-labelled pig kidney Na+,K+-ATPase. (A) Spectra are for
samples with enzyme (36 µM) in buffer and 0.8 mM ouabain labelled with C-1 at
concentrations of 12 µM (black line), 36 µM (red line) and 72 µM (cyan line),
respectively. The approximate positions of loci B and C are indicated. (B) Spectra
are for the same samples in the absence of ouabain. Samples were centrifuged
before EPR. The approximate positions of locus A are indicated both in the low-field
and the high-field regions. Temperature 26 oC. Total scan width 100 G.
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Fig. 6. EPR spectra of spin-labelled pig kidney Na+,K+-ATPase. (A) Spectra are for
samples with enzyme (12 µM) in buffer and 0.5 mM ouabain, labelled with 4 µM C-4
(black), C-5 (red), C-6 (cyan) and C-7 (blue) respectively (top to bottom). The peaks
marked "B", "C" and "D" in the C-4 spectrum are related to non-specifically bound
spin-label at loci B or C or label in aqueous solution (locus D, see Fig. 3). (B)
Spectra are for samples with enzyme (12 µM) in buffer labelled with 4 µM C-4
(black), C-5 (red), C-6 (cyan) and C-7 (blue) respectively (top to bottom) in the
absence of ouabain. In addition, calculated spectra from non-specifically bound
labels are shown as dotted lines. The approximate positions of the peaks originated
from locus A are indicated. Samples were centrifuged before EPR. Temperature 26
oC. Total scan width 100 G. (C) The ratio of the lineheights at about 3299 Gauss and
at 3290.5 Gauss (read from spectra shown in Figs. 4 and 6) are shown for the 7 spin
labels C-1 to C-7 in the prescence of 0.5 mM ouabain (open black squares) as well
as in the absence of ouabain (for C-5 to C-7, filled black squares). In addition the
linewidth of the sharp line at about 3299 G is given for spin-label in the prescence of
0.5 mM ouabain (open red circles) as well as in the absence of ouabain (for C-5 to
C-7, filled red circles).
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Fig. 7. Binding of the spin-labeled cinobufagins C-4 and C-7 within the CTS-binding
site of Na+,K+-ATPase. Transmembrane helices and loops on the extracellular side
of the α- and β-subunits of the enzyme in the E2P-bufalin complex (PDB ID 4RES)
are depicted in grey and green, respectively. Purple spheres represent K+ ions in
sites I and II. Five representative poses are depicted for C-4 (panel A) and C-7
(panel B) revealing possible variations in their binding modes compared to that of
bufalin (cyan sticks) in the crystal structure. Panel C depicts a set of more extended
forms of C-7. The steroid moiety is docked as in panels A and B, but the spin-label
group is stretched into the aqueous phase.
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Fig. 8. Synthetic procedure of nitroxides with different linkers. Reagents and
conditions: (a) SOCl2, CH3OH, 0 °C -r.t. (b) Bromoacetic acid, EDCI, DMAP, DCM,
r.t. (c) Bromoacetic acid, NaH, THF, reflux. (d) IBCF, NMM, DMF/DCM, 0 °C -r.t. (e)
LiOH, CH3OH/H2O, r.t.
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Fig. 9. Synthetic procedure of spin-labeled cinobufagin. Reagents and conditions: (a)
DIC, DMAP, DCM, 0 °C - r.t. (b) Succinic anhydride, DMAP, DCM, reflux. (c) 4-OH-
TEMPO, EDCI, DMAP, DCM, r.t. (d) compound 3, K2CO3, DMF, r.t.
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