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Soluble epoxide hydrolase limits mechanical hyperalgesia during inflammation
Christian Brenneis*†, Marco Sisignano†, Ovidiu Coste, Kai Altenrath, Michael J Fischer, Carlo Angioni, Ingrid Fleming, Ralf P Brandes, Peter W Reeh, Clifford J Woolf, Gerd Geisslinger and Klaus Scholich
Corresponding author:
† Equal contributors
Pharmazentrum Frankfurt/ZAFES, Institute of Clinical Pharmacology, Johann Wolfgang Goethe-University, Frankfurt, Germany
F. M. Kirby Neurobiology Center, Department of Neurology, Children's Hospital Boston, Boston, MA, USA
Department of Pharmacology, University of Cambridge, Cambridge, UK
Institute for Vascular Signalling, ZAFES, Faculty of Medicine, Johann Wolfgang Goethe-University, Frankfurt, Germany
Institute for Cardiovascular Physiology, ZAFES, Faculty of Medicine, Johann Wolfgang Goethe-University, Frankfurt, Germany
Department of Physiology and Pathophysiology, Friedrich-Alexander-University Erlangen-Nürnberg, Erlangen, Germany
For all author emails, please .
Molecular Pain 2011, 7:78&
doi:10.69-7-78
The electronic version of this article is the complete one and can be found online at:
Received:14 June 2011
Accepted:4 October 2011
Published:4 October 2011
& 2011 B licensee BioMed Central Ltd.
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Background
Cytochrome-P450 (CYP450) epoxygenases metabolise arachidonic acid (AA) into four different
biologically active epoxyeicosatrienoic acid (EET) regioisomers. Three of the EETs
(i.e., 8,9-, 11,12- and 14,15-EET) are rapidly hydrolysed by the enzyme soluble epoxide
hydrolase (sEH). Here, we investigated the role of sEH in nociceptive processing during
peripheral inflammation.
In dorsal root ganglia (DRG), we found that sEH is expressed in medium and large diameter
neurofilament 200-positive neurons. Isolated DRG-neurons from sEH-/- mice showed higher EET and lower DHET levels. Upon AA stimulation, the largest changes
in EET levels occurred in culture media, indicating both that cell associated EET
concentrations quickly reach saturation and EET-hydrolyzing activity mostly effects
extracellular EET signaling. In vivo, DRGs from sEH-deficient mice exhibited elevated 8,9-, 11,12- and 14,15-EET-levels.
Interestingly, EET levels did not increase at the site of zymosan-induced inflammation.
Cellular imaging experiments revealed direct calcium flux responses to 8,9-EET in
a subpopulation of nociceptors. In addition, 8,9-EET sensitized AITC-induced calcium
increases in DRG neurons and AITC-induced calcitonin gene related peptide (CGRP) release
from sciatic nerve axons, indicating that 8,9-EET sensitizes TRPA1-expressing neurons,
which are known to contribute to mechanical hyperalgesia. Supporting this, sEH-/- mice showed increased nociceptive responses to mechanical stimulation during zymosan-induced
inflammation and 8,9-EET injection reduced mechanical thresholds in naive mice.
Conclusion
Our results show that the sEH can regulate mechanical hyperalgesia during inflammation
by inactivating 8,9-EET, which sensitizes TRPA1-expressing nociceptors. Therefore
we suggest that influencing the CYP450 pathway, which is actually highly considered
to treat cardiovascular diseases, may cause pain side effects.
Keywords: sEH; EET; CYP450; TRPA1; hyperalgesiaBackground
Inflammatory responses after tissue damage or infection cause the release of arachidonic
acid (AA) and its subsequent metabolism to biologically active lipids in activated
immune cells and hyperactive neurons []. Free AA is a major substrate for cyclooxygenases (COX), lipoxygenases (LOX) and
cytochrome P450 (CYP450) epoxygenases which metabolize it to prostanoids, leukotrienes
and epoxyeicosatrienoic acids (EETs) and hydroxyeicosatetranoic acids (HETES), respectively
[-]. At the site of injury and in the CNS prostanoids and leukotrienes are inflammatory
and pain mediators that attract and activate immune cells as well as directly sensitize
nociceptive neurons [,].
Recently, those EETs involved in vascular homeostasis and coronary physiology have
been shown also to influence nociceptive processing [,]. EETs either bind intracellular targets, are released to act as auto- or paracrine
mediators or are stored in cell membranes esterified to phospholipids []. They activate PPARγ or the cAMP/PKA pathway and modulate and/or activate a variety
of channels including several transient receptor potential (TRP) channel isoforms,
large-conductance Ca2+-activated K+ channels (BK(Ca)) and L-type voltage gated calcium channels (Ca(v)) [].
Three EETs (8,9-, 11,12- and 14,15-EET) are metabolized by soluble epoxide hydrolase
(sEH) to their corresponding dehydro metabolites (DHET), which are thought to be less
active []. Increasing EET bioavailability by blocking sEH activity or expression can be used
to study the actions of EETs in vivo. sEH inhibition strongly reduces nociceptive responses, suppresses COX-2 expression
and up-regulates the acute neurosteroid-producing gene StARD1, when applied either
topically to the inflammation site or injected intrathecally in a model of inflammatory
and neuropathic pain [,]. Furthermore, a conditional knockout of cytochrome P450 reductase (CRP) in CNS neurons,
which blocks CYP450 activity, largely abolishes morphine-induced anti-nociception,
suggesting that EETs have an anti-nociceptive activity downstream of the μ-opioid
receptor [].
EETs can however, directly activate TRPV4 channels as well as elicit rapid membrane
insertion of TRPV4 and TRPC6 channels [-] which may be pro-nociceptive. In DRG-neurons, calcium influx after TRP-channel opening
leads to activation of signaling molecules like p38 mitogen-activated protein-kinase,
which sensitize the neurons for further excitation []. TRPV4 and TRPC6, as other TRP- family channels including, TRPV1 and TRPA1, mediate
thermal and mechanical hyperalgesia in various disease models [,]. However, the mechanism how they are activated by specific endogenous agonists is
largely unknown.
We have now investigated EETs in nociceptive processing by phenotyping mice with targeted
deletion of sEH, and find that the EETs metabolized by sEH sensitize TRPA1 expressing
DRG neurons and that sEH-/- mice show an increased mechanical hyperalgesia.
sEH is expressed in primary afferent neurons
To determine if and how sEH could potentially contribute to nociceptive processing
we first analysed the cellular distribution of sEH by Western blotting and immunohistochemistry
in edematous paw and DRG tissue up to 48 h after an intraplantar zymosan injection.
Western blotting of tissue from wild- type mice revealed a 58 kDa band which corresponds
to the predicted size of sEH but was not regulated in paw or DRG tissue (Figure ). The band was specific for sEH, since it was absent in the sEH-/- animals (Figure ). Comparison of tissue sections from wild- type and sEH-/- animals by immunohistochemistry showed specific sEH staining in the DRG (Figure ). Notably, the distribution and localization of sEH immunoreactivity was not altered
in DRGs after zymosan treatment (Figure ). As indicated by co-staining with NeuN, sEH expression in the DRG was restricted
to a subset of neurons. To determine which cell types express sEH in DRGs, we combined
conventional immunohistochemistry with the Multi-epitope ligand cartography-(MELC)-system
which allows repeated immunostaining using FITC-labelled primary antibodies against
several cell type-specific markers [,]. We compared sEH expression with staining for NeuN as a marker for all neurons, GFAP
as a marker for satellite glial cells, IB4 as a marker for unmyelinated small diameter
nociceptive neurons and NF200 as a marker for myelinated medium and large diameter
neurons. The co-localisation pattern indicated that sEH is predominantly expressed
in NF200-positive neurons (Figure ). These medium and large sized neurons have myelinated axons forming Aβ-and Aδ-fibers,
with the latter predominatly involved in fast nociception []. Further, a communication between innocuous Aβ- and noxious C- or Aδ-fibers can play
a role in the development of allodynia [].
Expression pattern of sEH in primary afferent neurons. (A) Representative western blot analysis of sEH-expression during zymosan induced
inflammation in paw and DRG-tissue 0 - 48h after zymosan injection. (B) Western blot
detection of sEH expression in tissues from dorsal root ganglia and paw oedema tissue
of wild type and sEH-deficient mice 48h after zymosan injection. Arrowhead indicates
a signal appearing only in wild- type tissue at approximately 58 kDa. (C) Expression
pattern of sEH in dorsal root ganglia neurons. Immunofluorescence staining of L4 or
L5 dorsal root ganglia cryosections during zymosan induced inflammation (0h, 1h, 6h
and 48h zymosan) was performed with antibodies against sEH (green) and NeuN (red).
At time point 48h post zymosan injection wildtype and sEH-deficient DRG-tissue were
compared. (D) Selective sEH expression in NF200 positive neurons. Combination of conventional
immunofluorescence staining for sEH as in (C) with the MELC technology using 7AAD
and FITC-labeled primary antibodies against NeuN, glial fibrillary acidic protein
(GFAP), isolectin B4 (IB4) and neurofilament 200 (NF200).
sEH activity in isolated DRG neurons and its contribution to EET levels during inflammation
To investigate if sEH contributes to steady state EET levels in vivo, we extracted lipids form DRG tissue of naive wild type and sEH-/- mice. EET quantification by LC-MS/MS analysis revealed that deletion of sEH increased
8,9-EET, 11,12-EET and 14,15-EET tissue concentrations (Figure ) while 5,6-EET levels were unaffected. DHETs were not detected in DRG tissue. To
test if sEH expression is regulated during peripheral inflammation and if differences
in EET levels become more pronounced, we analysed DRG tissue samples 48h after intraplantar
zymosan injection. We found no difference in 5,6-EET, 8,9-EET and 11,12-EET tissue
concentrations (Figure ). However, 14,15-EET concentrations were reduced below the detection limit. We next
investigated the functional contribution of sEH to EET hydrolysis in DRG neurons by
comparing basal EET synthesis and that after stimulation with 0.5 μM AA in DRGs from
wild type and sEH deficient mice. In neuronal cell lysates, we detected only a small
increase in the levels of some EETs, and only 8,9-EET was elevated in sEH-/- compared to sEH+/+ cells (Figure ). However, the level of DHET markedly increased when AA was added, and this was significantly
higher in wild type than in sEH-/- cells (Figure ).
Contribution of sEH to EET release from sensory neurons and DRG tissue levels during
inflammation. (A) Basal EET levels extracted from tissues of DRGs from L4 and L5 segments from
wild type and sEH deficient mice were determined by LC-MS/MS analysis. (B) EET-tissue
levels in DRGs 48h after intraplantar zymosan injection. Data shown represent the
mean ± SEM from 6 animals. (C) EET and DHET (D) levels in lysates of sensory neurons.
Neuron enriched cultures from DRGs of wild- type and sEH-deficient mice were stimulated
with 0.5 μM arachidonic acid and incubated for 2 h until extraction of EETs from cells.
Data shown represent the mean ± SEM from 5 culture dishes. (E) EET and DHET (F) release
from sensory neurons.. EETs were extracted from culture media of cells used in panel
(C). Data shown represent the mean ± SEM from 5 culture dishes. Student's t test: *, p ≤ 0.05; **, p ≤ 0.01.
In contrast, we found dramatically elevated EET levels (around 100-fold) after AA
stimulation in the cell culture medium (Figure ). This indicates that cell associated EET concentrations quickly reach saturation
and that higher AA availability increases EET release more than its levels within
neurons. Interestingly, the extracellular DHET concentrations were strongly induced
upon AA stimulation, an effect which was markedly reduced in sEH-/- neurons (Figure ). These data suggest that after increases in AA availability, EETs are released from
sensory neurons and that a deficiency in cellular hydrolysis by deletion of sEH amplifies
this effect.
Role of sEH on EET and prostaglandin levels at the inflammation side
To elucidate if EET levels differ at the site of inflammation, we determined their
levels in zymosan-inflamed paws. First, we compared EET levels in wild- type mice
at the onset (1-2 h), peak (6 h) and recovery phases (24-48 h) of inflammation after
zymosan injection. We found, no significant changes in EET levels up to 6 hours after
zymosan injection (Figure ). However, at later time points i.e., during the recovery phase, 8,9-EET and 14,15-EET
levels decreased significantly (Figure ). Next, we determined how EET levels were altered by sEH deletion, 48 h after zymosan
injection. In sEH-/- mice, 8,9- and 14,15-EET levels were significantly increased while the corresponding
DHETs were significantly reduced (Figure ). 11,12-EET levels and 5,6-EET were unaffected.
Consequences of sEH deletion on EET and prostaglandin levels in the inflamed paw. (A-C) Changes in EET levels during zymosan induced inflammation. Levels of 8,9-
(A), 11,12-(B) and 14,15-EET (C) levels were determined by LC-MS/MS in paw tissues
at different time points after zymosan injection. (D) Changes in EET and DHET (E)
levels after sEH deletion. Four different regioisomers were determined by LC-MS/MS
analysis in paw tissue from wild type and sEH-/- mice 48 h after intraplantar zymosan injection. Data shown represent the mean ± SEM
from tissues of 4-5 animals. Student's t test: *, p ≤ 0.05. (F) Effect of sEH deletion on COX-2 expression. COX-2 protein levels were
determined by western blot in paw tissues from wild- type and sEH deficient mice 48
h after intraplantar zymosan injection. (G) Effect of sEH deletion on prostaglandin
synthesis in the inflamed paw. Prostaglandins were determined 48h after zymosan injection
by LC-MS/MS analysis. Data shown represent the mean ± SEM from tissues of 3-5 animals.
(H) Effect of sEH deletion on prostaglandin synthesis in the spinal cord. Prostaglandins
were determined from lumbal spinal cord tissue 48h after zymosan injection. Data shown
represent the mean ± SEM from tissues of 4 animals.
Recent studies suggest that sEH inhibition has an antinociceptive effect in LPS-induced
hyperalgesia and in streptozocin-induced diabetic neuropathy [,,,]. One explanation for these findings is a direct anti-inflammatory effect of the sEH
inhibitors followed by suppression of COX-2 expression in the inflamed paw and spinal
cord [,]. Since COX-2 is also strongly upregulated in the zymosan model for up to 96 hours
[] and hyperalgesia in this model strongly depends on its activity [,], we examined COX-2 expression and prostaglandin synthesis in the sEH-/- mice. However, 48 h after zymosan injection, a time point at which COX-2 expression
remains upregulated in inflamed paws, no differences in COX-2 expression or PGE2, PGD2, TXB2, PGF2α or 6-keto-PGF1α levels at the site of inflammation or in the spinal cord were observed in wild type
and sEH knockout mice (Figure ).
Effects of EETs on nociceptive neurons
Next we determined whether increases in EET levels in sensory ganglia after sEH deletion
alters nociceptive neurons. To do this, we applied 8,9-, 11,12- or 14-15-EET to cultured
DRG neurons and determined calcium fluxes using fura-2-AM. At 1 μM none of the EETs
induced significant changes in intracellular calcium levels. However, 10 μM 8,9-EET,
but not 11,12- or 14-15-EET, increased intracellular calcium concentrations when applied
for 10 sec (Figure ). This calcium transient was observed in 4.7% of all neurons as determined by the
responsiveness of the cells to 40 mM KCl. We further characterized the responding
cells by co-stimulation with menthol, capsaicin and AITC to identify TRPM8, TRPV1
and TRPA1 expressing neurons respectively (Figure ). 8,9-EET responding neurons were predominantly capsaicin- and AITC-sensitive (92.3%)
showing that these cells are nociceptive neurons.
Activation of primary afferent neurons by 8,9-EET. (A) 8,9-EET induces direct calcium influx in sensory neurons. Calcium concentrations
were monitored by ratiometric imaging using fura-2 in cultivated neurons from DRGs
of adult wild- type mice. Shown is a representative trace. 40 mM KCL solution was
used to identify all viable neurons (B). Average values of peak normalized to baseline
ratios of experiments shown in panel B. Data shown represent the mean ± SEM from 12
experiments. Student's t test: *, p ≤ 0.05. (C) Phenotypic characterisation of 8,9-EET responsive neurons. Cultivated
DRG neurons were stimulated with 10 μM 8,9-EET, 200 μM menthol, 300 nM capsaicin,
100 μM AITC and 40 mM KCL for 10 sec. Representative traces are shown. Relative amount
of cells grouped according to their responsiveness to 8,9-EET and TRP-agonists. Percent
values were calculated from 280 KCl responsive neurons. Values below the dotted line
are the percentage of all 8,9-EET responding cells.
Next, we determined whether or not low concentrations of 8,9-EET contribute to the
sensitization of nociceptive neurons. Therefore, we studied the effect of 1 μM 8,9-EET
on AITC-induced calcium increases. We found that 8,9-EET significantly increased the
amplitude of AITC evoked calcium increases in cultured DRG neurons (Figure ). To further investigate nociceptor activation by 8,9-EET, we measured the release
of neuropeptide from freshly isolated sciatic nerves []. TRPA1 channels are expressed along peptidergic nerve fibers and neuropeptide release
can be stimulated by incubation with AITC []. To test, whether 8,9-EET can sensitize neuropeptide release from those fibers, we
incubated sciatic nerves with 10 μM 8,9-EET or with 3.2% ethanol (vehicle) before
stimulation with 50 μM AITC. While 8,9-EET had no direct effect, it significantly
sensitized AITC-induced CGRP release from the nerves (Figure ).
8,9-EET sensitizes AITC induced TRPA1 response. (A) Representative calcium imaging experiment of 8,9-EET dependent TRPA1-sensitization.
DRG cells were stimulated with AITC twice (50 μM, 15 sec) with 10 minutes interval.
1 μM 8,9-EET or vehicle was perfused for two minutes prior to the second AITC stimulation.
(B) Statistical analysis comparing the amplitudes of AITC induced TRPA1 response.
Data shown represent the mean ± SEM of 21-23 cells. Student's t test: *, p ≤ 0.05 (C) 8,9-EET sensitizes AITC induced CGRP release from sciatic nerve axons.
Freshly isolated sciatic nerves were incubated with 10 μM 8,9-EET (ipsilateral) or
3.2% EtOH (vehicle, contralateral) and stimulated with 50 μM AITC. CGRP was determined
by ELISA from extracellular solutions. Data shown represent the mean ± SEM from nerves
of 3-4 animals. Two way ANOVA with Bonferroni post test: *, p ≤ 0.05.
Effects of EETs and sEH deletion on nociceptive thresholds during inflammation
TRPA1-expressing neurons appear to have an important contribution to the development
of mechanical hyperalgesia [-]. To test whether 8,9-EET, which activates and sensitizes TRPA1-expressing neurons,
also induces mechanical hyperalgesia, we injected 10 μM of 8,9-EETs intraplantarily
and determined mechanical thresholds using the Dynamic Plantar Aesthesiometer. Injection
of vehicle (3.2% ethanol (v/v)) reduced mechanical thresholds 30 minutes after injection
(Figure ). However, this effect recovered after 1 hour while mice receiving 8,9-EET showed
significantly lower mechanical thresholds for up to 2 hours. In contrast, injections
of 11,12- and 14,15-EET had no effect on nociceptive behavior (Figure ).
Effects of EETs and sEH deletion on mechanical pain thresholds. (A) 8,9-EET induces mechanical hyperalgesia. 20 μl of 10 μM 8,9-EET was injected
intraplantar and mechanical thresholds were determined by the Dynamic Plantar Test.
Control animals received vehicle solution containing the corresponding volumes of
ethanol (3.2% v/v) and were tested in parallel. Data shown represent the mean ± SEM
from 8 animals per group. Two way Anova with Bonferroni post test: *, p ≤ 0.05; **, p ≤ 0.01. (B) Same experiments than in (A) but other EET regioisomers were used. (C)
Thermal thresholds after 8,9-EET injection. 20 μl of 10 μM 8,9-EET or vehicle was
injected and thermal thresholds were determined by the Hargreaves test. Data shown
represent the mean ± SEM from 4 animals. (D) Effect of sEH deletion on mechanical
hyperalgesia after zymosan injection. Mechanical thresholds of both hind paws were
tested by the dynamic plantar test after unilateral intraplantar zymosan injection
and compared between sEH-/- mice and wild type C57BL/6 controls. Data shown represent the mean ± SEM from 8-9
animals per group. Two way ANOVA with Bonferroni post test: **, p ≤ 0.01.
Interestingly, after 8,9-EET injection we did not detect any significant differences
in thermal thresholds, indicating that EETs affect noxious mechanical perceptions
specifically (Figure ). To investigate the role of sEH in nociceptive processing, we used the mice with
a targeted gene deletion of sEH, which exhibit much higher 8,9-EET levels in DRGs
after intraplantar injection of zymsoan A. Within the first 10 hours, wild type and
sEH-/- mice developed a comparable degree of hyperalgesia (Figure ). However, from 24 to 96 hours after the zymosan-injection, sEH-deficient mice exhibited
strongly reduced mechanical thresholds compared to the WT mice, indicating that deletion
of sEH prolongs mechanical hyperalgesia during peripheral inflammation.
Taken together, these data suggests that 8,9-EET sensitizes TRPA1 expressing primary
afferent neurons and that this may reduce mechanical thresholds. Moreover, sEH-/- animals which have significantly elevated 8,9-EET-levels in DRGs and paw show elevated
mechanical hyperalgesia during zymosan induced inflammation.
Discussion
In the present study we used a genetic mouse model to characterise the role of sEH
in nociceptive processing. Our findings suggest that the sEH is expressed in a subpopulation
of myelinated primary sensory neurons. There, it negatively regulates steady state
EET tissue levels as well as release of EETs from neurons. Specifically we show that
8,9-EET activates TRPA1-expressing neurons and sensitizes neuropetide release from
TRPA1-expressing fibers. Finally, we find that 8,9-EET can induce mechanical hyperalgesia
and that sEH deletion prolongs mechanical hyperalgesia during inflammation.
sEH exhibits lipid phosphatase as well as epoxide hydrolase activity, with the latter
resulting in rapid inactivation of EETs and epoxyoctadecenoic acids (EpOMEs) linoleic
acid metabolites [,]. Although a putative lipid phosphatase activity may have an impact on nociceptive
processing, we focused on the epoxide hydrolase activity of sEH and on EETs/DHETs
since they mediate most sEH functions in the cardiovascular system [,].
The EETs are generally attributed with anti-inflammatory effects, largely on the basis
of the fact that 11,12-EET can inhibit the IκB kinase and NF-κB signaling []. However, although 11,12-EET appears to be the most potent with respect to anti-inflammatory,
anti-migratory, and pro-fibrinolytic effects [], it has also been reported to increase COX2 expression in endothelial cells, a phenomenon
linked to angiogenesis []. This finding is of relevance, as one major consequence of immune cell activation
after zymosan injection is induction of COX-2 expression at the inflammation site
and in the spinal cord to sensitize nociceptive processing [,]. However, we found no obvious change in COX-2 expression or in prostaglandin synthesis
in paw tissue from zymosan-treated sEH-/- mice that could be attributed to the higher EET levels in these animals. Our results
suggest that sEH deletion, which mainly increases levels of 8-9- and 14,15-EET in
the tissue studied, does not alter zymosan-induced inflammatory hyperalgesia by an
inhibition of prostaglandin synthesis. These findings contrast with those made by
Incoeglu et al. who described that inhibition of sEH reduces PGD2 synthesis and hyperalgesia in a LPS model []. While these reports are difficult to reconcile, it is possible that differences
in the mechanisms of immune cell activation between the two models may be a determinant
factor since LPS and zymosan selectively activate toll like receptor 4 (TLR-4) and
TLR-2, respectively [] leading to expression and release of a different set of proinflammatory mediators,
which may be differentially affected by EETs.
All studies reporting antinociceptive effects of sEH are based on a pharmacological
inhibition of the enzyme [,,]. The model used in the present investigation was that of Ephx gene deletion resulting in the loss of the N-terminal lipid phosphatase activity and
C-terminal soluble epoxide hydrolase activity. Thus, one major difference between
our study and those performed previously is the fact that the sEH-associated lipid
phosphatase activity was also inhibited. As no endogenous substrates or pharmacological
inhibitors have been identified that target sEH lipid phosphatase activity, it is
currently not possible to distinguish between the contributions of the lipid phosphatase
and the epoxide hydrolase activities to the phenotype of sEH-/- mice []. Another possible explanation for the differences between this and previous studies
could be related to off-target effects of the urea-based inhibitors used in the pharmacological
studies. Although specificity for sEH over other epoxide hydrolases such as mEH is
generally good for most inhibitors, some compounds including 12-(3-adamantan-1-yl-ureido)-dodecanoic
acid (AUDA) can also activate PPAR□, which represses COX-2 expression [,].
We did not detect the sEH in invading immune cells within inflamed paw tissue. In
contrast, we found prominent expression of sEH in primary afferent neurons in the
DRG. It seems that also other components of the EET generating/metabolising pathway
are present in sensory neurons. The epoxygenases CYP2J3 and CYP2J4 are expressed together
with the sEH in trigeminal ganglia, indicating that the CYP450/sEH pathway is a common,
integral component of peripheral sensory neuron signaling [].
In the nociceptive models used in this study mechanical or heat stimuli are applied
to the plantar surface of the hind paw. Here, nociception is processed by small and
medium diameter DRG-neurons with C- and A-δ fibres. We found that the sEH was mainly
expressed in NF200 positive large and medium diameter DRG neurons. Most of these cells
transmit proprioceptive and low threshold mechanical but not noxious sensations. However,
we found that sEH deficient neurons have a higher steady state release of EET into
the extracellular medium. Further, we showed that an increase of AA bioavailability
predominately affects EET release instead of intracellular accumulation. This implies
that the sEH present in myelinated neurons may also affect C-fibre neurons by paracrine
signaling. Neuronal activity as well as inflammation activates PLA2 in primary afferents causing increased synthesis of eicosanoids. The restricted expression
pattern of sEH in a subset of DRG neurons and its hydrolyzing activity on EET, preventing
their release, suggests that the sEH may act to limit EET signaling to nociceptors.
Using cellular imaging experiments we identified 8,9-EET as the sEH substrate most
likely to sensitize nociceptor function. 8,9-EET induced a direct calcium influx in
≈5% of sensory neurons. Co-stimulation with different TRP-channel agonists revealed
that 8,9-EET only activates a subset of capsaicin responsive nociceptors and not large
non nociceptive neurons. Further, we found that within the capsaicin-responsive group
most were also AITC-responsive (TRPA1-positive neurons) (92.3% positive cells). Notably,
Kwan et al. reported that only AITC-sensitive neurons that also respond to TRPV1 express
TRPA1 []. Even though we did not further address the specific target of 8,9-EET in this TRPA1-expressing
cell population, multiple studies have shown that calcium transients in nociceptive
neurons induce plasticity changes through the activation of PKCε, p38 MAPK or ERK
which result in reduced activation thresholds and increased firing rates []. It could be speculated that 8,9-EET may directly activate TRPA1 at high concentrations
although various other possible targets exist. However, due to its electrophilic character,
8,9-EET can potentially sensitize TRPA1 by direct interaction with its intracellular
cysteins as previously described for other lipids like 4-HNE or cyclopentons [,]. In addition to a potential direct activation of the TRPA1-positive cell population,
we found that lower doses of 8.9-EET potentiate AITC-induced calcium flux. Here, 8,9-EET
may modulate TRPA1 indirectly via G-protein coupled receptors as described for bradykinin
[]. Moreover, to increase functionality of certain TRP-channels such as TRPC6, EETs
have already been shown to promote membrane translocation []. Finally, other TRPA1 independent downstream processes, such as sensitized voltage
gated calcium channels or calcium transporters may be involved in the observed increased
calcium responses.
We also investigated whether 8,9-EET modulates the activation of TRPA1-expressing
neurons. 8,9-EET application to isolated sciatic nerves caused an increased neuropeptide
release in response to AITC. Peripheral nerve axons resemble peripheral sensory terminals
in their common properties of sensory and signal transduction and CGRP neuropeptides
are stored all along axons of small diameter peptidergic neurons []. Stimulation of those cells by AITC induces a translocation of TRPA1 to the membrane
where it can be activated resulting in calcium influx, subsequent vesicle fusion and
neurotransmitter release. CGRP release from sciatic nerves can be sensitized by activation
of G-protein-coupled receptors and related protein kinases [] and that CGRP can induce mechanical hyperalgesia and central sensitization []. TRPA1 expressing neurons appear to mediate mechanotransduction and blockade of TRPA1
attenuates the development of mechanical hyperalgesia [,,]. Our finding that 8,9-EET increases CGRP release from TRPA1 expressing neurons strongly
suggests that sEH activity in primary afferents may prevents mechanical hypersensitivity.
In keeping with this, 8,9-EET injection into a hind paw lowered the mechanical but
not thermal threshold in wild- type mice. Wild- type mice recover from zymosan-induced
mechanical hyperalgesia 2 to 4 days after injection, a time during which 8,9-EET levels
decrease in the paw tissue. Our finding that 8,9-EET sensitizes primary afferents
and reduces mechanical thresholds suggested that hydrolysis of 8,9-EET could potentially
contribute to the resolution of mechanical hyperalgesia during inflammation. In accordance
with this hypothesis, sEH-deficient mice, which exhibit elevated 8,9-EET levels during
inflammation, show a dramatically reduced recovery from mechanical hyperalgesia.
Conclusion
sEH is expressed in sensory ganglia where it contributes to the metabolism of 8,9-EET
which can sensitize nociceptors. As a consequence of genetic deletion of sEH mice
exhibit exaggerated hyperalgesia during inflammation underlining the importance of
the antinociceptive function sEH. Pharmacological interventions influencing the EET-pathway,
which are actually highly considered to treat cardiovascular diseases, should therefore
be taken with care in terms of pain side effects.
sEH-/- mice [] were crossbred for 10 generations onto the C57BL/6 background. In all experiments
the ethical guidelines for investigations in conscious animals of the NIH and International
Association for the Study of Pain were followed, and the procedures were approved
by the local Ethics Committee. sEH-/- mice were compared with strain-, age-, and sex-matched C57BL/6 control mice.
Behavioral tests
To determine mechanical hyperalgesia mice were kept in test cages for 2 h on day one
for habituation. On day two, baseline paw withdrawal latencies (PWL) in response to
mechanical stimulation were determined. Briefly, animals were placed on an elevated
wire grid and the plantar surface of the paw stimulated using a Dynamic Plantar Aesthesiometer
(Ugo Basile, Comerio VA, Italy). We used a force increasing by 0.5 g every second
with an upper limit of 5 g. Paw withdrawal latencies were measured in sec ± 0.1 with
a cut off of 20 sec.
To test the effects of EETs on mechanical hyperalgesia, 20 μl of either 8,9-, 11,12-
or 14,15-EET (10 μM) were injected subcutaneously into the mid plantar side of the
left hind paw. Control animals received a corresponding volume of ethanol (3,2% v/v).
Mechanical hyperalgesia was assessed from 0.5 to 4 h after EET- injection using the
Dynamic Plantar Aesthesiometer.
Mechanical hyperalgesia after zymosan-induced inflammation has been described previously
[,]. 20 μl of a zymosan A (Sigma, Deisenhofen, Germany) suspension (12.5 mg/ml in phosphate
buffered saline) was injected subcutaneously into the plantar side of one hind paw.
Mechanical hyperalgesia was assessed from 0.5 h to 96 h after zymosan injection using
the Dynamic Plantar Aesthesiometer as described above.
To exclude gender differences only male mice were used. The non-injected and injected
paws were measured alternately at intervals of 5-10 min. For all behavioral tests
the observer was unaware either of genotypes or treatment.
Western blot analysis
For Western blot analysis we dissected skin tissue from the mid-plantar region of
the paw (1-3 mm deep) and collected DRGs from L4-6 segments. Tissues were homogenised
and sonicated in PBS and whole cell lysates containing 30 μg protein used for separation
on a 15% SDS-polyacrylamide gel. After blotting, COX-2 was detected with a polyclonal
antibody (1:500) from Cayman (Ann Arbor, MI). sEH was detected with a polyclonal anti-mouse
sEH antibody (dilution of 1:2000) raised against recombinant murine sEH produced in
a baculovirus expression system, and then purified to apparent homogeneity by affinity
chromatography. Antibodies against ss-actin (1:5000), HSP90 (1:2000) or ERK-2 (1:500)
(all from Santa Cruz, CA) were used to control for loading.
Immunohistochemistry using Multi-epitope ligand cartography (MELC)
DRGs and paws (48h after zymosan) were dissected from mice intracardially perfused
with 0.9% saline followed by 4% PFA/PBS (pH 7.4). After 24 h incubation in 30% sucrose/PBS
the tissue was cryostat sectioned at 10 μm and stored at 4° C until use.
To analyse sEH expression we combined conventional immunohistochemistry for sEH staining
with the MELC-technique using FITC-labelled antibodies against cell type specific
markers [,]. For sEH-staining we blocked the slices with 10% BSA and 1% mouse serum in PBS followed
by a 15 h incubation with a polyclonal rabbit anti sEH antibody (1:1000) diluted in
the blocking solution and a 1 h incubation with a anti-rabbit-Cy3 antibody (1:1000)
(Sigma, Deisenhofen, Germany) diluted in 1% BSA/PBS. To compare the sEH signal between
wt and sEH-/- mice the DRG slices were costained with NeuN and DAPI []. For comparison with multiple markers the slices stained for sEH were then transferred
to the MELC-system. Here, we used monoclonal anti-neuronal nuclei (NeuN) (Millipore,
Billerica, MA), monoclonal anti NF200 (AbD Serotec, Oxford, UK), monoclonal anti glial
fibrillary acidic protein (GFAP) (Sigma, Deisenhofen, Germany) and Isolectin B4 (IB4)
(Sigma, Deisenhofen, Germany) which were all directly labelled with FITC as described
Primary dorsal root ganglia (DRG)-cultures
DRGs were dissected from all spinal segments of adult mice and transferred to ice
cold HBSS with CaCl2 and MgCl2 (Invitrogen, Carsbad, CA, USA). For dissociation, isolated DRGs were treated for 90
min with collagenase/dispase (500 U/ 2.5 U/ml dispase) followed by
a 20 minutes incubation with 0.05% Trypsin (Invitrogen, Carsbad, CA, USA). After removal
of the enzyme-solutions, cells were washed twice with neurobasal medium containing
10% FCS respectively. Then cells were mechanically dissociated by pipetting (Gilson
1000 μl) and plated on culture dishes or poly-l-lysine (Sigma, Deisenhofen, Germany)
coated glass cover slips for calcium imaging. After two hours incubation, neurobasal
medium containing 2 mM L-glutamine, 100 U/ml penicillin, 100 μg/ml streptomycin, 50
μg/ml gentamicin and supplement B27 (all from Invitrogen, Carsbad, CA, USA) were added
and cells incubated for 24-48 h at 37° C without serum, NGF or any other neurotrophins.
Calcium imaging experiments
Calcium imaging experiments were performed on DRG cultures 24 h to 48 h after preparation.
Cells were loaded with 5 μM Fura-2-AM-Ester containing 0.02% Pluronic F-127 (both
from Biotrend, K?ln, Germany) and incubated for 30 to 45 min at 37° C. For baseline
measurements extracellular solution (145 mM NaCl,1.25 mM CaCl2, 1 mM MgCl2, 5 mM KCl, 10 mM D-glucose, 10 mM HEPES, pH 7.3) was added by bath application at
a flow-rate of 1-2 ml/min. EETs were dissolved in the extracellular solution at a
concentration of 1 and 10 μM. Cells were stimulated for 30 seconds. For control the
corresponding volume of ethanol was applied in the same way. At the end of each measurement,
cells were stimulated with 40 mM KCl for 30 seconds to identify viable neurons. Images
were taken with an Axioscope 2 upright microscope (Zeiss, Jena, Germany) using a 10x
Achroplan water immersion objective (Zeiss). The microscope was equipped with an Imago
CCD camera, a Polychrome IV monochromator (all TILL Photonics, Gr?felfing, Germany).
Images were acquired and processed using Tillvision software []. Quantitative characterization of 8,9-EET responding cells was done using a Nikon
Ti-E PFS Large Research Microscope cells were recorded every 8 sec. with a QImaging
EXI Aqua Digital Camera (Q-imaging, Surrey, BC, Canada) after alternating excitation
with 340nm and 380nm by a Ti-FL Epi-Fluorescence Illuminator (Sutter Instrument Company
Novato, CA) regulated by a SmartShutter Controller Unit (Sutter Instrument Company
Novato, CA). Emission was filtered by an ET FURA-2 Hybrid Filter. Extracellular solutions
were applied via a 360 μm perfusion pencil tip and a valve bank control system (both
from AutoMate scientific inc., Berkeley, CA) with >> 1 drop/sec gravity flow. Fluorescence
intensities of single cells were calculated by the NIS-Elements (AR Advance Research
Acquisition and analysis) software and transferred to Microsoft Excel for further
statistical analysis.
The DRG neurons were stimulated for 10 sec with 10 μM 8,9-EET, 200 μM menthol, 0.3
μM capsaicin, 100 μM AITC and 40 mM KCL. Neurons from 2 different dissections and
6 different experiments were used for calculations. For sensitization experiments,
DRG cells were stimulated with 50 μM AITC for 15 seconds twice with an interval of
10 minutes. 1 μM 8,9-EET were perfused over the cells for two minutes prior to the
second AITC-stimulation.
Sciatic nerve CGRP-release measurements
The experimental procedures were performed exactly as described before []. C57BL/6 mice were sacrificed in CO2-. Sciatic nerves were excised and continuously rinsed with synthetic interstitial
fluid (SIF, Bretag, 1969) consisting (in mM) of 107.8 NaCl, 26.2 NaCO3, 9.64 Na-gluconate, 7.6 sucrose, 5.05 glucose, 3.48 KCl, 1.67 NaH2PO4, 1.53 CaCl2 and 0.69 MgSO4, gassed with 95% oxygen and 5% carbon dioxide creating pH 7.4. Preparations were
placed in SIF inside a thermostatic shaking bath at 32°C for a 30 min washout-phase
before the experiment. All experiments were performed with matched pairs of a treated
and untreated side from the same animal. An experiment consisted of four incubations
of 5 minutes in mesh wells filled with 100 μl carbogen-saturated SIF. After two incubations
to determine basal CGRP release and spontaneous variation at 32°C, the samples were
stimulated with AITC (50 μM). 10 μM 8,9-EET was added in steps two to four. Immediately
after removal of the nerve from an incubation tube a sample volume of 100 μl was processed
using a CGRP-EIA kit (SPIbio, France) []. The CGRP detection level of the method is ~2 pg/ml.
Determination of EETs and their metabolites by liquid chromatography-tandem mass spectrometry (LC/MS-MS)
Sampling of tissue from DRGs and inflamed paws
For the determination of EETs and prostanoids during inflammation, 20 μl of 10 mg/ml
Zymosan A was injected intraplantarily into hind paws of wild type and sEH-deficient
mice. At indicated time points, DRGs from L4-6 segments and the tissue from the mid-plantar
region of the paw (1- 3 mm deepma) was dissected, weighed and directly transferred
to organic extraction solvents including 20 μl internal standards (5,6 EET-d11, 8,9
EET-d8, 11,12 EET-d8 and 14,15 EET-d8 all with a concentration of 200 ng/ml in methanol)
and stored at -80° C. The next day, tissue samples were homogenized in extraction
solvents using a Retsch Mixer-Mill MM 200 (Retsch, Haan, Germany).
Sampling of conditioned media from DRG cultures
For the determination of EET-release from sensory neurons we used primary" DRG-neurons
from wild type and sEH-deficient mice. After 24 h in culture, cells were stimulated
with 0.5 μM arachidonic acid and incubated for 2 h. Then, the cell culture supernatant
was removed, internal standards added, and samples directly extracted.
Sample extraction and standards
All standard substances were obtained from Cayman Chemical, Ann Arbor, MI, USA. Stock
solutions with 2500 ng/ml of all analytes were prepared in methanol. Working standards
were obtained by further dilution with a concentration range of 0.1-250 ng/ml for
EETs respectively.
Sample extraction was performed with liquid-liquid-extraction. Therefore tissue or
cell culture medium was extracted twice with ethyl acetate. The combined organic phases
were removed at a temperature of 45° C under a gentle stream of nitrogen. The residues
were reconstituted with 50 μl of methanol/water/(50:50, v/v), centrifuged for 2 minutes
at 10,000x g and then transferred to glass vials (Macherey-Nagel, Düren, Germany)
prior to injection into the LC-MS/MS system. Extraction and LC-MS/MS-instrumentation
for measurment of prostanoids was described previously [,].
Instrumentation for measuring 5,6-EET, 8,9-EET, 11,12-EET, 14,15-EET and their dehydro
metabolites
The LC/MS-MS system comprised an API 4000 triple quadrupole mass spectrometer (Applied
Biosystems, Darmstadt, Germany), equipped with a Turbo-V-source operating in negative
ESI mode, an Agilent 1100 binary HPLC pump and degasser (Agilent, Waldbronn, Germany)
and an HTC Pal autosampler (Chromtech, Idstein, Germany) fitted with a 25 μL LEAP
syringe (Axel Semrau GmbH, Sprockh?vel, Germany). High purity nitrogen for the mass
spectrometer was produced by a NGM 22-LC/MS nitrogen generator (cmc Instruments, Eschborn,
For the chromatographic separation a Gemini NX C18 column and precolumn were used
(150 mm × 2 mm i. d., 5 μm particle size and 110? pore size from Phenomenex, Aschaffenburg,
Germany). A linear gradient was employed at a flow rate of 0.5 ml/min mobile phase
with a total run time of 17.5 minutes. Mobile phase A was water/ammonia (100:0.05,
v/v) and B acetonitrile/ammonia (100:0.05, v/v).
The gradient started from 85% A to 10% within 12 min. This was held for 1 min at 10%
A. Within 0.5 min the mobile phase shifted back to 85% A and was held for 3.5 min
to equilibrate the column for the next sample. The injection volume of samples was
Quantification was performed with Analyst Software V 1.4.2 (Applied Biosystems, Darmstadt,
Germany) employing the internal standard method (isotope- dilution mass spectrometry).
Ratios of analyte peak area and internal standard area (y-axis) were plotted against
concentration (x-axis) and calibration curves were calculated by least square regression
with 1/concentration2 weighting.
Competing interests
The authors declare that they have no competing interests.
Authors' contributions
CB, KS, GG and CWJ conceived and designed the experiments. MS performed the behavior,
calcium imaging, histology, tissue sampling and CGRP release experiments. CB performed
Western blot, histology and calcium imaging experiments. OC performed behavior experiments.
KA performed Western blot experiments. IF and RPB provided the sEH knockout mice and
sEH antibodies. CA and HS performed the LC-MS/MS measurements. PWR and MJF introduced
the CGRP-release experiments. CB, KS, MS and CJW wrote the manuscript. All authors
read and approved the final manuscript.
Acknowledgements
This work was supported by the Deutsche Forschungsgemeinschaft stipend BR 2923/1-
and the LOEWE Lipid Signaling Forschungszentrum Frankfurt (LiFF). We thank Wiebke
Becker for technical assistance.
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