Alkylphenol inverse agonists of HCN1 gating: H-bond propensity, ring\nsaturation and adduct geometry differentially determine efficacy and\npotency
Background and purpose In models of neuropathic pain, inhibition of HCN1 is anti-hyperalgesic. 2,6-di-iso-propyl phenol (propofol) and its non-anesthetic congener, 2,6-di-tert-butyl phenol, inhibit HCN1 channels by stabilizing closed state(s). Experimental approach Using in vitro electrophysiology and kinetic modeling, we systematically explore the contribution of ligand architecture to alkylphenol-channel coupling. Key results When corrected for changes in hydrophobicity (and propensity for intra-membrane partitioning), the decrease in potency upon 1-position substitution (NCO~OH >> SH >>> F) mirrors the ligands\xe2\x80\x99 H-bond acceptor (NCO > OH > SH >>> F) but not donor profile (OH > SH >>> NCO~F). H-bond elimination (OH to F) corresponds to a \xce\x94\xce\x94G of ~4.5 kCal mol\xe2\x88\x921 loss of potency with little or no disruption of efficacy. Substitution of compact alkyl groups (iso-propyl, tert-butyl) with shorter (ethyl, methyl) or more extended (sec-butyl) adducts disrupts both potency and efficacy. Ring saturation (with the obligate loss of both planarity and \xcf\x80 electrons) primarily disrupts efficacy. Conclusions and implications A hydrophobicity-independent decrement in potency at higher volumes suggests the alkylbenzene site has a volume of \xe2\x89\xa5 800 \xc3\x853. Within this, a relatively static (with respect to ligand) H-bond donor contributes to initial binding with little involvement in generation of coupling energy. The influence of \xcf\x80 electrons/ring planarity and alkyl adducts on efficacy reveals these aspects of the ligand present towards a face of the channel that undergoes structural changes during opening. The site\xe2\x80\x99s characteristics suggest it is \xe2\x80\x9cdruggable\xe2\x80\x9d; introduction of other adducts on the ring may generate higher potency inverse agonists.
Introduction<!>Molecular Biology<!>Electrophysiology<!>Paradigms and analysis<!>Modeling of alkylphenol and alkylcyclohexanol coupling to channel\ngating<!>Physicochemical parameter estimates<!>Data and statistical analysis<!>Phenols:<!>Isocyanates:<!>Thiols:<!>Fluoro:<!>Cyclohexanols:<!>H-bond interactions with position 1 confer potency<!>Compensating for partitioning reveals the 1-position adduct influences\npotency through action as an H-bond acceptor<!>Are the effects of alterations in position 1 chemistry energetically\nconsistent with disruption of an H-bond?<!>Do differences in pKa influence the H-bonding profile of the phenyl and\ncyclohexanol derivatives?<!>Does the H-bond contribute to coupling energy?<!>The alkylbenzene site is large<!>Alkylbenzenes act via four, radially-arrayed,\nstructurally-defined, sites<!>What is the basis for degradation of coupling energy upon conversion of a\nphenol to a cyclohexanol?<!>Extended alkyl chains disrupt ligand association with HCN1<!>Discussion<!>Reasonableness of the common site, S=4, gating parameters<!>How does ring planarity and/or the \xcf\x80 electron cloud contribute to\ncoupling energy?<!>Do the chemical characteristics identified here correspond to known\nalkylbenzene binding sites?<!>Mapping the isoform divergence onto the HCN1 cryo-EM structure<!>Alkylphenols as novel antihyperalgesics and addiction potential
<p>Neuropathic pain is pain caused by a lesion of, or damage to, the
somatosensory nervous system [1, 2]. Neuropathic pain is a pathologic feature of
numerous conditions, including postherpetic neuralgia, trigeminal neuralgia, sensory
(painful) radiculopathy, painful diabetic neuropathy, peripheral nerve injury,
stroke, and treatment with antineoplastics [2–4]. Globally, it is
estimated that 7– 10% of the population lives with chronic pain with
neuropathic features [2–6]. Cardinal symptoms of neuropathic pain are spontaneous
pain, abnormal response to non-painful (allodynia) or painful (hyperalgesia)
stimuli, dysesthesia (unpleasant/strange sensation – "tingling"
"pins and needles"), and enhanced thermal sensitivity; numerous
pharmacologic approaches to relieving neuropathic pain have been proposed [3], but their efficacy is limited, resulting in
a large unmet need with respect to providing meaningful pain relief [7].</p><p>Recent work suggests that neuropathic pain might better be understood at the
mechanistic level as a function of sensory profiling rather than disease etiology
[8]. Hyperexcitability of primary afferent
neurons is thought to be an early causal factor leading to the development and
maintenance of peripheral neuropathic pain [1,
9]. Numerous ion channel families
contribute to this hyperexcitability, including the hyperpolarization-activated,
cyclic nucleotide-gated (HCN) channel family [10]. In animal models of neuropathic pain, non-selective HCN block with
either ZD7288 [11–14] or the clinically-available drug ivabradine [15–17] results in significant anti-hyperalgesia. HCN1 expression is
upregulated in multiple animal models of neuropathic pain including those wherein
the initiating injury was mechanical damage [11–13], chemotherapy
administration [16–19], or diabetes [20]. Anti-hyperalgesic activity is retained in HCN1-selective molecules
[21–24], suggesting that HCN1-selective blockade has
therapeutic potential.</p><p>Previously we demonstrated that the intravenous general anesthetic
2,6-di-iso-propyl phenol (propofol, DIP-P) and its closely
related, non-anesthetic congener 2,6-di-tert-butyl phenol (DTB-P),
were HCN1-selective inhibitors of channel gating [22, 25]. Such inhibition resulted
from ligand-mediated stabilization of channel closed state(s), which was solely
dependent upon interactions with the membrane embedded channel core [25]. Both DIP-P and DTB-P provided mechanical and thermal
anti-hyperalgesia in mouse peripheral nerve injury neuropathic pain models [22, 23].</p><p>In silico modelling and crystallography demonstrate
alkylphenols dock in cavities in GABAA receptors (GABAA-Rs)
[26, 27] where they likely displace either native acyl chains [28] or water [29–31]. Crystallography
further reveals acyl chains associate with, and intercalate between, the voltage
sensor and pore domains of Kv channels, indicating specific packing at
the lipid-channel interface is thermodynamically favored [32]. Given the structural homology between HCN1 and
Kv channels [10, 33], we posit that specific acyl (and water)
filled cavities are also present on HCN channels. Consequently, the HCN1-alkylphenol
association is likely similar in nature, but likely unique in detail, to the
alkylphenol interaction with GABAA-Rs, and that the alkylphenol:HCN1
association can be defined and optimized.</p><p>In the present study, we demonstrate that H-bonding, ring planarity and/or
π electron interactions and coupling to ring adducts at the 2 and 6 positions
all contribute to HCN1 inhibition at a site with a volume ≥800
Å3. Such characteristics are consistent with alkylphenols
acting at a defined site within the membrane-embedded core of HCN1 channels.
Examination of HCN sequences in conjunction with the cryo-EM structure of HCN1,
shows multiple sites of divergence within this region. Together, the data suggest
that alkylphenol-mediated inhibition of HCN1 channel opening occurs
via a defined and "druggable" site which, if
correct, may open the way to generation of a novel class of high-potency,
non-opioid, therapeutics for neuropathic pain.</p><!><p>Methods for heterologous expression were as previously described [22]. Briefly, cDNA encoding murine HCN1
channels were sub-cloned into a pGH19 vector and amplified in STBL2 cells
(Invitrogen Corporation, Grand Island, NY, USA). cRNA was transcribed from NheI
linearized DNA using T7 RNA polymerase (Message Machine; Ambion, Houston, TX,
USA).</p><!><p>Recordings were made from Xenopus oocytes (Ecocyte,
Austin, TX) 1–3 days after cRNA injection (1–50 ng/oocyte). Cells
were maintained in L-15 media without Ficoll (Specialty Media, Phillipsburg, NJ)
at 16 °C until use.</p><p>Two-electrode voltage clamp (TEVC) data were acquired using a Warner
Instruments (Hamden, CT) OC-725C amplifier, filtered at 1 kHz (Frequency Devices
Bessel model 902, Ottawa, IL) digitized at 2 kHz by an ITC-18 interface
(Instrutech Corporation, Port Washington, NY) and recorded with Pulse software
(HEKA Elektronik, Lambrecht/Pfalz, Germany).</p><p>Microelectrodes were fabricated from 1B120-F4 borosilicate glass (World
Precision Instruments, Sarasota, FL) and had resistances of 0.1 – 0.5
MΩ (I passing) and 1 – 4 MΩ (V sensing) when filled with 3
M KCl. An active virtual ground was used to clamp the bath. Ag-AgCl ground
wire(s) were connected to the bath solution by 3 M KCl-2% agar salt bridges
placed downstream of, but close to, the oocyte. Oocytes were bathed in a
recording solution of (in mM): 107 NaCl, 5 KCl, 1 CaCl2, 1
MgCl2, 10 HEPES-free acid pH 7.4 (NaOH). Recordings were obtained
at room temperature. In all cases, voltages are reported as the command
potentials. The observed potential was within 1% of the command.</p><!><p>The holding potential was −30 mV. Tail currents were recorded at
0 mV. Steady-state activation properties were determined from tail current
activation curves (see Riegelhaupt et al. for a discussion on
the criteria adopted to eliminate collection and analysis bias - [34]). Fits of these relations with the
Boltzmann function (equation 1
where A1 is the current offset and A2 the maximal
amplitude and V the step voltage) yielded the V1/2 (the activation
mid-point) and the slope factor (equal to RT/ZF where R, T and F have their
usual meaning and Z is an approximation of the gating charge associated with
activation).</p><p>Gating of HCN channels is slowed in the presence of in the presence of
inverse agonists. To ensure the determined Boltzmann parameters reflect
thermodynamic behavior free of kinetic contamination, we routinely determined
the gating parameters using step durations of 3 s and 5 s. In no case was there
a systematic difference in fit parameters. This observation is consistent with
activation being equilibrated at all voltages under all conditions tested. The
two determinations were averaged and treated as a single measurement. Data
reported as shifts in the V1/2 (ΔV1/2) were
determined with respect to vehicle control recordings performed before and after
the experimental determinations. Standard errors around vehicle populations were
less than 1 mV.</p><p>Concentration response data were fit with the Hill equation (equation 2), where
ΔV1/2 is as defined above, ΔV1/2 max the
maximal change as determined from the fit, IC50 the concentration of
ligand required for half maximal inhibition of gating and h the Hill
coefficient. In all fits, ΔV1/2 in the absence of ligand was
constrained to zero. For several ligands, estimates of IC50 were
determined assuming ΔV1/2 max and h were equal to those
obtained for DTB-P (see thin green lines on Figure
3 and discussion thereof).</p><!><p>To consider the mechanistic basis by which chemical modification(s) of
the alkylphenol structure perturbs coupling of the ligands to HCN1, we examined
models of the form shown schematically in Figure
1.</p><p>Here, CR, CA and O represent the closed-resting,
closed-activated and open states, of the channel. L is the voltage independent
equilibrium constant that describes opening of activated channels. Activation of
voltage sensors is described by the voltage-dependent equilibrium constant
KV (equation 3)
where KV0 is KV in the absence of an applied
field.</p><p>I and H are full and partial inverse agonists, respectively. Consistent
with previous work wherein we showed DIP-P was indifferent to the activation
status of the voltage sensors [25], here
we assume neither I nor H alter KV. Which is to say both I and H are
considered to bind equally well (or poorly) to CR and CA.
I binds tightly to CR and CA (as defined by the
association constant KCI) but poorly to O (as defined by
the association constant KOI) and thus acts as a full
inverse agonist of opening. For the partial inverse agonist, H, the loss of
efficacy must arise from proportionally weaker binding to C than to O when
compared to binding of I. While it is critical to understand how such partial
behavior arises, particularly when considering rational drug design, it is not
possible to a priori intuit whether it arises due to a loss of
function (a proportionally greater decrement in KC versus
KO) or a gain of function (a proportionally greater improvement
in KO versus KC).</p><p>Within this overall scheme, we examined two limiting conditions.
Specifically, we assumed I and H bind to common site(s) or to distinct site(s).
Here, the definition of common site(s) includes pocket(s) with over-lapping
structures, occupancy of which are mutually-exclusive. In contrast, distinct
sites are, de facto, non-overlapping. In the common site model,
the sum of p and q cannot exceed S (where p and q represent the number of
binding events of I and H, respectively, and S is the number of sites
per channel) and the equation determining the change in the
V1/2 (ΔV1/2) due to the presence of I and/or H
is given by equation 4. In the
distinct site model, p and q can both equal S and ΔV1/2 is
given by equation 5.</p><p>In both equations, A = [H]*KOH, B =
[I]*KOI, C = [H]*KCH and D =
[I]*KCI. Note that in the presence of only H or I (and
the absence of both) equations 4
and 5 become equivalent. It is
the behavior of gating in the combined presence of H and I that distinguishes
between these two mechanisms.</p><p>To allow us to simultaneously fit the behavior of
iso-propyl and tert-butyl analogues, we
defined two sets of equilibrium constants wherein KOH,
KOI, KCH and
KCI describe tert-butyl association
and QOH, QOI,
QCH and QCI describe
iso-propyl association. Versions of equations 4 and 5 describing individual ligand effects and
additivity were generated by substitution of the appropriate equilibrium
constants (and corresponding ligand concentration terms). In simultaneous fits
to the iso-propyl and tert-butyl data, all
eight binding constants and L were free parameters. Importantly, equations 4 and 5 show that the degrees of freedom in the
common and distinct site models are identical and that this is independent of S.
As such, measures of the goodness of fit are directly comparable. As is evident
from inspection of equations 4
and 5, in models where inhibitors
do not discriminate between CR and CA, KV does
not contribute to their influence on ΔV1/2.</p><p>We applied only three constraints. First, S (the number of sites
per channel), was held to be either 1 or 4. While
arbitrary, this is consistent with the tetrameric nature of HCN channels. An S
of 1 is consistent with the site being at the four-fold axis of symmetry, that
is, at a point of juxtaposition of all four subunits. An S of 4 is consistent
with the sites lying within or between channel subunits and distant (with
respect to ligand size) from the central axis. Second, the gating valence Z was
held equal to 2.82. This value was determined from fits of the Boltzmann
function to control GV relations. While not arbitrary, it is well understood
that, in a complex system such as exists for HCN gating, the slope of such a fit
is a secondary, and imprecise, measure that will likely underestimate the total
charge moved [35, 36]. However, given that our three-state gating
scheme is a clear simplification in and of itself ([37] and references therein) and in the model Z serves
only as a proportionality term (equations 4 and 5),
an error in the estimation of Z will have no impact on the qualitative integrity
of our analysis (assuming Z is constant). Is an assumption of constancy of Z
reasonable? At high concentrations, DIP-P and DTB-P both decrease the steepness
of the gating relation (i.e., the Boltzmann slope term gets
larger, meaning the estimated value of Z would get smaller – not shown).
Would use of experimentally determined values of Z under each condition be
better? We think not; while a change in the GV slope could imply Z has changed,
it is highly improbable that the fundamental motions of the sensors (and of the
net charge moved) is radically altered by the presence of an alkylbenzene. A
more reasonable interpretation of changes in the activation slope is that this
reflects altered co-operativity between subunits, an effect the Boltzmann
equation does not account for. For these reasons, we consider use of a stable
measure of charge moved to be the conservative assumption. Third, and finally,
values of all equilibrium constants were constrained to be >0, a
physically reasonable expectation.</p><p>To examine the secondary butyl analogue of DTB-P
(2,6-di-sec-butyl phenol; DSB-P) we defined a third set of
equilibrium constants, JO and JC. As responses to the
DSB-P were only observed at high concentrations and were, therefore, relatively
poorly determined, these data were not included in the simultaneous fit of
iso-propyl and tert-butyl responses as
per
Figure 5. For the DSB-P fit, all parameters
other than JO and JC were held equal to the values used
and/or determined for and from the fits shown in Figure 5 and reported in Table
4.</p><p>Association constants determined from the fits of the models were
transformed to free energy terms according to equation 6.</p><p>It is important to note that within both classes of models the maximal
ΔV1/2 is limited to RT/ZF * Ln 1/1+L (equations 4 and 5) and, ipso facto, the
maximal observed response of the most effective inverse agonist will strongly
influence the estimate of L. This arises because the inverse agonists are
stabilizing closing and, therefore, can only register an inhibitory voltage
shift to the extent permitted by opposing the effect of a favorable opening
transition (large L) that normally drags the activation transition forward. What
does this mean with respect to the behavior expected within competition
experiments? Intuitively, for a channel behaving according to the common site
model, the main phenotype will be for the partial inverse agonist to have no
effect in the face of an intermediate concentration of the full inverse agonist
(or suppress that if high enough concentrations of the partial inverse agonist
can be achieved and its maximal response is lower than that of the included
concentration of full inverse agonist). In contrast, while the actions of
partial and full inverse agonists acting via distinct sites will be (as
expected) energetically additive, this will only be revealed as an additive
effect on the ΔV1/2 at sub-maximal concentrations of the full
inverse agonist. Together, these considerations reveal examination of ligands of
unknown properties need to be considered in the face of a low concentration of
DTB-P. We selected 3 Î¼M.</p><p>Theoretically, an independent measure of L can be determined from the
maximal open probability (PMAX). While conceptually straightforward,
determining PMAX of HCN1 channels is difficult. First, the single
channel conductance of HCN channels is very low, making direct analysis of
single channel records problematic. Second, wild-type HCN1 channels cluster and
gating is slow making noise analysis similarly challenging. We have previously
shown that deletion of the N and C termini of HCN1 generates HCN1 channels that
are amenable to non-stationary fluctuation analysis (NSFA; [25]). When determined in excised inside-out patches
in high K+ recording conditions (to maximize the current amplitude),
the PMAX of HCN1-ΔNvΔC is 0.88, which corresponds to a
value of L of 7.3 (equation
7).</p><p>However, it should be noted that analysis of HCN1-ΔNvΔC in
the excised patch configuration as used for NSFA will underestimate the likely
value of L of wtHCN1 as the effect of at least three factors that stabilize
gating (internal cAMP, internal 4,5-PIP2, and a physiological level
of external Na+ and K+) are lost. This disruption is
reflected in a 40 mV hyperpolarization of gating of HCN1 upon patch excision. If
these positive regulators all acted via enhancing L
(stabilizing opening), equation 7
reveals L would be 93-fold higher in intact cells than reported by NSFA in
excised patches. Further, it is clear that while the effect of
4,5-PIP2 seems to involve, at least in part, a surface charge
contribution, the positive effects of cAMP (5–7 mV right-shift in the
V1/2 for HCN1) and physiological external Na+ and
K+ concentrations (10–12 mV) do indeed arise from a
stabilization of opening [38–41] (and our unpublished observations).
Thus, the model determined value of L of 125 is completely consistent with
independent experimental evidence wherein endogenous positive modulation of L
accounts for 25 mV of the observed 40 mV difference between recording modes.</p><p>Equation 8 determines the
value of L in an intact cell under TEVC (LT) as a function of the
value of L determined from NSFA in excised inside out patches (LI)
and V(T-I), the difference in V1/2 between these modes.
Equation 8 assumes
KV0 is unaltered by patch excision.</p><!><p>We found determinations of ligand physicochemical parameters used in
this study to be sensitive to the analysis engine used (not shown but compare
values in Tables 1 and 2 to those of Krasowski et al. and Woll et al.
[42, 43]). To provide a unified set of values, we redetermined chemical
properties using a single engine, the Maestro 2018–2 package from
Schrödinger. LigPrep was used to generate all possible states at a target
pH of 7.4 ± 0.2. QikProp was used to calculate the reported properties of
all generated states. Properties are reported in Tables 1 (phenyl derivatives) and 2 (cyclohexanol derivatives). LogP is log of the calculated
octanol/water partition coefficient. Values for all ligands in the matrix are
provided to facilitate cross-study comparisons.</p><!><p>Data analysis was performed in PulseFit (HEKA Elektronik) or using
custom analysis routines written in IgorPro (Wavemetrics Corporation, Lake
Oswego, OR). Data were typically collected and analyzed independently by two or
more experimenters with some recordings randomly selected for independent
reanalysis. Data are presented as mean ± SEM with the numbers of
observations per data set reported in figure legends. Errors around Hill fit
parameters and the equilibrium constants determined from fits of the gating
model are non-linear regression approximations of the standard deviation.</p><p>Fits of the Hill equation were to the individual data. To minimize the
effect of different population sizes on the global minimization, fits of the
models were to mean behavior. As all models have the same degrees of freedom and
were fit to a common data set, the goodness of fits are directly comparable and
allow for robust model discrimination. A two-tailed t-test was used to compare
two populations. Comparisons between multiple populations and a control were by
one-way ANOVA using Dunnett's test for multiple comparisons. Statistical
tests were performed in SigmaStat 4.0 (Systat Software).</p><!><p>2,6-di-methyl phenol (DM-P; Sigma-Aldrich Cat. # D175005),
2,6-di-ethyl phenol (DE-P; Sigma-Aldrich Cat. # CBR02039),
2,6-di-iso-propyl phenol (DIP-P; Sigma-Aldrich Cat. #
D126608), 2,6-di-tert-butyl phenol (DTB-P; Sigma-Aldrich
Cat. # D48400), 2,6-di-sec-butyl phenol (DSB-P; Acros
Organics, Fairlawn, NJ, USA Cat. #AC15283).</p><!><p>2,6-di-methyl phenyl-isocyanate (DM-PIC; Sigma-Aldrich, Cat. #
252336), 2,6-diethyl phenyl-isocyanate (DE-PIC; Alfa Aesar, Ward Hill, MA,
USA Cat. # L11339), 2,6-di-isopropyl phenyl-isocyanate
(DIP-PIC; Sigma-Aldrich Cat. # 376930).</p><!><p>2,6-di-methyl phenyl-thiol (DM-PT; Sigma-Aldrich Cat. # 306940);
2,6-di-iso-propyl phenyl-thiol (DIP-PT) was synthesized
within the Milstein Chemistry Core Facility at Weill Cornell Medicine. LCMS,
1H-NMR, and 13C-NMR analysis conformed to
literature precedent [44] and
confirmed purity at >99%.</p><!><p>2-Fluoro-1,3-di-iso-propylbenzene was synthesized
as previously described [43]. For
clarity, here we use a nomenclature that accords with the other molecules
studied, that is, 2,6-di-iso-propyl phenyl-fluorine
(DIP-PF).</p><!><p>2,6-di-methyl cyclohexanol (DM-CH) was obtained from Acros Organics
(Thermo Fisher Scientific, New Jersey, USA; Cat. # 149810050); 2,6-di-ethyl
cyclohexanol (DE-CH), 2,6-di-iso-propyl cyclohexanol
(DIP-CH) and 2,6-di-sec-butyl cyclohexanol (DSBCH) were as
previously described [45].
2,6-di-tert-butyl cycloxhexanol (DTB-CH) was
synthesized within the Milstein Chemistry Core Facility at Weill Cornell
Medicine. LCMS, 1H-NMR, and 13C-NMR analysis conformed
to literature precedent [45] and
confirmed purity at >99%.</p><p>All ligands were prepared as stock solutions in DMSO then dispersed
in the bath solution with vigorous stirring for at least 60 minutes. In the
absence of any other excipient, the final DMSO concentration was 0.2%
(V/V). 2-Hydroxypropyl-β-cyclodextrin
(HPÎ²CD; Sigma-Aldrich, St. Louis, MO;H107) can increase the
solubility of ligands such as DTB-P and DIP-P by ≥200-fold and do so
with no effect on pharmacokinetics ([22]; and references therein). In an attempt to extend the range
of the concentration responses we could examine, in some experiments we
included HPÎ²CD. While the presence of HPÎ²CD did not
qualitatively alter the behavior of any ligand, its presence did cause a
modest right-shift in the inverse agonist concentration response curves when
it was included. However, as the right-shift was not demonstrably outside
the normal scatter observed in the data between recordings from different
donor frogs on different days and was far smaller than the variance in the
model fit parameters, the different vehicle conditions were combined.</p><!><p>Figure 2 shows representative
families of voltage-clamp recordings (Figure
2A) in either vehicle or vehicle plus 10 Î¼M of one of four
2,6-di-iso-propyl phenyl derivatives (Figure 2B, also Figure
3). Inspection of the records suggests that systematic perturbation
of the H-bonding capability of the adduct at the 1-position leads to a
disruption of inhibition of HCN1 gating. This inference is confirmed by
inspection of the activation curves (Figure
2C).</p><p>Diminished inhibition could be due to reduced efficacy, a weakening of
potency, or both. To address this, we constructed complete
concentration-response relations for the four ligands (Figure 3 central column, first four rows). Inspection
confirms there is a loss of potency upon 1-position substitution. The
progressive loss of potency (DIP-P > DIP-PT > DIP-PIC
>>> DIP-PF) evident in Figure
3 and 4A (symbols within the
dashed ellipse – see also Table 3)
does not track the ability of the ligands to act as either a H-bond donor (DIP-P
> DIP-PT >>> DIP-PIC ~ DIPPF) or acceptor
(DIP-PIC > DIP-P > DIP-PT >>> DIP-PF) (Table 1).</p><p>It should be noted, however, that changing the 1-position adduct will
alter not just the H-bonding profile of the ligand but other aspects of its
physical chemistry. Thus, the four ligands here will have modestly different
volumes, pKa's and markedly divergent hydrophobicities (Table 1), and each of these properties is examined
below.</p><!><p>Due to partitioning, at an equivalent aqueous concentration, more
hydrophobic molecules will have a higher intra-plasma membrane concentration. As
alkylphenols act by coupling to the membrane embedded core of HCN1 channels
[25], we next asked whether
partitioning per se alters either the rank order and/or
energetic strength of 1-position adduct substitutions.</p><p>To address this, we assumed the within-membrane concentration of each
ligand at its aqueous IC50 can be represented by the aqueous
IC50 multiplied by the calculated partition coefficient (P). The
dashed ellipse-encircled symbols in Figure
4B plot the partition-corrected IC50 values for the four
ligands considered in Figure 2 (and
DIP-CH). The progressive loss of potency (DIP-PIC ~ DIP-P
>> DIP-PT >>> DIP-PF) qualitatively tracks
closer to the ability of the ligand to act as an H-bond acceptor (DIP-PIC
> DIP-P > DIP-PT >>> DIP-PF) than a donor
(DIP-P > DIP-PT >>> DIP-PIC ~ DIP-PF) with
the most glaring potency-acceptor inconsistency being the behavior of isocyanate
compared to hydroxyl. While isocyanate can theoretically accept 3 H-bonds but
hydroxyl only one, DIP-PIC and DIP-P exhibit essentially identical potency. We
posit that the similarity of DIP-PIC and DIP-P is consistent with the 1-adduct
acting as an H-bond acceptor in an environment where there is only a single
H-bond donor available. Such a conclusion is inconsistent with ligand coupling
to water and suggests the bond is a ligand-protein interaction.</p><!><p>Replacement of the 1-OH by F eliminates the capacity of this adduct to
engage in H-bonding - either as a donor or an acceptor. Importantly, it does so
with little effect on most other physicochemical properties of the molecule
[43, 46]. Consequently, we can discretely ascribe alterations in the
IC50 to disruption of H-bonding. Based on the data in Figures 3 and 4, we estimate that the shift in IC50 in response to
replacing OH with F is between 46 to 1,870-fold as determined from the ratios of
the observed IC50s for DIP-P and DIP-PF (Figure 4A) and the partition-corrected
IC50s for those ligands (Figure
4B), respectively. These changes correspond to a 2.3 to 4.5 kCal
mol−1 destabilization of association; energetically, the
observed loss of potency is entirely consistent with the disruption of an
H-bond.</p><!><p>As pKa's of the phenol and cyclohexanol derivatives are orders of
magnitude larger than the experimental pH, the concentrations of ionized species
will be very low. Differential presentation of the hydroxyls is unlikely to
contribute to the observed behavior. While the pKa of sulphydryls is in the
experimental range, as the site of action of the ligands appears to be in the
largely anhydrous membranous space, differential ionization is unlikely to be a
factor beyond influencing the partitioning coefficient – an effect
already accounted for in the data in Tables
1 and 2 and Figure 4B.</p><!><p>At the highest concentrations, DIP-P (and DTB-P) elicit an ~
−40 mV shift in the V1/2 of HCN1 gating. While we were unable
to reach concentrations sufficient to obviously saturate the concentration
response curves for DIP-PT, DIP-PIC or DIP-PF, the first two appear to trend
towards an efficacy similar to that of DIP-P (and DTB-P). In contrast, the
concentration response relation of DIP-PF appears to be flatter than the
hydroxyl, thiol or isocyanate iso-propyl analogues, and it
appears it may tend to a plateau less than −40 mV; this suggests the
interaction energy at the 1-position does not absolutely partition towards
initial binding but, rather, the 1-position contributes modestly to coupling
energy.</p><!><p>To examine the role of ligand volume, we systematically and
symmetrically varied the nature of the alkyl adducts on positions 2 and 6 and
did so in the background of not only a hydroxyl on position 1 (top row) but also
in the background of both a thiol (second row) and an isocyanate (third
row).</p><p>Inspection of the hydroxyl-containing substituents appears to show a
U-shaped dependence of potency on the alkyl identity (Figure 3). Iso-propyl and
tert-butyl derivatives are comparable with respect to
potency but inclusion of either smaller (ethyl) or more extended
(sec-butyl) side-chains results in a marked right-shift in
the concentration-response curves. A plot of IC50 as a function of
molecular volume supports this impression (Figure
4A). Although we do not have access to the complete alkyl adduct
series for 1-position thiol or isocyanate families, these partial series exhibit
a comparable trend to higher potency as the alkyl group tends towards the
iso-propyl : tert-butyl apparent optimum
(Figures 3, 4A).</p><p>These data suggest that there is a narrowly-defined pocket that
alkylbenzene derivatives dock within. If the alkyl groups are too small the
ligand does not orient correctly while if the groups get above a threshold, the
ability of the ligands to enter the pocket is diminished. Such a conclusion
would be consistent with the well described "cut-off" effect of
anesthetics at other molecular targets [47–50] with the caveat
that the changes considered here (increasing the number of carbons in the 2 and
6 alkyl adducts) increases not only ligand volume, they increase ligand
hydrophobicity and, as such, increases the ligand's partition
coefficient.</p><p>Figure 4B plots the
partition-corrected IC50 values of the matrix of ligands as a
function of their molecular volume. When we consider only those ligands where a
clear inflection and rising phase is discernable in the concentration response
relation (disregarding DM-P, DM-PIC, DM-CH, DE-CH, and DSB-CH) the dependence on
volume below the apparent iso-propyl :
tert-butyl optimum largely disappears.</p><p>There is one anomaly within this analysis. DM-PT exhibits a clear
inverse agonist activity whereas DM-P and DM-PIC do not. Given that both
hydroxyl and isocyanate are better H-bond acceptors than is a thiol, and
hydroxyl a better donor (Table 1), this
preference for thiol in the di-methyl family is unexpected. As DM-PT is more
hydrophobic (logP 2.87) than either DM-P (logP 2.22) or DM-PIC (logP 1.49; Table 1), it is possible that the apparent
discontinuity reflects an edge effect wherein DM-PT reaches an effective
within-membrane concentration whereas what would otherwise be effective
molecules (DM-P and DM-PIC) do not. However, this is unlikely to be the sole
determinant. Thus, the observations that DE-PIC is effective despite a logP of
1.93 would argue against inadequate ligand partitioning into the membrane being
the sole reason DM-P and DM-PIC are ineffective in comparison with DM-PT. The
basis for the additional advantage afforded to DM-PT (or disadvantage of the
hydroxyl and isocyanate dimethyl derivatives) is unclear.</p><p>What about the apparent suppression of effectiveness at alkyl sizes
above that of tert-butyl? The response curve for DSB-P suggests
it could be either a low potency and high efficacy full inverse agonist (the
data in Figure 3 represent the foot of a
curve that has a right-shifted IC50 of ~355 Î¼M as shown
by the thin green line) or a partial inverse agonist that has a higher potency
and lower efficacy (the data in Figure 3
represent the maximal ΔV1/2 with an IC50 of
~30 Î¼M). Given the low aqueous solubility of butyl phenols
(~200 Î¼M; [51]) we were
unable to reliably address this question by simply increasing the range of the
concentration response curve.</p><p>Importantly, as an IC50 of either 30 Î¼M or 355
Î¼M is markedly larger than that of DTB-P (IC50 = 4.8
Î¼M) the upturn seen at larger volumes in Figure 4 is likely to be real. This conclusion receives further
support from the additivity/competition approach adopted below, that suggests
DSB-P is indeed a low potency full inverse agonist. Together, these data are
consistent with the notion that, while large (≥ 800
Å3), the inverse agonist alkylphenol site has a defined volume
and geometry.</p><!><p>The observation that various congeners of DIP-P and DTB-P may exhibit
partial inverse agonist behavior has important implications with respect to
understanding the molecular interactions between this class of molecules and
HCN1 and, hence, the development of a SAR profile for the presumptive inverse
agonist site(s). Accordingly, we next examined whether the behavior of
corresponding cyclohexanols and alkyl adduct-modified ligands could further
inform us about the nature of the alkylbenzene:HCN1 channel interaction.
Specifically, we sought to determine the answers to four fundamental questions:
1) Are the cyclohexanols and the short and long alkyl group phenols truly
partial inverse agonists? 2) Does the behavior of cyclohexanols and phenols
further validate the notion of a structurally-defined alkylbenzene site on HCN1
as opposed to a generalized perturbation effect? 3) Can the data inform us as to
the stoichiometry and location of the presumptive alkylbenzene site(s)? 4) Can
we elucidate the mechanistic basis for the loss of efficacy in the
cyclohexanols?</p><p>The simplest ligand to consider is DE-P. The observed
ÎV1/2 appears to a saturate at 100 Î¼M. Indeed, in
paired experiments we find the ΔV1/2 in the face of 100 and
300 μM to be indistinguishable (−21.6 mV ± 4.2, n = 8 and
−21.2 mV ± 4.6, n = 9; respectively; P = 0.95, two-tailed t-test).
Importantly, the aqueous solubility limit of DE-P (~10 mM; https://comptox.epa.gov/dashboard/dsstoxdb/results?search=DTXSID5061401#comments)
is two-orders of magnitude higher than the functional saturation observed here.
That is, DE-P is indeed a partial inverse agonist with a maximal efficacy
~2-fold lower than the maximal effect seen with DIP-P and DTB-P (see also
Table 3). These data show the 2
and/or 6 adducts can "sense" changes in channel structure upon
channel opening and, in this case, the effect of the smaller alkyl adduct is to
reduce coupling energy.</p><p>With respect to the cyclohexanols, the maximal ΔV1/2
determined from fits of the Hill equation suggest there to be a >2-fold
decrease in efficacy in the di-iso-propyl background and a
>4-fold decrease in the tert-butyl background (Figure 3 and Table 3). This decrement is associated with little
(tert-butyl) or only modest (iso-propyl)
decrement in potency (see the green dashed lines in Figure 3). However, the low aqueous solubility of
these cyclohexanols raises the concern that the apparent saturation of their
concentration response curves is a function of ligand availability and not
channel responsiveness. To address this, we presented HCN1 channels with a
submaximal concentration of DTB-P in the absence and presence of increasing
concentrations of DIP-CH and DTB-CH (as well as DIP-P, DSB-P and DE-CH) and
considered the results within the context of the gating models outlined in Figure 1.</p><p>The symbols in Figure
5A–B reproduce the
observed concentration response data (from Figure
3) for DIP-CH and DIP-P (Figure
5A) and DTB-CH and DTB-P (Figure
5B). Figure 5C–E present the shift in the V1/2
elicited by increasing concentrations of DIP-CH (Figure 5C), DTB-CH (Figure 5D)
and DIP-P (Figure 5E) in the presence of 3
Î¼M DTB-P. The solid blue lines in Figure
5A–E represent the
simultaneous fit of the common site model at S=4.</p><p>Qualitatively, the behaviors of DIP-CH and DTB-CH are exactly those
expected for partial inverse agonists while DIP-P behaves exactly as expected
for a second full inverse when all act at the same site(s) as DTB-P (Methods).
Quantitatively, the optimized values of the ligand association constants are
physically reasonable and the errors around each term are relatively modest
(Table 4; Figure 5F). In contrast, the fit of the common site
model at S=1 is inferior (χ2 is 147 vs. 91 for S=4) and L
adopts a physically unreasonable value of 7 * 108 ± 5 *
108.</p><p>We examined the reasonableness of the distinct-site models in two ways.
First, we used the parameters reported in Table
4 and Figure 5F to predict the
behavior of the distinct-site model at S=4. As anticipated (Methods) models of
this class are, by definition, additive in the domain of inverse agonist
concentrations where the ΔV1/2 is submaximal; the dashed blue
lines in Figure 5C–E show poor or no correspondence to the observed data.
Second, we fit the distinct-site model to the data. The fits were inferior to
those of the single site model (χ2 at S=4 was 263 and 809 at
S=1). Furthermore, at both S=1 and S=4, one or more ligand association constants
dropped to zero while, at S=1, L was unreasonably large and very poorly defined
(4.1 * 105 ± 9.3 * 108). It is important to note
that as the distinct-site model is conceptually identical to a generalized
membrane perturbation model, failure of the distinct model provides additional
evidence for the existence of specific, structurally-defined, alkylbenzene
site(s) on HCN1.</p><p>Consistent with the above findings, DE-CH exhibited neither additivity
nor competition with 3 Î¼M DTB-P (data not shown). While DE-CH appears to
be only weakly effective as an inverse agonist (displaying an inflexion in the
concentration response curve only at the highest concentration achieved - Figure 3) the lack of effect of DE-CH in the
presence of DTB-P provides an additional control for potential non-specific
effects of loading the cell membrane with these small molecules.</p><p>The simplest interpretation of these findings is that DTB-P (and DIP-P)
act as full inverse agonists by interacting with four radially-arrayed binding
sites lying within or between each HCN subunit, and that cyclohexanols (and
ligands with altered alkyl adducts) associate with these same sites but generate
less coupling energy than do the iso-propyl and
tert-butyl phenols.</p><!><p>Figure 5F shows this arises from a
preferential degradation in the association of the ligand with the closed state
with a more modest disruption of open state affinity. In the background of
iso-propyl and tert-butyl ligand families,
the effect of ring saturation on closed and open state association is largely
independent of the alkyl chain background (as shown by the comparable shifts in
ΔG - Figure 6F). This is consistent
with these alkyl groups contributing comparably to channel-ligand association
energies.</p><!><p>Figure 6 shows the results of an
additivity experiment directly comparable to that in Figure 5. While the achievable concentration range of
DSB-P is a limiting factor and the fits are, consequently, poorly determined,
the data are consistent with DSB-P being a low affinity full inverse agonist. As
with the data for DIP-CH, DTB-CH and DIP-P, the behavior of DSB-P was best
described by the common-site model S=4.</p><!><p>Four key chemical characteristics define congeners of DIP-P - H-bond
propensity, ring saturation/planarity, hydrophobicity, and molecular volume. We find
that: 1) There is an alkylbenzene site on HCN1 channels that is relatively large,
having a volume ≥ 800 Å3; 2) The site contains an H-bond
donor that interacts with the 1-position adduct on the ligand in a manner that
contributes to initial binding energy; 3) The 1-position adduct is, at best, poorly
involved with generation of coupling energy indicating the site's hydrogen
bond donor is relatively immobile (with respect to ligand) during channel opening;
4) The presence of a π electron cloud is coupled to the generation of
coupling energy suggesting that these electrons (or the imposed ring planarity) is
important in establishing a preferential association between the ligands and the
closed state(s) of the channel; 5) Substituents at the 2 and 6 positions of the
ligand (and possibly others excluding the 1-position) interact with aspects of the
channel that are structurally dynamic during opening; 6) The data are most readily
reconciled with the existence of four radially-arrayed alkylbenzene sites suggesting
the sites lie within each HCN1 subunit or at the inter-subunit interface and these
sites are distant (in terms of ligand geometry) from the central axis of the
channel; 7) Addition of adducts capable of forming more specific interactions than
permitted by alkyl chemistry may better exploit the state dependent channel surface
and generate a higher potency class of inverse agonists.</p><!><p>The best test of a model is to have it predict data not included in the
fit. While we cannot readily gain independent measures of the association
constants, we can independently estimate L. Based on the maximal open
probability (PMAX) we estimate L to be between 7.3 and 679 (Methods).
That our model-determined measure of L (125) falls within this range suggests
the model determined parameters are consistent with the energetics of HCN1
gating and drug effects thereon.</p><p>Does our use of a simple three-state gating scheme undermine the
validity of this analysis? While the model shown in Figure 1 does not reproduce the nuanced kinetic
behavior of HCN channels ([37] and
references therein), the general form of the relationships between L,
PMAX and ΔV1/2 in more complex,
kinetically-faithful, models will be comparable to equations 7 and 8 (albeit such solutions will contain
proportionality terms based on the presence of additional coupled equilibria).
Any quantitative impact on the estimates of L from such an increase in the
complexity of the basal gating model will equivalently impact its determination
from the additivity data in Figure 5 and
the NSFA studies. Accordingly, we posit that the analysis of inverse agonists
and partial inverse agonists predicts a physiologically-reasonable determination
of the fully-activated open-closed equilibrium which will remain true with any
such model expansion.</p><!><p>What is the likely basis for the preferential loss of closed state
binding upon ring saturation? One possibility is that the π electrons
form an electrostatic bond with the channel that is stronger in the closed state
than in the open state. The ~1 kCal mol−1 degradation
in Kc on shifting from the phenolic structure to the cyclohexanol is
energetically consistent with such a scenario [52]. An alternative scenario is that the rigid structure of the
phenolic ring acts as a better "foot-in-the-door" than does the
more flexible cyclohexanol. Distinguishing between these mechanisms could be
aided by use of fluorine for ring-hydrogen substitution analogues [53, 54] in the background of either DIP-P or DTB-P, but those studies
are beyond the scope of the current manuscript.</p><!><p>The existence of spatially-defined cavities (i.e.,
presumptive functional binding sites) that can accommodate small anesthetic
molecules have been identified in a variety of proteins, including firefly
luciferase, human serum albumin, and both ligand- and voltage-gated ion channels
[55–57]. Modeling based on molecular
mechanics–quantum mechanics calculations suggest that anesthetic-protein
complexes share a set of common characteristics, or "binding
motifs"; these motifs are "polar and nonpolar interactions within
an amphiphilic binding cavity, including the presence of weak hydrogen bond
interactions with amino acids and water molecules" [58]. The results presented herein suggest the
existence of a spatially-defined cavity in HCN1 channels that accommodates small
lipophilic molecules and that the underlying interactions conform to the
previously proposed "binding motifs."</p><!><p>HCN1 is the isoform most sensitive to alkylphenols with HCN2, 3 and 4
being progressively less sensitive albeit with some dependence on the alkyl side
chain identity (iso-propyl versus tert-butyl)
[22]. With respect to trying to
identify key residues that determine the potency and efficacy of alkylphenol
inhibition, this profile suggests residues that are unique to HCN1 may not be
the complete picture but that locations that follow a divergence in conservation
may also be important.</p><p>Figure 7 shows within the
transmembrane structure of HCN1-ΔNvΔC, the minimal HCN1 channel
that retains alkylphenol sensitivity, there are residues unique to HCN1 and
others divergent between the four isoforms. While our hypothesis is that
residues contacting, or within, the S5-S6 motif are most likely to contribute to
the site, it is clear that there are too many divergent locations to justify a
site-directed analysis (with all its inherent caveats of interpretation).
Accordingly, identification of the alkylbenzene sites will likely be best
determined using photolabeling and chimeric approaches combined with molecular
docking strategies once the open and closed architectures of the HCN1 channel
are known [55]. The results presented
here will allow for rationale interpretation of the findings of any, and all,
such studies.</p><!><p>Might propofol, and by extension other alkylphenols, interact with other
signaling pathways, especially those centered on the opiate receptor system?
In vitro, propofol has no direct effect on G-protein
coupled inwardly rectifying K+ channels (GIRKs), nor does a
clinically relevant concentration of propofol (2 Î¼M) potentiate
D-Ala2,N-Me-Phe4,Gly5-ol]enkephalin
(DAMGO)-evoked currents generated by GIRK2 channels [59]; importantly, and in contrast, supratherapeutic
propofol concentrations (up to 100 μM propofol, a 50×-clinically
relevant concentration) inhibit DAMGO-evoked currents generated by chimeric mu
opiate (Î¼OR-Gqi5) receptors [60]. Thus, there is no evidence that propofol potentiates mu opiate
receptor (mOR) function at the molecular level [61]. As to other alkylphenols, DTB-P, like propofol, inhibits HCN1
gating [22], but unlike propofol, it has
minimal activity at GABAA receptors [42] although it increases open channel probability (Po)
of α3β glycine receptors [23]; there are no published data on the effects of DTB-P on
G-protein coupled receptors in general, and mORs in particular. Despite
increasing Po of α3β glycine receptors,
α3β glycine receptors do not contribute to either DTB-P-mediated
mechanical or thermal anti-hyperalgesia following chronic constriction nerve
injury [23], an effect comparable to our
observations using a partial sciatic nerve ligation model of neuropathic pain
[22].</p><p>With regards to analgesia and modulation of pain pathways in
vivo, propofol suppresses activity of spinal cord dorsal horn
neurons in response to noxious stimulation in goats [62] and rats [63], and this effect appears to be partly GABAergic in origin [63]. In human subjects, however, a
meta-analysis of thirty-nine clinical trials with a combined subject population
of 4,520 patients indicated that propofol-based anesthetics do not result in
less postoperative pain than those utilizing volatile anesthetics [64], suggesting that propofol per
se does not produce meaningful analgesia that extends into the
immediate postoperative period. Such an observation does not preclude the
development of alkylphenols as anti-hyperalgesics as anti-hyperalgesia is not
the same as analgesia, reflecting the fact that pain -
"an unpleasant sensory and emotional experience associated with actual or
potential tissue damage" is not equivalent to
hyperalgesia – "increased pain from a
stimulus that normally provokes pain" (https://www.iasp-pain.org/terminology?navItemNumber=576). The
in vivo data comport with the in vitro
data and suggest that propofol, and presumably related alkylphenols, should have
a good therapeutic index with respect to the risk of addiction as the relevant
circuitry underlying opioid-dependent addiction [65, 66] and propofol
mechanisms of action [67, 68] do not overlap.</p><p>Our data suggest that a spatially defined cavity that accommodates
alkylphenols exists in the transmembrane or interfacial region of HCN1. Whether
the binding site is similar to comparable binding sites in other channels,
including GABAA [26, 69], TRPA1 [70, 71], and glycine [23], receptors remains to be determined.
Given the size of the pocket and the lack of specific chemical interactions with
alkyl adducts, the findings suggest it may be possible to develop a higher
potency analogue of DTB-P with drug-like properties, particularly as the HCN1
cryo-EM structure is now available [33].
Such experiments have the potential to reveal a new, opioid-independent approach
to treating neuropathic pain.</p>