Next Article in Journal
Polycomb Repressor Complex 2 in Genomic Instability and Cancer
Next Article in Special Issue
Modulation of Astrocyte Activity by Cannabidiol, a Nonpsychoactive Cannabinoid
Previous Article in Journal
Stonin 2 Overexpression is Correlated with Unfavorable Prognosis and Tumor Invasion in Epithelial Ovarian Cancer
Previous Article in Special Issue
Cannabinoid Receptor 1 and Fatty Acid Amide Hydrolase Contribute to Operant Sensation Seeking in Mice
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:

Interactions between the Kynurenine and the Endocannabinoid System with Special Emphasis on Migraine

MTA-SZTE Neuroscience Research Group, University of Szeged, H-6725 Szeged, Hungary
Faculty of Health Sciences and Social Studies, University of Szeged, H-6726 Szeged, Hungary
Institute of Biochemistry, Biological Research Center, Hungarian Academy of Sciences, H-6726 Szeged, Hungary
Department of Neurology, Faculty of Medicine, Albert Szent-Györgyi Clinical Center, University of Szeged, H-6725 Szeged, Hungary
Author to whom correspondence should be addressed.
These authors contributed equally to this work.
Int. J. Mol. Sci. 2017, 18(8), 1617;
Received: 30 June 2017 / Revised: 18 July 2017 / Accepted: 21 July 2017 / Published: 30 July 2017
(This article belongs to the Special Issue Cannabinoid Signaling in Nervous System)


Both the kynurenine and the endocannabinoid systems are involved in several neurological disorders, such as migraine and there are increasing number of reports demonstrating that there are interactions of two systems. Although their cooperation has not yet been implicated in migraine, there are reports suggesting this possibility. Additionally, the individual role of the endocannabinoid and kynurenine system in migraine is reviewed here first, focusing on endocannabinoids, kynurenine metabolites, in particular kynurenic acid. Finally, the function of NMDA and cannabinoid receptors in the trigeminal system—which has a crucial role in the pathomechanisms of migraine—will also be discussed. The interaction of the endocannabinoid and kynurenine system has been demonstrated to be therapeutically relevant in a number of pathological conditions, such as cannabis addiction, psychosis, schizophrenia and epilepsy. Accordingly, the cross-talk of these two systems may imply potential mechanisms related to migraine, and may offer new approaches to manage the treatment of this neurological disorder.

Graphical Abstract

1. Introduction

The endocannabinoid system is involved in several neurological pathological conditions including neuropathic pain, inflammatory diseases, movement disorders (Parkinson’s disease and Huntington’s disease) and multiple sclerosis [1,2,3]. Cannabis has been used for a long time to treat nausea and vomiting, and to treat pain and migraine since the 6th century [4]. Migraine is one of the most prevalent neurological disorders, which affects about 16% of the population [5]. The total cost of healthcare for patients with migraine in Europe in 2010 was 18.4 billion € [6]. Growing evidence implies that endocannabinoid and glutamatergic systems are connected to migraine pathophysiology.
Human and animal data show that migraine is presumably hyperexcitability disorder, which means that the glutamatergic system is overactive [7]. In addition, an increasing amount of evidence suggests that migraine could alsobe linked to the kynurenine pathway (KP) itself [8].
Endocannabinoids, also known as “the body own cannabinoids” [9], and its receptors, have an extensive link with other endogenous receptors, such as opioid and glutamate ones, and especially the N-methyl-d-aspartate (NMDA) receptors. One of the endogenous NMDA receptor antagonists is kynurenic acid (KYNA), which is generated through tryptophan (Trp) metabolism. KYNA has a neuroprotective function and it might prove to be a future candidate in the treatment of migraine possibly by its NMDA antagonism.
The aim of this review is to demonstrate the interaction between the endocannabinoid and kynurenine system in relation to migraine. The review will discuss the involvement of each system in migraine separately, focusing on glutamate, kynurenines, endocannabinoids and the role of NMDA and cannabinoid receptors in the trigeminal system. Finally, we will cover the already demonstrated or possible interactions of the kynurenine and endocannabinoid systems, which can be potentially relevant to migraine.

2. Glutamate and Migraine

Glutamate is an ionic form of the nonessential amino acid glutamic acid, and it is the main excitatory neurotransmitter in the central nervous system [10]. Thus, it excites nearly every neuron contributing the primary neural transmission and pain perception [11,12]. As a neurotransmitter, glutamate is synthesized from glutamine, by the mitochondrial enzyme glutaminase, and is stored in synaptic vesicles. During neurotransmission, it is released from the stores to the synaptic cleft and removed by the presynaptic glutamate transporter and the transporter located on the neighboring glial cells. In glial cells, glutamate is converted to glutamine by glutamine synthetase. Thereafter, glutamine is transported out of the glia and picked up by nerve cells and transformed to glutamate [10].
Glutamate receptors can be divided into ionotropic and metabotropic receptors. The ionotropic receptors, namely NMDA, α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid (AMPA) and kainate receptors are ligand-gated ion channels. The metabotropic receptors are G-protein coupled receptors (GPCRs), which mean that their activations depend on a biochemical cascade [13].
Glutamate excitotoxicity is related to the hyperexcitability of NMDA receptors, as described in 1969 by Olney [14]. During this process, high glutamate stimulation leads to a large amount of Ca2+ is entering the cell [15] influencing many enzyme functions, such as phospholipases, proteases and endonucleases [16]. These mechanisms have a pivotal role damaging cell structures and DNA causing neuronal cell death. These receptors, especially the NMDA receptors have a crucial role in the pathomechanisms of migraine [17], supported by various experimental observations showing increased levels of glutamate in plasma, cerebrospinal fluid and platelets in migraineurs [18,19,20].
Glutamate has also a relevant role in the peripheral and central sensitization of the trigeminal system, is crucial in the pathomechanism of migraine [21]. Activation of NMDA receptors is one of the most important steps in initiating and maintaining the central sensitization [22], which can be blocked by competitive (D-CPP) and non-competitive (MK801) NMDA receptor antagonists in rats [23]. In addition, a conditional deletion of the NR2 subunit of NDMA receptors inhibits the synaptic inputs through NMDA receptors and the central sensitization in rats [24]. Besides NMDA, the metabotropic glutamate receptors also contribute to the mechanical allodynia [25]. Data from the human studies showed that the levels of glutamate were higher in the plasma, cerebrospinal fluid and platelets in migraine patients compared with non-migraineurs [20,26,27], which could indicate an increased activation of glutamate receptors, thus hyperexcitability [8].

3. Kynurenine Pathway (KP) and Migraine

KP is a dominant part of Trp metabolism, since 95% of Trp metabolizes this way. Trp, an essential amino acid, is transformed to N-formyl-l-kynurenine by tryptophan 2,3-dioxygenase (TDO) and indoleamine 2,3-dioxygenase (IDO), which are the rate-limiting enzymes of the KP. N-formyl-l-kynurenine can be further metabolized by formamidase to l-kynurenine (l-KYN), which is the precursor of KYNA (synthetized by kynurenine aminotransferases (KAT)s). l-KYN can also be degraded to anthranilic acid (ANA) by l-kynurenine hydrolase (KYNU) or to 3-hydroxy-l-kynurenine (3-HK) by kynurenine 3-monooxygenase (KMO). ANA and 3-HK are then further transformed to 3-hydroxyanthranilic acid (3-HA), which metabolizes to quinolinic acid (QUIN). 3-HK can be converted to xanthurenic acid as well. In thelast step of KP, QUIN is converted to nicotinamide adenine dinucleotide (NAD+) (Figure 1).
Among the KP metabolites, many compounds are biologically active. 3-HK and 3-HA are able to raise the formation of free radicals, yielding oxidative stress [28]. KYNA can exert its effect through NMDA and other glutamate receptors, namely AMPA [29] and kainate receptors [30].
In addition, KYNA has an agonistic effect of on the G protein coupled receptor 35 (GPR35) [31], which was found for a long time only in the gastrointestinal and in the immune system, namely in the crypts of Lieberkühn [32]. However, recent experiments showed that this receptor can also be found in the nervous system [33], and it has a relevant role in pain processing and neuroinflammation [34].
Opposite to KYNA, QUIN is an agonist of NDMA receptors and it can induce neuronal cell death [35] and lipid peroxidation [36]. In addition, QUIN is able to inhibit glutamate uptake in rats, resulting in raised extracellular glutamate levels [37] (Figure 1).
Several animal studies indicate that kynurenines, its analogs and halogenated derivatives have a future potential therapeutic action in the treatment of migraine. Since KYNA has a poor ability to cross the blood-brain barrier, its analogs and derivatives are tested experimentally. Halogenated derivatives, 4,6-dichlorokynurenine and 4-chlorokynurenine are converted to KYNA derivatives (7-chlorokynurenic acid and 5,7-dichlorokynurenic acid), which have increased affinity to the glycine-binding site of NMDA receptors [38,39].
One of the human and animal models of migraine is the administration of nitroglycerin (NTG), which is a nitric oxide (NO) donor. NTG is able to activate and sensitize the trigeminal system, which are the crucial mechanisms in the pathophysiology of migraine [40,41]. Administration of l-KYN and probenecid (an inhibitor of KYNA secretion from the central nervous system (CNS) together or KYNA analogs ((N-(2-N,N-dimethylaminoethyl)-4-oxo-1H-quinoline-2-carboxamide hydrochloride (KA1) and N-(2-N-pyrrolidinylethyl)-4-oxo-1H-quinoline-2-carboxamide hydrochloride (KA2)) were effective to inhibit the NTG caused morphological and behavioral changes in rats likely via the inhibition of NDMA receptors [42,43,44]. In this model, NTG was able to decrease the expression of kynurenine aminotransferase II (KATII) [45], which is the key enzyme of KYNA production. In a recent study, it was also shown that NTG altered the expression of other enzymes of the KP, namely TDO, IDO, KYNU and KMO, suggesting that NTG/NO has an influence on the KP [46].
Another animal model of trigeminal activation and sensitization is the application of Complete Freund’s Adjuvant (CFA) on the dural surface causing inflammation. In this experimental setting, Lukács and her colleagues showed that KA1 was able to attenuate the CFA-caused inflammation [47].
The orofacial formalin test is also a suitable model for mimicking the trigeminal activation and sensitization [48]. In this model, probenecid decreased the nociceptive behavior in rats, probably via increasing the concentration of KYNA [42]. In a recent study KA1 and KA2 were able to abolish the formalin induced behavioral and morphological changes, and increased KYNA levels [49]. On the other hand, in the combined model of NTG and formalin, KA1 also inhibited the behavioral and morphological alterations [50]. In addition, in the electrical stimulation model of the trigeminal activation a decreased KAT immunoreactivity has been shown in the rat’s dura mater [51].
Cortical spreading depression (CSD) is a self propagating wave in the cortical areas of the brain and it has a relevant role in the pathomechanism of migraine [52], as it is well accepted that CSD is the basis of the aura phenomena [53]. In the model of CSD, two new KYNA analogs inhibited the propagation of CSD waves [54], likely by the inhibition of glutamate receptors. Since glutamate receptors have a crucial role in the propagation and generation of CSD [55], they possibly represent a link between migraine and CSD.
It is also important to note that, the levels of the metabolites of the KP were found to be changed in migraineurs. Curto and her colleagues discovered decreased level of kynurenine metabolites in the serum of patients with chronic migraine and cluster headache [56,57], the data of which are in accordance with animal results from the NTG model of migraine [45]. These findings suggest that the decreased levels of KYNA mean that the glutamatergic system is overactive in chronic migraine as well as cluster headache.
The role of KYNA and its metabolites in the pathomechanisms of migraine is still not fully known. The effect of KYNA could manifest through both peripheral and central mechanisms. On the periphery, KYNA can modulate glutamate receptors, principally, NMDA receptors localized in the dorsal root and trigeminal ganglia [58]. The other peripheral place, where KYNA and its analogs can exert their effect is GPR35, which is also present in the dorsal root ganglion (DRG) [59]. Besides the peripheral effects, KYNA and its analogs have an impact on second-order neurons too, proven by that KYNA can decrease mechanical allodynia and pain sensitivity in the hot-plate and tail-flick tests [60,61].
To summarize the human and animal data, we can conclude that the KP has a relevant role in the pathomechanism of migraine, and it might be promising future therapeutic target in the treatment of headaches.

4. The Endocannabinoid System and Migraine

4.1. The Endocannabinoid System

The endocannabinoid system comprises of the endogenous ligands called endocannabinoids, the enzymes which synthesize and degrade them and the receptors, to which these ligands bind. Two types of cannabinoid receptors have been cloned so far, the type 1 and type 2 cannabinoid receptor (CB1 and CB2) from rat cerebral cortex [62] and human promyelocytic leukaemia cells, respectively [63]. They are class A GPCRs and belong to the Gi/o-coupled GPCR superfamily and they couple to Gi/o type inhibitory G-protein, thus their activation inhibits cyclic adenosine monophosphate (cAMP) production and stimulates mitogen-activated protein kinases (MAP). The CB1 receptor is the most abundant GPCR in the CNS and its density is comparable to that of the glutamate, γ-Aminobutyric acid (GABA) and dopamine receptors [62,64,65]. It can also be found in the periphery such as in the liver, adipose tissues, muscles, cardiovascular and gastrointestinal system [66,67]. The CB2 receptors are mainly present on immune and hematopoietic cells [2,68], but can also be found in the CNS especially in microglia [69], in the periphery on myocardial cells [2,66] and in the endothelium [66,70]. CB1 receptors are responsible for mood regulation and can also induce antinociception, regulate energy balance and endocrine functions [66,67,71]. The major function of the CB2 receptor is the control of cytokine release and immune cell migration (reduce inflammation-induced pain, reveal peripheral antinociception, inhibition of tumor growth) [72]. CB2 receptor can decrease nociception so far without any detectable tolerance [72,73] and side effects [74]. Activation of this receptor caused analgesia in the tail flick and the orofacial formalin test [75], as well.
Endocannabinoids are lipid-derived hydrophobic compounds, among them N-arachidonoylethanolamine (anandamide, AEA) and 2-arachidonoylglycerol (2-AG) are the most studied [76,77,78]. AEA is a full agonist of CB1 and partial agonist of CB2 receptors. AEA is synthesized from membrane phospholipids via N-acyl transacylase and N-acyl phosphatidylethanolamine-phospholipase D [79]. In the degradation of AEA is proceeded mainly by fatty acid amide hydrolase (FAAH), which degrades AEA to arachidonic acid and ethanolamine [79,80]. Blocking the FAAH enzyme by irreversible inhibitors such as URB597 has been reported to be a promising treatment for smoking addiction [66,81,82] and it also enhanced opioid analgesia [83,84]. 2-AG is formed from the omega-6 fatty acid arachidonic acid and glycerol [79]. 2-AG can be found in relatively high concentration in the nervous system [85] and it is mainly degraded by monoacylglycerol lipase (MAGL) [86,87].

4.2. The Role of Endocannabinoids in Migraine

Cannabis has been used for migraine medication since 6th century and the deficiency of endocannabinoid system contributes to the pathophysiology of the disorder [4]. This system has a crucial role in the pathomechanisms of pain [88] and its activation is essential in the inhibition of trigeminal neurons [89].
Clinical studies shown that the formation of AEA and 2-AG were down-regulated in migraineurs [90,91]. In patients with chronic migraine a decreased AEA level was found in the cerebrospinal fluid [92], as well. In addition, in the blood of female migraine patients, a raised FAAH enzyme level and a decreased AEA level has been shown [93]. The reduced levels of AEA might promote the hyperactivity of the trigeminal system and the reduced inhibitory impact of the endocannabinoid system [94,95], both contributing to migraine development.
Numerous studies show that endocannabinoids are effective in the animal models of migraine. Endocannabinoids control the cerebrovascular tone and contribute to NO production [88]. In the NTG model of migraine, AEA was able to inhibit the NTG induced increase of c-Fos expression in rats [95], one of the markers of trigeminal neuronal activation. Besides that, AEA can also abolish the NTG caused elevation of the levels of the sensitization markers in rats [45] and also inhibit the NO induced dural vasodilatation [96]. AEA is effective in the inhibition of the NTG induced KAT-II expression decrease [45], suggesting an influence on the KP. In these above-mentioned experiments, the main modulatory effect was probably achieved by the CB1 receptor, which can be activated by increased AEA levels [97]. This notion is supported by the experiments in which CB1 receptor activation was able to alleviate the KCl-induced CSD [98] and repressed the Aδ neuron activity in rats [99]. In addition, AEA can alter NO and calcitonin gene-related peptide (CGRP) induced dural vasodilatation [96]. CGRP is the main peptide in the pathomechanisms of migraine, its level is raised in external jugular veins in migraineurs [100] and it co-localizes with CB1 receptors [88]. On the other hand, 2-AG created an anti-nociceptive effect in the formalin test [101] and endocannabinoid uptake inhibitors were effective in pain relief in the formalin test [102], which shows that the levels of endocannabinoid are important in the pain inhibition process. WIN55121, a potent cannabinoid agonist can inhibit the wind-up process, which is a centrally mediated enhance of C-fibers and contributes to the improvement of allodynia [103].
The molecular or genetic inhibition of the AEA degradation enzymes enhance cannabinoid signaling and raise the levels of AEA in the brain [80], thus making an opportunity to influence the levels of the endocannabinoids in many experimental protocols. Indeed, FAAH inhibitors have an analgesic effect in the inflammatory and the neuropathic models of pain [104]. In addition, NTG was able to enhance the enzyme levels of FAAH and MAGL [95], thus contributed to the decreased AEA and 2-AG levels in animals.

5. Cannabinoid and Glutamate Receptors in the Trigeminal System

In migraine pathomechanism, trigeminal system has a pivotal role. The trigeminal system consists of the peripheral Aδ and C-fibers, which convey to the trigeminal ganglion from the peripheral skin and meningeal blood vessels. The brainstem includes the second-order trigeminal neurones, which receive nociceptive afferents from trigeminal ganglion and modulating afferents from other brainstem structures, as locus coeruleus, nucleus raphe magnus and periaqueductal grey (PAG), which also called migraine generators. The information from brainstem nuclei proceeds to the somatosensory cortex via the third-order neurons located in the thalamus.
Cannabinoid receptors can be found throughout the trigeminal system and in the migraine generators, as well. In the trigeminal system CB1 is present in the trigeminal ganglion, in the rostral and caudal PAG and on the peripheral and central axon terminals of trigeminal primary sensory neurons [88]. They are also present in the human or rat thalamus and PAG [105,106] and rostral ventromedial medulla [88], from where trigeminal system receives inputs. CB2 is also present in the nervous system, can be found in the afferent fibers in the dorsal horn of the spinal cord [107]. Thus, activation of these receptors may modulate the neuronal firing of the trigeminal system [94]. Activation of glutamatergic projections is able to facilitate the synthesis of endocannabinoids in the glutamatergic terminals. CB1 receptors are located on these terminals and their activation can decrease the excitatory transmission and glutamate induced hyperexcitability [108]. On the other hand, a genetic association study found strong haplotypic associations between the CB1 gene and the three prognostic symptoms of migraine, as disability, nausea and photophobia [109], suggesting that the haplotype causes diminished CB1 function or expression.
Glutamate receptors are also found in the trigeminal system. NMDA, AMPA, and kainate receptors are present within the brainstem nuclei of the trigeminal system [110] and NMDA receptor mRNA was found in the trigeminal ganglion [111]. NMDA, AMPA, and kainate receptors can also be found in the superficial layers of the spinal cord [112], where the brainstem trigeminal nuclei extend.
Several experimental data show that the levels of glutamate and endocannabinoid are altered in migraine patients. To summarize human and animal data with kynurenines and cannabinoids, we can conclude that they have a relevant role in the pathomechanisms of migraine, probably also by their interaction.

6. Known and Potential Functional Interactions betweenthe Endocannabinoid and Kynurenine System: Possible Pharmacological Targets against Migraine

Thus far there are rather limited data demonstrating the interaction between the kynurenine and endocannabinoid systems [108] and their implication in migraine has not yet been investigated at all. Nevertheless, the functional co-operation between the two systems may reveal other, yet unknown mechanisms, which might be involved in migraine and may display novel potential therapeutic targets. There are studies describing connections between exogenous cannabinoids and the enzymes of the KP, whereas other data demonstrate the anatomical and functional interactions between CB1 and NMDA receptor, in which KYNA is used more as a tool to indicate the involvement of the NMDA receptor in the described effects [108]. Both possibilities will be discussed in this section, together with potential interplay between the cannabinoid and further target receptors of KYNA. An additional interaction will also be proposed among the µ opioid, the CB1 and the NMDA receptor.

6.1. The Relationship between Enzymes of the KP and Exogenous Cannabinoids

Enzymes of the endocannabinoid and kynurenine system have been demonstrated to be potential therapeutic targets in several pathological conditions [113,114,115,116], including migraine [116,117]. Thus far no direct interaction has been described between these enzymes, however there issome evidence for overlapping expression profile in certain neurons. For instance, IDO and FAAH enzymes are both expressed in neurons of the hippocampus and dentate gyrus [118,119]. On the other hand, Jenny M. Santer et al. demonstrated that Δ9-tetrahydrocannabinol (THC) or cannabidiol (phytocannabinoids of the cannabis sativa plant) in nanomolar concentrations can enhance mitogen-stimulated IDO enzyme activity (Figure 1), which was dependent from CB1 or CB2 receptor activity. In contrast, in micromolar concentrations both compounds suppressed the activity of the IDO enzyme independently from cannabinoid receptors, consequently enhancing TRP levels for serotonin synthesis (Figure 1), which overall may contribute to improve mood disturbance [120]. Incidentally, cross-talks between the CB1 and serotonin (5-hydroxytryptamine) receptors has been also described previously [99,121,122].
Furthermore, Justinova and co-workers reported that by enhancing brain KYNA levels with a KMO (Figure 1) inhibitor (Ro 61-8048) attenuated cannabinoid-induced increase in dopamine levels in the nucleus accumbens (NAc) shell and thus reduced the rewarding effects of THC and WIN55,212-2 [123]. This study also demonstrated that the anti-abuse action of elevated brain KYNA levels by KMO inhibition is due to KYNA-induced negative allosteric modulation on the α7 nicotinic acetylcholine receptor (α7nAChR), since this effect was prevented by a α7nAChR selective positive allosteric modulator (PNU120596) [123]. Thus, the selective elevation of brain KYNA levels can be suggested a potential novel strategy for treating human marijuana dependence and it can be considered a pharmacologically safe approach since Ro 61-8048 has not been associated with adverse side effects [123].
Accordingly, cannabinoids can alter the activity of certain enzymes of the KP and vice versa, manipulating KYNA levels through enzyme inhibition in the kynurenine pathway can also alter exogenous cannabinoid activity. However, it is yet to be examined whether altering the levels of endocannabinoids for instance via FAAH or MAGL inhibitors induces any changes in the kynurenine pathway and vice versa.

6.2. The Type 1 Cannabinoid Receptor-N-methyl-d-aspartate (CB1-NMDA) Receptor Complex

Among the known receptor targets for KYNA, the NMDA receptor is the most cited and studied and therapeutically one of the most relevant targets in terms of neuropathic pain and migraine [116]. The NMDA receptor is a ligand-gated cation channel, permeable for monovalent and Ca2+ ions. The ion permeability is co-activated by glutamate and glycine through separate binding sites. NMDA receptors consist of three types of subunits, NMDA receptor subunit 1 (NR1), NMDA receptor subunit 2 (NR2) and NMDA receptor subunit 3 (NR3), which form a functional tetramer of an obligatory pair of NR1 and two NR2 or NR3 subunits [124]. KYNA binds to the glycine B binding site of the NMDA receptor [125] with micromolar affinity [126,127], antagonizing the effects of the receptor [128].
The CB1 and the NMDA receptor systems are involved in multiple processes, such as learning and memory [129,130,131], drug reinforcement [132,133,134], or nociception [88,135,136]. Nevertheless, the most widely studied and known pathological disorders in which these two receptors interact are psychosis, schizophrenia and epilepsy [124,137].
NMDA and CB1 receptors have been demonstrated to be post and pre-synaptically co-localized [138,139,140,141,142] allowing them to functionally interact in both sides of the synapse [143]. Moreover, a physical association has been previously described between the C-terminal of the CB1 receptor and NR1 subunit of the NMDA receptor [144]. Incidentally, NMDA receptors are known to interact physically with other GPCRs as well, such as the μ opioid receptor (MOR), dopamine D1 receptor, group 1 metabotropic glutamate receptor [145,146,147]. The CB1-NMDA receptor complex requires the presence of the histidine triad nucleotide-binding protein 1-σ receptor type 1 (HINT1-σ1R) protein tandem [142,148], which works as an on-off switch, connecting and disconnecting the two receptors, which can be regulated by calcium and exogenous σ1R ligands [149]. The HINT1-σ1R protein tandem has been described in the MOR-NMDA receptor complex as well [150].
It is now well-established that both endogenous and exogenous cannabinoids reduce the activity of NMDA receptors through the CB1 receptor [142,151,152] by reducing pre-synaptic glutamate release or alter post-synaptic NMDA receptor mediated signaling pathways [143,152] and it also requires both HINT1 and σ1R proteins [150,153]. Additionally, exogenous cannabinoids effectively induce CB1 receptor internalization, which disassembles and deactivates the CB1-NMDA receptor complex, dampening NMDA receptor activity, thus reducing the risk of NMDA receptor mediated excitotoxicity [142,154]. NMDA receptor hyperactivity is one of the main characteristics of epilepsy and NMDA receptor antagonists display antiepileptic effects in clinical and preclinical studies, unfortunately associated with serious side-effects, such as memory dysfunctions or motor disturbances [155]. Phytocannabinoids, such as THC and cannabidiol have been reported to reduce epileptic seizures, utilizing the CB1-NMDA receptor complex, thus introducing a possible new therapeutic approach for epilepsy [124]. However, excessively reducing NMDA receptor activity can lead to NMDA receptor hypofunction, which is implicated with psychosis and schizophrenia [137]. Indeed, cannabis abuse has been associated with these two mental disorders [156,157]. The HINT1 and σ1R genes have also been implicated in schizophrenia [158,159,160], moreover the σ1R agonist pregnenolone and antagonists showed promising results in reducing the symptoms of schizophrenia in clinical trials [161,162]. Additionally, the CB1-NMDA receptor complex has been implicated in analgesia as well. Although cannabinoid mediated analgesia does not require the association of CB1 receptor with the NMDA receptor, when coupled, NMDA receptor antagonists can significantly reduce cannabinoid-induced analgesia, the effect of which is HINT1 protein dependent [142].
Thus, the CB1-NMDA receptor complex, including the HINT1-σ1R protein tandem offers a promising new approach for the therapeutic management of certain neurological disorders [124]. NMDA receptor hyperactivity is one of the main pathomechanisms of migraine [8] and since CB1 receptor has been demonstrated to hinder this activity, the CB1-NMDA receptor complex could be a potential therapeutic target for migraine. Furthermore, based on the previous section and the data above mentioned, this can be extended by the inhibition of endocannabinoid metabolizing enzymes or by manipulating the enzymes of the kynurenine pathway. This way, the enhanced levels of endocannabinoids, or exogenously administered cannabinoids via CB1 receptor and/or enhanced levels of KYNA may represent an alternative approach for the reduction of NMDA receptor hyperactivity and thus against migraine (Figure 2). Additionally, the NMDA receptor being one of the key receptor target for KYNA, bifunctional KYNA-cannabinoid/σ1R ligands or co-administration of KYNA and cannabinoids might be a further therapeutic tool to utilize the CB1-NMDA receptor association in migraine or other neurological disorders.

6.3. Possible Interaction between the μ Opioid, the CB1 and the NMDA Receptor

There are extensive data describing the interaction between the kynurenine and the endogenous opioid system [60,163,164,165,166,167]. It has also been demonstrated that KYNA and its analog KA1 can indirectly alter the G-protein signaling of opioid receptors through the NMDA receptor depending on the opioid receptor type (μ, κ or δ) and brain region (cortex or striatum) [168,169]. More importantly, the opioid, cannabinoid and the NMDA receptors are known to be co-localized and functionally interact with each other pair-wise in areas relevant to opioid dependence, tolerance and antinociception [84,142,170,171,172,173,174,175]. Additionally, similar to CB1 receptor, MOR also physically associates with the NR1 subunit of the NMDA receptor, and it is also under the control of the HINT1 and σ1R proteins [149]. Interestingly, in contrast to CB1, MOR promotes the activity of the NMDA receptor in this complex [150]. Thus, KYNA might alter the functionality of cannabinoid and opioid receptors via the NMDA receptor and might form a functional “triangle”. Such a proposal has been reviewed previously between opioid, cannabinoid receptors and the transient receptor potential vanilloid type 1 (TRPV1) channel in terms of analgesia [176].

6.4. The Interaction between the CB1 and the α7nACh Receptor

There are numerous publications reviewing the interaction between the cholinergic and endocannabinoid system which has been reviewed elsewhere [177]. In this section we are focusing on the α7nAChR, which KYNA directly inhibits as firstly reported by Hilmas et al. [178] and further confirmed by several in vitro and in vivo reports (for review see [179]). However, other studies also questioned this effect [180,181]. A recent review corroborated that KYNA can be considered a bona fide endogenous modulator for α7nAChR, but established as a complex phenomenon, depending on mostly methodological considerations [179].
The α7nAChR belongs to the neuronal type nAChRs and shares the five transmembrane subunit structure with this subfamily, but it consists of only α7 subunits, creating a homomeric structured ion channel [182]. Additionally, among the neural homomeric nAChRs, α7nAChR is the most abundant in the mammalian brain [182,183]. As all nAChRs, the α7nAChR is too a cation sensitive ion channel, with a high calcium permeability. They are expressed extensively in the cortex and hippocampus and in neurons of the mesostriatal dopaminergic system [182], which also overlaps with the expression of the cannabinoid receptors. Similar to the CB1 receptor [184], the α7nAChR are presynaptic receptors and modulate the release of GABA, dopamine, noradrenaline and serotonin neurotransmitters, however in contrast to CB1 it enhances their release [185]. Hence, the α7nAChR share a number of functions with the CB1R including learning, memory or nicotine addiction [182].
The α7nAChR and cannabinoids have been in focus in terms of cannabis abuse. Solinas et al. demonstrated that the blockade of α7nAChR reversed the discriminative effects of THC and the synthetic cannabinoid agonist WIN55,212-2. Additionally, the α7nAChR antagonist methyllycaconitine also reduced WIN55,212-2 self-administration and blocked THC-induced enhanced dopamine levels in the NAc shell, which is an important region of rewarding effects [186]. Importantly these effects were observed at doses that do not induce psychiatric side-effects or toxicity [186].

6.5. GPR35: A Possible Interactional Partner for Cannabinoid Receptors

Among the direct receptor targets of KYNA, the GPR35 stands out being the only metabotropic receptor. GPR35 was cloned as an “orphan” GPCR [32] and it is reported to couple to Gαi/o and Gα13 type G-proteins [31,33,187,188,189]. KYNA has been the first endogenous agonist ligand to be described for GPR35 [31], also the presence of KYNA overlaps with the expression of GPR35 in many tissues and organs (e.g., spleen, colon, and brain) [190]. However, the GPR35 endogenous ligand designation of KYNA is still a matter of debate, mainly because it activates the receptor in very high micromolar concentrations and it has been reported to be significantly more potent on rat than human GPR35 [31]. Several other endogenous GPR35 ligands have been described [191], among them the lysophosphatidic acid is the other well studied endogenous ligand besides KYNA [192].
Both cannabinoid receptors and the GPR35 distribution overlaps in certain cells or tissues such as in leukocytes in the immune system (CB2 receptors), the gastrointestinal system (GI) or in neurons of DRG and both have similar signaling pathways (coupling to Gαi/o, inhibiting adenylate cyclase activity etc.). Although there are no data referring to GPR35 and CB1 receptor being co-localized or to functionally interact in these areas, there are studies indicating its possibility. For instance, in DRG neurons, there is evidence that both CB1 and GPR35 are co-localized or co-expressed together with the TRPV1 channel in the small DRG neurons [59,126]. Additionally, both GPR35 and CB1 receptors mediate peripheral nociception [191,193], which in case of GPR35 might be induced by KYNA [59]. Another potential region for an anatomical and functional interaction of GPR35 and cannabinoid receptors could be the small intestines, colon or stomach, where both receptors are expressed in significant quantities [31,66,127,194], however the cell specific expression of GPR35 within the GI system has not been established yet in contrast to cannabinoid receptors [66]. At the same time, both GPR35 and cannabinoid receptors have been implicated in inflammatory bowel disease [195,196]. Further, both GPR35 via KYNA and CB2 receptors expressed on leukocytes have been reported to be involved in leukocyte recruitment [197,198,199,200,201,202,203,204]. Finally, although KYNA is not implicated in this finding, a review suggested a linkage between GPR35 and cannabinoid receptors through the interconversion of their endogenous ligands, the 2-acyl lysophosphatidic acid (GPR35) and 2-AG (cannabinoid receptors) [191].

7. Final Remarks and Conclusions

In summary, both the endocannabinoid and kynurenine systems can interact with each other in several levels, which might be relevant to migraine. However, the data are so far limited and some areas of this interaction are yet undiscovered, for instance the cross-talk between the endocannabinoids and their enzymes with the kynurenine pathway. Nevertheless, previous successful preclinical or clinical studies in regard of other pathological disorders demonstrated that the endocannabinoid and kynurenine system are potential therapeutic targets. Presently, the well-described NMDA-CB1 receptor complex might be the most promising therapeutic target against migraine, by either manipulating the endogenous levels of KYNA and endocannabinoids (Figure 2) and/or using specific exogenous compounds to target this receptor complex.
It is important to further investigate the mechanisms and interactional partners involved in the cross-talk between the endocannabinoid and kynurenine system, as it will also possibly reveal more information regarding the individual function of these two systems and also their connections with migraine.


This work was supported by the Hungarian Brain Research Program (Grant No. KTIA_13_NAP-A-III/9); the EUROHEADPAIN (FP7-Health 2013-Innovation; Grant No. 602633) and GINOP-2.3.2-15-2016-00034. Árpád Párdutz was supported by the Bolyai Scholarship Programme of the HungarianAcademy of Sciences. Szabolcs Dvorácskó was supported by the ÚNKP-16-3 New National Excellence Program of the Ministry of Human Capacities of Hungary.

Conflicts of Interest

The authors declare no conflict of interest.


3-HA3-hydroxyanthranilic acid
AMPAα-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid
ANAanthranilic acid
cAMPcyclic adenosine monophosphate
CB1type 1 cannabinoid receptor
CB2type 2 cannabinoid receptor
CFAcomplete Freund’s Adjuvant
CGRPcalcitonin gene-related peptide
CSDcortical spreading depression
DRGdorsal root ganglion
FAAHfatty acid amide hydrolase
GABAγ-Aminobutyric acid
GPCRG-protein coupled receptor
GPR35G protein coupled receptor 35
HINT1histidine triad nucleotide-binding protein 1
IDOindoleamine 2,3-dioxygenase
KA1N-(2-N,N-dimethylaminoethyl)-4-oxo-1H-quinoline-2-carboxamide hydrochloride
KA2N-(2-N-pyrrolidinylethyl)-4-oxo-1H-quinoline-2-carboxamide hydrochloride
KATIIkynurenine aminotransferase II
KMOkynurenine 3-monooxygenase
KPkynurenine pathway
KYNAkynurenic acid
KYNUl-kynurenine hydrolase
MAGLmonoacylglycerol lipase
MAPmitogen-activated protein kinases
MORμ opioid receptor
NAD+nicotinamide adenine dinucleotid
NOnitric oxide
NAcnucleus accumbens
PAGperiaqueaductal grey
QUINquinolinic acid
TDOtryptophan 2,3-dioxygenase
α7nAChRα7 nicotinic acetylcholine receptor
σ1Rσ receptor type 1


  1. Rodríguez de Fonseca, F.; Del Arco, I.; Bermudez-Silva, F.J.; Bilbao, A.; Cippitelli, A.; Navarro, M. The endocannabinoid system: Physiology and pharmacology. Alcohol Alcohol. 2005, 40, 2–14. [Google Scholar] [CrossRef] [PubMed]
  2. Pacher, P.; Bátkai, S.; Kunos, G. The endocannabinoid system as an emerging target of pharmacotherapy. Pharmacol. Rev. 2006, 58, 389–462. [Google Scholar] [CrossRef] [PubMed]
  3. Di Marzo, V.; Bifulco, M.; De Petrocellis, L. The endocannabinoid system and its therapeutic exploitation. Nat. Rev. Drug Discov. 2004, 3, 771–784. [Google Scholar] [CrossRef] [PubMed]
  4. Russo, E.B. Clinical endocannabinoid deficiency (CECD): Can this concept explain therapeutic benefits of cannabis in migraine, fibromyalgia, irritable bowel syndrome and other treatment-resistant conditions? Neuro Endocrinol. Lett. 2004, 25, 31–39. [Google Scholar] [PubMed]
  5. Smitherman, T.A.; Burch, R.; Sheikh, H.; Loder, E. The prevalence, impact, and treatment of migraine and severe headaches in the United States: A review of statistics from national surveillance studies. Headache 2013, 53, 427–436. [Google Scholar] [CrossRef] [PubMed]
  6. Olesen, J.; Gustavsson, A.; Svensson, M.; Wittchen, H.U.; Jonsson, B. The economic cost of brain disorders in Europe. Eur. J. Neurol. 2012, 19, 155–162. [Google Scholar] [CrossRef] [PubMed]
  7. Vikelis, M.; Mitsikostas, D.D. The role of glutamate and its receptors in migraine. CNS Neurol. Disord. Drug Targets 2007, 6, 251–257. [Google Scholar] [CrossRef] [PubMed]
  8. Vecsei, L.; Majlath, Z.; Balog, A.; Tajti, J. Drug targets of migraine and neuropathy: Treatment of hyperexcitability. CNS Neurol. Disord. Drug Targets 2015, 14, 664–676. [Google Scholar] [CrossRef] [PubMed]
  9. Grotenhermen, F.; Muller-Vahl, K. The therapeutic potential of cannabis and cannabinoids. Dtsch. Arztebl. Int. 2012, 109, 495–501. [Google Scholar] [PubMed]
  10. Greenamyre, J.T.; Porter, R.H. Anatomy and physiology of glutamate in the CNS. Neurology 1994, 44, S7–S13. [Google Scholar] [PubMed]
  11. Meldrum, B.S. Glutamate as a neurotransmitter in the brain: Review of physiology and pathology. J. Nutr. 2000, 130, 1007S–1015S. [Google Scholar] [PubMed]
  12. Ziff, E.B. Recent excitement in the ionotropic glutamate receptor field. Ann. N. Y. Acad. Sci. 1999, 868, 465–473. [Google Scholar] [CrossRef] [PubMed]
  13. Brauner-Osborne, H.; Egebjerg, J.; Nielsen, E.O.; Madsen, U.; Krogsgaard-Larsen, P. Ligands for glutamate receptors: Design and therapeutic prospects. J. Med. Chem. 2000, 43, 2609–2645. [Google Scholar] [CrossRef] [PubMed]
  14. Olney, J.W.; Sharpe, L.G. Brain lesions in an infant rhesus monkey treated with monsodium glutamate. Science 1969, 166, 386–388. [Google Scholar] [CrossRef] [PubMed]
  15. Choi, D.W. Glutamate neurotoxicity in cortical cell culture is calcium dependent. Neurosci. Lett. 1985, 58, 293–297. [Google Scholar] [CrossRef]
  16. Farooqui, A.A.; Ong, W.Y.; Horrocks, L.A. Biochemical aspects of neurodegeneration in human brain: Involvement of neural membrane phospholipids and phospholipases A2. Neurochem. Res. 2004, 29, 1961–1977. [Google Scholar] [CrossRef] [PubMed]
  17. Gasparini, C.F.; Griffiths, L.R. The biology of the glutamatergic system and potential role in migraine. Int. J. Biomed. Sci. 2013, 9, 1–8. [Google Scholar] [PubMed]
  18. Cananzi, A.R.; D’Andrea, G.; Perini, F.; Zamberlan, F.; Welch, K.M. Platelet and plasma levels of glutamate and glutamine in migraine with and without aura. Cephalalgia 1995, 15, 132–135. [Google Scholar] [CrossRef] [PubMed]
  19. D’Eufemia, P.; Finocchiaro, R.; Lendvai, D.; Celli, M.; Viozzi, L.; Troiani, P.; Turri, E.; Giardini, O. Erythrocyte and plasma levels of glutamate and aspartate in children affected by migraine. Cephalalgia 1997, 17, 652–657. [Google Scholar] [CrossRef] [PubMed]
  20. Martinez, F.; Castillo, J.; Rodriguez, J.R.; Leira, R.; Noya, M. Neuroexcitatory amino acid levels in plasma and cerebrospinal fluid during migraine attacks. Cephalalgia 1993, 13, 89–93. [Google Scholar] [CrossRef] [PubMed]
  21. Edvinsson, L.; Villalon, C.M.; MaassenVanDenBrink, A. Basic mechanisms of migraine and its acute treatment. Pharmacol. Ther. 2012, 136, 319–333. [Google Scholar] [CrossRef] [PubMed]
  22. Latremoliere, A.; Woolf, C.J. Central sensitization: A generator of pain hypersensitivity by central neural plasticity. J. Pain 2009, 10, 895–926. [Google Scholar] [CrossRef] [PubMed]
  23. Ma, Q.P.; Woolf, C.J. Noxious stimuli induce an N-methyl-d-aspartate receptor-dependent hypersensitivity of the flexion withdrawal reflex to touch: Implications for the treatment of mechanical allodynia. Pain 1995, 61, 383–390. [Google Scholar] [CrossRef]
  24. Hughes, D.I.; Scott, D.T.; Todd, A.J.; Riddell, J.S. Lack of evidence for sprouting of abeta afferents into the superficial laminas of the spinal cord dorsal horn after nerve section. J. Neurosci. 2003, 23, 9491–9499. [Google Scholar] [PubMed]
  25. Soliman, A.C.; Yu, J.S.; Coderre, T.J. Mglu and NMDA receptor contributions to capsaicin-induced thermal and mechanical hypersensitivity. Neuropharmacology 2005, 48, 325–332. [Google Scholar] [CrossRef] [PubMed]
  26. D’Andrea, G.; Cananzi, A.R.; Joseph, R.; Morra, M.; Zamberlan, F.; Ferro Milone, F.; Grunfeld, S.; Welch, K.M. Platelet glycine, glutamate and aspartate in primary headache. Cephalalgia 1991, 11, 197–200. [Google Scholar] [CrossRef] [PubMed]
  27. Rothrock, J.F.; Mar, K.R.; Yaksh, T.L.; Golbeck, A.; Moore, A.C. Cerebrospinal fluid analyses in migraine patients and controls. Cephalalgia 1995, 15, 489–493. [Google Scholar] [CrossRef] [PubMed]
  28. Dykens, J.A.; Sullivan, S.G.; Stern, A. Oxidative reactivity of the tryptophan metabolites 3-hydroxyanthranilate, cinnabarinate, quinolinate and picolinate. Biochem. Pharmacol. 1987, 36, 211–217. [Google Scholar] [CrossRef]
  29. Birch, P.J.; Grossman, C.J.; Hayes, A.G. Kynurenate and FG9041 have both competitive and non-competitive antagonist actions at excitatory amino acid receptors. Eur. J. Pharmacol. 1988, 151, 313–315. [Google Scholar] [CrossRef]
  30. Perkins, M.N.; Stone, T.W. Actions of kynurenic acid and quinolinic acid in the rat hippocampus in vivo. Exp. Neurol. 1985, 88, 570–579. [Google Scholar] [CrossRef]
  31. Wang, J.; Simonavicius, N.; Wu, X.; Swaminath, G.; Reagan, J.; Tian, H.; Ling, L. Kynurenic acid as a ligand for orphan G protein-coupled receptor GPR35. J. Biol. Chem. 2006, 281, 22021–22028. [Google Scholar] [CrossRef] [PubMed]
  32. O’Dowd, B.F.; Nguyen, T.; Marchese, A.; Cheng, R.; Lynch, K.R.; Heng, H.H.; Kolakowski, L.F.; George, S.R. Discovery of three novel G-protein-coupled receptor genes. Genomics 1998, 47, 310–313. [Google Scholar] [CrossRef] [PubMed]
  33. Guo, J.; Williams, D.J.; Puhl, H.L.; Ikeda, S.R. Inhibition of n-type calcium channels by activation of GPR35, an orphan receptor, heterologously expressed in rat sympathetic neurons. J. Pharmacol. Exp. Ther. 2008, 324, 342–351. [Google Scholar] [CrossRef] [PubMed]
  34. Mackenzie, A.E.; Milligan, G. The emerging pharmacology and function of GPR35 in the nervous system. Neuropharmacology 2017, 113, 661–671. [Google Scholar] [CrossRef] [PubMed]
  35. Guidetti, P.; Schwarcz, R. 3-hydroxykynurenine potentiates quinolinate but not NMDA toxicity in the rat striatum. Eur. J. Neurosci. 1999, 11, 3857–3863. [Google Scholar] [CrossRef] [PubMed]
  36. Behan, W.M.; McDonald, M.; Darlington, L.G.; Stone, T.W. Oxidative stress as a mechanism for quinolinic acid-induced hippocampal damage: Protection by melatonin and deprenyl. Br. J. Pharmacol. 1999, 128, 1754–1760. [Google Scholar] [CrossRef] [PubMed]
  37. Tavares, R.G.; Tasca, C.I.; Santos, C.E.; Wajner, M.; Souza, D.O.; Dutra-Filho, C.S. Quinolinic acid inhibits glutamate uptake into synaptic vesicles from rat brain. NeuroReport 2000, 11, 249–253. [Google Scholar] [CrossRef] [PubMed]
  38. Kemp, J.A.; Foster, A.C.; Leeson, P.D.; Priestley, T.; Tridgett, R.; Iversen, L.L.; Woodruff, G.N. 7-chlorokynurenic acid is a selective antagonist at the glycine modulatory site of the N-methyl-d-aspartate receptor complex. Proc. Natl. Acad. Sci. USA 1988, 85, 6547–6550. [Google Scholar] [CrossRef] [PubMed]
  39. Sas, K.; Robotka, H.; Rozsa, E.; Agoston, M.; Szenasi, G.; Gigler, G.; Marosi, M.; Kis, Z.; Farkas, T.; Vecsei, L.; et al. Kynurenine diminishes the ischemia-induced histological and electrophysiological deficits in the rat hippocampus. Neurobiol. Dis. 2008, 32, 302–308. [Google Scholar] [CrossRef] [PubMed]
  40. Di Clemente, L.; Coppola, G.; Magis, D.; Gerardy, P.Y.; Fumal, A.; De Pasqua, V.; Di Piero, V.; Schoenen, J. Nitroglycerin sensitises in healthy subjects CNS structures involved in migraine pathophysiology: Evidence from a study of nociceptive blink reflexes and visual evoked potentials. Pain 2009, 144, 156–161. [Google Scholar] [CrossRef] [PubMed]
  41. Tassorelli, C.; Joseph, S.A. Systemic nitroglycerin induces Fos immunoreactivity in brainstem and forebrain structures of the rat. Brain Res. 1995, 682, 167–181. [Google Scholar] [CrossRef]
  42. Fejes-Szabo, A.; Bohar, Z.; Vamos, E.; Nagy-Grocz, G.; Tar, L.; Veres, G.; Zadori, D.; Szentirmai, M.; Tajti, J.; Szatmari, I.; et al. Pre-treatment with new kynurenic acid amide dose-dependently prevents the nitroglycerine-induced neuronal activation and sensitization in cervical part of trigemino-cervical complex. J. Neural Transm. (Vienna) 2014, 121, 725–738. [Google Scholar] [CrossRef] [PubMed]
  43. Vamos, E.; Pardutz, A.; Fejes, A.; Tajti, J.; Toldi, J.; Vecsei, L. Modulatory effects of probenecid on the nitroglycerin-induced changes in the rat caudal trigeminal nucleus. Eur. J. Pharmacol. 2009, 621, 33–37. [Google Scholar] [CrossRef] [PubMed]
  44. Vamos, E.; Pardutz, A.; Varga, H.; Bohar, Z.; Tajti, J.; Fulop, F.; Toldi, J.; Vecsei, L. l-kynurenine combined with probenecid and the novel synthetic kynurenic acid derivative attenuate nitroglycerin-induced nNOS in the rat caudal trigeminal nucleus. Neuropharmacology 2009, 57, 425–429. [Google Scholar] [CrossRef] [PubMed]
  45. Nagy-Grocz, G.; Tar, L.; Bohar, Z.; Fejes-Szabo, A.; Laborc, K.F.; Spekker, E.; Vecsei, L.; Pardutz, A. The modulatory effect of anandamide on nitroglycerin-induced sensitization in the trigeminal system of the rat. Cephalalgia 2016, 36, 849–861. [Google Scholar] [CrossRef] [PubMed]
  46. Nagy-Grócz, G.; Laborc, K.F.; Veres, G.; Bajtai, A.; Bohar, Z.; Zádori, D.; Fejes-Szabó, A.; Spekker, E.; Vécsei, L.; Párdutz, Á. The effect of systemic nitroglycerin administration on the kynurenine pathway in the rat. Front. Neurol. 2017, 8, 278. [Google Scholar] [CrossRef] [PubMed]
  47. Lukacs, M.; Warfvinge, K.; Kruse, L.S.; Tajti, J.; Fulop, F.; Toldi, J.; Vecsei, L.; Edvinsson, L. KYNA analogue SZR72 modifies CFA-induced dural inflammation- regarding expression of pERK1/2 and IL-1beta in the rat trigeminal ganglion. J. Headache Pain 2016, 17, 64. [Google Scholar] [CrossRef] [PubMed]
  48. Clavelou, P.; Pajot, J.; Dallel, R.; Raboisson, P. Application of the formalin test to the study of orofacial pain in the rat. Neurosci. Lett. 1989, 103, 349–353. [Google Scholar] [CrossRef]
  49. Veres, G.; Fejes-Szabo, A.; Zadori, D.; Nagy-Grocz, G.; Laszlo, A.M.; Bajtai, A.; Mandity, I.; Szentirmai, M.; Bohar, Z.; Laborc, K.; et al. A comparative assessment of two kynurenic acid analogs in the formalin model of trigeminal activation: A behavioral, immunohistochemical and pharmacokinetic study. J. Neural Transm. (Vienna) 2017, 124, 99–112. [Google Scholar] [CrossRef] [PubMed]
  50. Greco, R.; Demartini, C.; Zanaboni, A.M.; Redavide, E.; Pampalone, S.; Toldi, J.; Fulop, F.; Blandini, F.; Nappi, G.; Sandrini, G.; et al. Effects of kynurenic acid analogue 1 (KYNA-A1) in nitroglycerin-induced hyperalgesia: Targets and anti-migraine mechanisms. Cephalalgia 2016, in press. [Google Scholar] [CrossRef] [PubMed]
  51. Knyihar-Csillik, E.; Chadaide, Z.; Okuno, E.; Krisztin-Peva, B.; Toldi, J.; Varga, C.; Molnar, A.; Csillik, B.; Vecsei, L. Kynurenine aminotransferase in the supratentorial dura mater of the rat: Effect of stimulation of the trigeminal ganglion. Exp. Neurol. 2004, 186, 242–247. [Google Scholar] [CrossRef] [PubMed]
  52. Ayata, C. Cortical spreading depression triggers migraine attack: Pro. Headache 2010, 50, 725–730. [Google Scholar] [CrossRef] [PubMed]
  53. Lauritzen, M. Pathophysiology of the migraine aura. The spreading depression theory. Brain 1994, 117 Pt 1, 199–210. [Google Scholar] [CrossRef] [PubMed]
  54. Knapp, L.; Szita, B.; Kocsis, K.; Vecsei, L.; Toldi, J. Nitroglycerin enhances the propagation of cortical spreading depression: Comparative studies with sumatriptan and novel kynurenic acid analogues. Drug Des. Dev. Ther. 2017, 11, 27–34. [Google Scholar] [CrossRef] [PubMed]
  55. Lauritzen, M.; Rice, M.E.; Okada, Y.; Nicholson, C. Quisqualate, kainate and NMDA can initiate spreading depression in the turtle cerebellum. Brain Res. 1988, 475, 317–327. [Google Scholar] [CrossRef]
  56. Curto, M.; Lionetto, L.; Negro, A.; Capi, M.; Fazio, F.; Giamberardino, M.A.; Simmaco, M.; Nicoletti, F.; Martelletti, P. Altered kynurenine pathway metabolites in serum of chronic migraine patients. J. Headache Pain 2015, 17, 47. [Google Scholar] [CrossRef] [PubMed][Green Version]
  57. Curto, M.; Lionetto, L.; Negro, A.; Capi, M.; Perugino, F.; Fazio, F.; Giamberardino, M.A.; Simmaco, M.; Nicoletti, F.; Martelletti, P. Altered serum levels of kynurenine metabolites in patients affected by cluster headache. J. Headache Pain 2015, 17, 27. [Google Scholar] [CrossRef] [PubMed][Green Version]
  58. Sato, K.; Kiyama, H.; Park, H.T.; Tohyama, M. AMPA, KA and NMDA receptors are expressed in the rat DRG neurones. Neuroreport 1993, 4, 1263–1265. [Google Scholar] [CrossRef] [PubMed]
  59. Ohshiro, H.; Tonai-Kachi, H.; Ichikawa, K. GPR35 is a functional receptor in rat dorsal root ganglion neurons. Biochem. Biophys. Res. Commun. 2008, 365, 344–348. [Google Scholar] [CrossRef] [PubMed]
  60. Mecs, L.; Tuboly, G.; Nagy, E.; Benedek, G.; Horvath, G. The peripheral antinociceptive effects of endomorphin-1 and kynurenic acid in the rat inflamed joint model. Anesth. Analg. 2009, 109, 1297–1304. [Google Scholar] [CrossRef] [PubMed]
  61. Zhang, Y.Q.; Ji, G.C.; Wu, G.C.; Zhao, Z.Q. Kynurenic acid enhances electroacupuncture analgesia in normal and carrageenan-injected rats. Brain Res. 2003, 966, 300–307. [Google Scholar] [CrossRef]
  62. Matsuda, L.A.; Lolait, S.J.; Brownstein, M.J.; Young, A.C.; Bonner, T.I. Structure of a cannabinoid receptor and functional expression of the cloned cdna. Nature 1990, 346, 561–564. [Google Scholar] [CrossRef] [PubMed]
  63. Munro, S.; Thomas, K.L.; Abu-Shaar, M. Molecular characterization of a peripheral receptor for cannabinoids. Nature 1993, 365, 61–65. [Google Scholar] [CrossRef] [PubMed]
  64. Herkenham, M.; Lynn, A.B.; Little, M.D.; Johnson, M.R.; Melvin, L.S.; de Costa, B.R.; Rice, K.C. Cannabinoid receptor localization in brain. Proc. Natl. Acad. Sci. USA 1990, 87, 1932–1936. [Google Scholar] [CrossRef] [PubMed]
  65. Piomelli, D. The molecular logic of endocannabinoid signalling. Nat. Rev. Neurosci. 2003, 4, 873–884. [Google Scholar] [CrossRef] [PubMed]
  66. Maccarrone, M.; Bab, I.; Bíró, T.; Cabral, G.A.; Dey, S.K.; Di Marzo, V.; Konje, J.C.; Kunos, G.; Mechoulam, R.; Pacher, P.; et al. Endocannabinoid signaling at the periphery: 50 years after THC. Trends Pharmacol. Sci. 2015, 36, 277–296. [Google Scholar] [CrossRef] [PubMed]
  67. Pagotto, U.; Marsicano, G.; Cota, D.; Lutz, B.; Pasquali, R. The emerging role of the endocannabinoid system in endocrine regulation and energy balance. Endocr. Rev. 2006, 27, 73–100. [Google Scholar] [CrossRef] [PubMed]
  68. Klein, T.W. Cannabinoid-based drugs as anti-inflammatory therapeutics. Nat. Rev. Immunol. 2005, 5, 400–411. [Google Scholar] [CrossRef] [PubMed]
  69. Van Sickle, M.D.; Duncan, M.; Kingsley, P.J.; Mouihate, A.; Urbani, P.; Mackie, K.; Stella, N.; Makriyannis, A.; Piomelli, D.; Davison, J.S.; et al. Identification and functional characterization of brainstem cannabinoid CB2 receptors. Science 2005, 310, 329–332. [Google Scholar] [CrossRef] [PubMed]
  70. Pacher, P.; Haskó, G. Endocannabinoids and cannabinoid receptors in ischaemia-reperfusion injury and preconditioning. Br. J. Pharmacol. 2008, 153, 252–262. [Google Scholar] [CrossRef] [PubMed]
  71. Pertwee, R.G. Cannabinoid pharmacology: The first 66 years. Br. J. Pharmacol. 2006, 147 (Suppl. 1), S163–S171. [Google Scholar] [CrossRef] [PubMed]
  72. Malan, T.P.; Ibrahim, M.M.; Deng, H.; Liu, Q.; Mata, H.P.; Vanderah, T.; Porreca, F.; Makriyannis, A. CB2 cannabinoid receptor-mediated peripheral antinociception. Pain 2001, 93, 239–245. [Google Scholar] [CrossRef]
  73. Romero, T.R.; Resende, L.C.; Guzzo, L.S.; Duarte, I.D. CB1 and CB2 cannabinoid receptor agonists induce peripheral antinociception by activation of the endogenous noradrenergic system. Anesth. Analg. 2013, 116, 463–472. [Google Scholar] [CrossRef] [PubMed]
  74. Vincenzi, F.; Targa, M.; Corciulo, C.; Tabrizi, M.A.; Merighi, S.; Gessi, S.; Saponaro, G.; Baraldi, P.G.; Borea, P.A.; Varani, K. Antinociceptive effects of the selective CB2 agonist MT178 in inflammatory and chronic rodent pain models. Pain 2013, 154, 864–873. [Google Scholar] [CrossRef] [PubMed]
  75. Greco, R.; Mangione, A.S.; Sandrini, G.; Nappi, G.; Tassorelli, C. Activation of CB2 receptors as a potential therapeutic target for migraine: Evaluation in an animal model. J. Headache Pain 2014, 15, 14. [Google Scholar] [CrossRef] [PubMed]
  76. Devane, W.A.; Hanus, L.; Breuer, A.; Pertwee, R.G.; Stevenson, L.A.; Griffin, G.; Gibson, D.; Mandelbaum, A.; Etinger, A.; Mechoulam, R. Isolation and structure of a brain constituent that binds to the cannabinoid receptor. Science 1992, 258, 1946–1949. [Google Scholar] [CrossRef] [PubMed]
  77. Mechoulam, R.; Ben-Shabat, S.; Hanus, L.; Ligumsky, M.; Kaminski, N.E.; Schatz, A.R.; Gopher, A.; Almog, S.; Martin, B.R.; Compton, D.R. Identification of an endogenous 2-monoglyceride, present in canine gut, that binds to cannabinoid receptors. Biochem. Pharmacol. 1995, 50, 83–90. [Google Scholar] [CrossRef]
  78. Sugiura, T.; Kondo, S.; Sukagawa, A.; Nakane, S.; Shinoda, A.; Itoh, K.; Yamashita, A.; Waku, K. 2-arachidonoylglycerol: A possible endogenous cannabinoid receptor ligand in brain. Biochem. Biophys. Res. Commun. 1995, 215, 89–97. [Google Scholar] [CrossRef] [PubMed]
  79. Lu, H.C.; Mackie, K. An introduction to the endogenous cannabinoid system. Biol. Psychiatry 2016, 79, 516–525. [Google Scholar] [CrossRef] [PubMed]
  80. Cravatt, B.F.; Giang, D.K.; Mayfield, S.P.; Boger, D.L.; Lerner, R.A.; Gilula, N.B. Molecular characterization of an enzyme that degrades neuromodulatory fatty-acid amides. Nature 1996, 384, 83–87. [Google Scholar] [CrossRef] [PubMed]
  81. Mechoulam, R.; Fride, E.; Di Marzo, V. Endocannabinoids. Eur. J. Pharmacol. 1998, 359, 1–18. [Google Scholar] [CrossRef]
  82. Justinova, Z.; Panlilio, L.V.; Moreno-Sanz, G.; Redhi, G.H.; Auber, A.; Secci, M.E.; Mascia, P.; Bandiera, T.; Armirotti, A.; Bertorelli, R.; et al. Effects of fatty acid amide hydrolase (FAAH) inhibitors in non-human primate models of nicotine reward and relapse. Neuropsychopharmacology 2015, 40, 2185–2197. [Google Scholar] [CrossRef] [PubMed]
  83. Haller, V.L.; Stevens, D.L.; Welch, S.P. Modulation of opioids via protection of anandamide degradation by fatty acid amide hydrolase. Eur. J. Pharmacol. 2008, 600, 50–58. [Google Scholar] [CrossRef] [PubMed]
  84. Bushlin, I.; Rozenfeld, R.; Devi, L.A. Cannabinoid-opioid interactions during neuropathic pain and analgesia. Curr. Opin. Pharmacol. 2010, 10, 80–86. [Google Scholar] [CrossRef] [PubMed]
  85. Kondo, S.; Kondo, H.; Nakane, S.; Kodaka, T.; Tokumura, A.; Waku, K.; Sugiura, T. 2-arachidonoylglycerol, an endogenous cannabinoid receptor agonist: Identification as one of the major species of monoacylglycerols in various rat tissues, and evidence for its generation through Ca2+-dependent and -independent mechanisms. FEBS Lett. 1998, 429, 152–156. [Google Scholar] [CrossRef]
  86. Karlsson, M.; Contreras, J.A.; Hellman, U.; Tornqvist, H.; Holm, C. cDNA cloning, tissue distribution, and identification of the catalytic triad of monoglyceride lipase. Evolutionary relationship to esterases, lysophospholipases, and haloperoxidases. J. Biol. Chem. 1997, 272, 27218–27223. [Google Scholar] [CrossRef] [PubMed]
  87. Dinh, T.P.; Carpenter, D.; Leslie, F.M.; Freund, T.F.; Katona, I.; Sensi, S.L.; Kathuria, S.; Piomelli, D. Brain monoglyceride lipase participating in endocannabinoid inactivation. Proc. Natl. Acad. Sci. USA 2002, 99, 10819–10824. [Google Scholar] [CrossRef] [PubMed]
  88. Pertwee, R.G. Cannabinoid receptors and pain. Prog. Neurobiol. 2001, 63, 569–611. [Google Scholar] [CrossRef]
  89. Akerman, S.; Holland, P.R.; Goadsby, P.J. Cannabinoid (CB1) receptor activation inhibits trigeminovascular neurons. J. Pharmacol. Exp. Ther. 2007, 320, 64–71. [Google Scholar] [CrossRef] [PubMed]
  90. Cupini, L.M.; Costa, C.; Sarchielli, P.; Bari, M.; Battista, N.; Eusebi, P.; Calabresi, P.; Maccarrone, M. Degradation of endocannabinoids in chronic migraine and medication overuse headache. Neurobiol. Dis. 2008, 30, 186–189. [Google Scholar] [CrossRef] [PubMed]
  91. Rossi, C.; Pini, L.A.; Cupini, M.L.; Calabresi, P.; Sarchielli, P. Endocannabinoids in platelets of chronic migraine patients and medication-overuse headache patients: Relation with serotonin levels. Eur. J. Clin. Pharmacol. 2008, 64, 1–8. [Google Scholar] [CrossRef] [PubMed]
  92. Sarchielli, P.; Pini, L.A.; Coppola, F.; Rossi, C.; Baldi, A.; Mancini, M.L.; Calabresi, P. Endocannabinoids in chronic migraine: CSF findings suggest a system failure. Neuropsychopharmacology 2007, 32, 1384–1390. [Google Scholar] [CrossRef] [PubMed]
  93. Cupini, L.M.; Bari, M.; Battista, N.; Argiro, G.; Finazzi-Agro, A.; Calabresi, P.; Maccarrone, M. Biochemical changes in endocannabinoid system are expressed in platelets of female but not male migraineurs. Cephalalgia 2006, 26, 277–281. [Google Scholar] [CrossRef] [PubMed]
  94. Greco, R.; Gasperi, V.; Maccarrone, M.; Tassorelli, C. The endocannabinoid system and migraine. Exp. Neurol. 2010, 224, 85–91. [Google Scholar] [CrossRef] [PubMed]
  95. Greco, R.; Gasperi, V.; Sandrini, G.; Bagetta, G.; Nappi, G.; Maccarrone, M.; Tassorelli, C. Alterations of the endocannabinoid system in an animal model of migraine: Evaluation in cerebral areas of rat. Cephalalgia 2010, 30, 296–302. [Google Scholar] [CrossRef] [PubMed]
  96. Akerman, S.; Kaube, H.; Goadsby, P.J. Anandamide is able to inhibit trigeminal neurons using an in vivo model of trigeminovascular-mediated nociception. J. Pharmacol. Exp. Ther. 2004, 309, 56–63. [Google Scholar] [CrossRef] [PubMed]
  97. Nozaki, S.; Endo, Y.; Nakahara, H.; Yoshizawa, K.; Hashiba, Y.; Kawashiri, S.; Tanaka, A.; Nakagawa, K.; Matsuoka, Y.; Kogo, M.; et al. Inhibition of invasion and metastasis in oral cancer by targeting urokinase-type plasminogen activator receptor. Oral Oncol. 2005, 41, 971–977. [Google Scholar] [CrossRef] [PubMed]
  98. Kazemi, H.; Rahgozar, M.; Speckmann, E.J.; Gorji, A. Effect of cannabinoid receptor activation on spreading depression. Iran. J. Basic Med. Sci. 2012, 15, 926–936. [Google Scholar] [PubMed]
  99. Akerman, S.; Holland, P.R.; Lasalandra, M.P.; Goadsby, P.J. Endocannabinoids in the brainstem modulate dural trigeminovascular nociceptive traffic via CB1 and “triptan” receptors: Implications in migraine. J. Neurosci. 2013, 33, 14869–14877. [Google Scholar] [CrossRef] [PubMed]
  100. Goadsby, P.J.; Edvinsson, L.; Ekman, R. Release of vasoactive peptides in the extracerebral circulation of humans and the cat during activation of the trigeminovascular system. Ann. Neurol. 1988, 23, 193–196. [Google Scholar] [CrossRef] [PubMed]
  101. Guindon, J.; Desroches, J.; Beaulieu, P. The antinociceptive effects of intraplantar injections of 2-arachidonoyl glycerol are mediated by cannabinoid CB2 receptors. Br. J. Pharmacol. 2007, 150, 693–701. [Google Scholar] [CrossRef] [PubMed]
  102. La Rana, G.; Russo, R.; Campolongo, P.; Bortolato, M.; Mangieri, R.A.; Cuomo, V.; Iacono, A.; Raso, G.M.; Meli, R.; Piomelli, D.; et al. Modulation of neuropathic and inflammatory pain by the endocannabinoid transport inhibitor am404 [N-(4-hydroxyphenyl)-eicosa-5,8,11,14-tetraenamide]. J. Pharmacol. Exp. Ther. 2006, 317, 1365–1371. [Google Scholar] [CrossRef] [PubMed]
  103. Strangman, N.M.; Walker, J.M. Cannabinoid win 55,212-2 inhibits the activity-dependent facilitation of spinal nociceptive responses. J. Neurophysiol. 1999, 82, 472–477. [Google Scholar] [PubMed]
  104. Ahn, K.; Johnson, D.S.; Mileni, M.; Beidler, D.; Long, J.Z.; McKinney, M.K.; Weerapana, E.; Sadagopan, N.; Liimatta, M.; Smith, S.E.; et al. Discovery and characterization of a highly selective FAAH inhibitor that reduces inflammatory pain. Chem. Biol. 2009, 16, 411–420. [Google Scholar] [CrossRef] [PubMed]
  105. Glass, M.; Dragunow, M.; Faull, R.L. Cannabinoid receptors in the human brain: A detailed anatomical and quantitative autoradiographic study in the fetal, neonatal and adult human brain. Neuroscience 1997, 77, 299–318. [Google Scholar] [CrossRef]
  106. Thomas, B.F.; Wei, X.; Martin, B.R. Characterization and autoradiographic localization of the cannabinoid binding site in rat brain using [3H] 11-OH-delta 9-THC-DMH. J. Pharmacol. Exp. Ther. 1992, 263, 1383–1390. [Google Scholar] [PubMed]
  107. Wotherspoon, G.; Fox, A.; McIntyre, P.; Colley, S.; Bevan, S.; Winter, J. Peripheral nerve injury induces cannabinoid receptor 2 protein expression in rat sensory neurons. Neuroscience 2005, 135, 235–245. [Google Scholar] [CrossRef] [PubMed]
  108. Colin-Gonzalez, A.L.; Aguilera, G.; Santamaria, A. Cannabinoids: Glutamatergic transmission and kynurenines. Adv. Neurobiol. 2016, 12, 173–198. [Google Scholar] [PubMed]
  109. Juhasz, G.; Lazary, J.; Chase, D.; Pegg, E.; Downey, D.; Toth, Z.G.; Stones, K.; Platt, H.; Mekli, K.; Payton, A.; et al. Variations in the cannabinoid receptor 1 gene predispose to migraine. Neurosci. Lett. 2009, 461, 116–120. [Google Scholar] [CrossRef] [PubMed]
  110. Tallaksen-Greene, S.J.; Young, A.B.; Penney, J.B.; Beitz, A.J. Excitatory amino acid binding sites in the trigeminal principal sensory and spinal trigeminal nuclei of the rat. Neurosci. Lett. 1992, 141, 79–83. [Google Scholar] [CrossRef]
  111. Watanabe, M.; Mishina, M.; Inoue, Y. Distinct gene expression of the N-methyl-d-aspartate receptor channel subunit in peripheral neurons of the mouse sensory ganglia and adrenal gland. Neurosci. Lett. 1994, 165, 183–186. [Google Scholar] [CrossRef]
  112. Furuyama, T.; Kiyama, H.; Sato, K.; Park, H.T.; Maeno, H.; Takagi, H.; Tohyama, M. Region-specific expression of subunits of ionotropic glutamate receptors (AMPA-type, KA-type and NMDA receptors) in the rat spinal cord with special reference to nociception. Brain Res. Mol. Brain Res. 1993, 18, 141–151. [Google Scholar] [CrossRef]
  113. Lomazzo, E.; Bindila, L.; Remmers, F.; Lerner, R.; Schwitter, C.; Hoheisel, U.; Lutz, B. Therapeutic potential of inhibitors of endocannabinoid degradation for the treatment of stress-related hyperalgesia in an animal model of chronic pain. Neuropsychopharmacology 2015, 40, 488–501. [Google Scholar] [CrossRef] [PubMed]
  114. Martin, G.; Nie, Z.; Siggins, G.R. Mu-opioid receptors modulate NMDA receptor-mediated responses in nucleus accumbens neurons. J. Neurosci. 1997, 17, 11–22. [Google Scholar] [PubMed]
  115. Patel, S.; Hill, M.N.; Cheer, J.F.; Wotjak, C.T.; Holmes, A. The endocannabinoid system as a target for novel anxiolytic drugs. Neurosci. Biobehav. Rev. 2017, 76, 56–66. [Google Scholar] [CrossRef] [PubMed]
  116. Vecsei, L.; Szalardy, L.; Fulop, F.; Toldi, J. Kynurenines in the CNS: Recent advances and new questions. Nat. Rev. Drug Discov. 2013, 12, 64–82. [Google Scholar] [CrossRef] [PubMed]
  117. Goadsby, P.J.; Holland, P.R.; Martins-Oliveira, M.; Hoffmann, J.; Schankin, C.; Akerman, S. Pathophysiology of migraine: A disorder of sensory processing. Physiol. Rev. 2017, 97, 553–622. [Google Scholar] [CrossRef] [PubMed]
  118. Basavarajappa, B.S. Critical enzymes involved in endocannabinoid metabolism. Protein Pept. Lett. 2007, 14, 237–246. [Google Scholar] [CrossRef] [PubMed]
  119. Guillemin, G.J.; Cullen, K.M.; Lim, C.K.; Smythe, G.A.; Garner, B.; Kapoor, V.; Takikawa, O.; Brew, B.J. Characterization of the kynurenine pathway in human neurons. J. Neurosci. 2007, 27, 12884–12892. [Google Scholar] [CrossRef] [PubMed]
  120. Jenny, M.; Santer, E.; Pirich, E.; Schennach, H.; Fuchs, D. Delta9-tetrahydrocannabinol and cannabidiol modulate mitogen-induced tryptophan degradation and neopterin formation in peripheral blood mononuclear cells in vitro. J. Neuroimmunol. 2009, 207, 75–82. [Google Scholar] [CrossRef] [PubMed]
  121. Aloyo, V.J.; Berg, K.A.; Clarke, W.P.; Spampinato, U.; Harvey, J.A. Inverse agonism at serotonin and cannabinoid receptors. Prog. Mol. Biol. Transl. Sci. 2010, 91, 1–40. [Google Scholar] [PubMed]
  122. Devlin, M.G.; Christopoulos, A. Modulation of cannabinoid agonist binding by 5-HT in the rat cerebellum. J. Neurochem. 2002, 80, 1095–1102. [Google Scholar] [CrossRef] [PubMed]
  123. Justinova, Z.; Mascia, P.; Wu, H.Q.; Secci, M.E.; Redhi, G.H.; Panlilio, L.V.; Scherma, M.; Barnes, C.; Parashos, A.; Zara, T.; et al. Reducing cannabinoid abuse and preventing relapse by enhancing endogenous brain levels of kynurenic acid. Nat. Neurosci. 2013, 16, 1652–1661. [Google Scholar] [CrossRef] [PubMed][Green Version]
  124. Rodríguez-Muñoz, M.; Sánchez-Blázquez, P.; Merlos, M.; Garzón-Niño, J. Endocannabinoid control of glutamate NMDA receptors: The therapeutic potential and consequences of dysfunction. Oncotarget 2016, 7, 55840–55862. [Google Scholar] [CrossRef] [PubMed]
  125. Stone, T.W.; Darlington, L.G. Endogenous kynurenines as targets for drug discovery and development. Nat. Rev. Drug Discov. 2002, 1, 609–620. [Google Scholar] [CrossRef] [PubMed]
  126. Ahluwalia, J.; Urban, L.; Capogna, M.; Bevan, S.; Nagy, I. Cannabinoid 1 receptors are expressed in nociceptive primary sensory neurons. Neuroscience 2000, 100, 685–688. [Google Scholar] [CrossRef]
  127. Turski, M.P.; Turska, M.; Paluszkiewicz, P.; Parada-Turska, J.; Oxenkrug, G.F. Kynurenic acid in the digestive system-new facts, new challenges. Int. J. Tryptophan Res. 2013, 6, 47–55. [Google Scholar] [PubMed]
  128. Perkins, M.N.; Stone, T.W. An iontophoretic investigation of the actions of convulsant kynurenines and their interaction with the endogenous excitant quinolinic acid. Brain Res. 1982, 247, 184–187. [Google Scholar] [CrossRef]
  129. Newcomer, J.W.; Krystal, J.H. NMDA receptor regulation of memory and behavior in humans. Hippocampus 2001, 11, 529–542. [Google Scholar] [CrossRef] [PubMed]
  130. Robinson, L.; McKillop-Smith, S.; Ross, N.L.; Pertwee, R.G.; Hampson, R.E.; Platt, B.; Riedel, G. Hippocampal endocannabinoids inhibit spatial learning and limit spatial memory in rats. Psychopharmacology 2008, 198, 551–563. [Google Scholar] [CrossRef] [PubMed]
  131. Wise, L.E.; Iredale, P.A.; Lichtman, A.H. The cannabinoid CB (1) receptor antagonist CE prolongs spatial memory duration in a rat delayed radial arm maze memory task. Eur. J. Pharmacol. 2008, 590, 246–249. [Google Scholar] [CrossRef] [PubMed]
  132. Allen, R.M.; Carelli, R.M.; Dykstra, L.A.; Suchey, T.L.; Everett, C.V. Effects of the competitive N-methyl-d-aspartate receptor antagonist, ly235959 [(−)-6-phosphonomethyl-deca-hydroisoquinoline-3-carboxylic acid], on responding for cocaine under both fixed and progressive ratio schedules of reinforcement. J. Pharmacol. Exp. Ther. 2005, 315, 449–457. [Google Scholar] [CrossRef] [PubMed]
  133. Allen, R.M.; Uban, K.A.; Atwood, E.M.; Albeck, D.S.; Yamamoto, D.J. Continuous intracerebroventricular infusion of the competitive NMDA receptor antagonist, LY235959, facilitates escalation of cocaine self-administration and increases break point for cocaine in sprague-dawley rats. Pharmacol. Biochem. Behav. 2007, 88, 82–88. [Google Scholar] [CrossRef] [PubMed]
  134. Soria, G.; Mendizábal, V.; Touriño, C.; Robledo, P.; Ledent, C.; Parmentier, M.; Maldonado, R.; Valverde, O. Lack of CB1 cannabinoid receptor impairs cocaine self-administration. Neuropsychopharmacology 2005, 30, 1670–1680. [Google Scholar] [CrossRef] [PubMed]
  135. De Vry, J.; Denzer, D.; Reissmueller, E.; Eijckenboom, M.; Heil, M.; Meier, H.; Mauler, F. 3-[2-cyano-3-(trifluoromethyl)phenoxy]phenyl-4,4,4-trifluoro-1-butanesulfonate (BAY 59-3074): A novel cannabinoid CB1/CB2 receptor partial agonist with antihyperalgesic and antiallodynic effects. J. Pharmacol. Exp. Ther. 2004, 310, 620–632. [Google Scholar] [CrossRef] [PubMed]
  136. Pelissier, T.; Infante, C.; Constandil, L.; Espinosa, J.; Lapeyra, C.D.; Hernández, A. Antinociceptive effect and interaction of uncompetitive and competitive NMDA receptor antagonists upon capsaicin and paw pressure testing in normal and monoarthritic rats. Pain 2008, 134, 113–127. [Google Scholar] [CrossRef] [PubMed]
  137. Sánchez-Blázquez, P.; Rodríguez-Muñoz, M.; Garzón, J. The cannabinoid receptor 1 associates with NMDA receptors to produce glutamatergic hypofunction: Implications in psychosis and schizophrenia. Front. Pharmacol. 2014, 4, 169. [Google Scholar] [CrossRef] [PubMed]
  138. Corlew, R.; Brasier, D.J.; Feldman, D.E.; Philpot, B.D. Presynaptic NMDA receptors: Newly appreciated roles in cortical synaptic function and plasticity. Neuroscientist 2008, 14, 609–625. [Google Scholar] [CrossRef] [PubMed]
  139. Hohmann, A.G.; Briley, E.M.; Herkenham, M. Pre- and postsynaptic distribution of cannabinoid and mu opioid receptors in rat spinal cord. Brain Res. 1999, 822, 17–25. [Google Scholar] [CrossRef]
  140. Marchalant, Y.; Cerbai, F.; Brothers, H.M.; Wenk, G.L. Cannabinoid receptor stimulation is anti-inflammatory and improves memory in old rats. Neurobiol. Aging 2008, 29, 1894–1901. [Google Scholar] [CrossRef] [PubMed]
  141. Salio, C.; Fischer, J.; Franzoni, M.F.; Conrath, M. Pre- and postsynaptic localizations of the CB1 cannabinoid receptor in the dorsal horn of the rat spinal cord. Neuroscience 2002, 110, 755–764. [Google Scholar] [CrossRef]
  142. Sánchez-Blázquez, P.; Rodríguez-Muñoz, M.; Vicente-Sánchez, A.; Garzón, J. Cannabinoid receptors couple to NMDA receptors to reduce the production of no and the mobilization of zinc induced by glutamate. Antioxid. Redox Signal 2013, 19, 1766–1782. [Google Scholar] [CrossRef] [PubMed]
  143. Liu, X.J.; Salter, M.W. Glutamate receptor phosphorylation and trafficking in pain plasticity in spinal cord dorsal horn. Eur. J. Neurosci. 2010, 32, 278–289. [Google Scholar] [CrossRef] [PubMed]
  144. Shyu, Y.J.; Suarez, C.D.; Hu, C.D. Visualization of ternary complexes in living cells by using a BiFC-based FRET assay. Nat. Protoc. 2008, 3, 1693–1702. [Google Scholar] [CrossRef] [PubMed]
  145. Fiorentini, C.; Gardoni, F.; Spano, P.; Di Luca, M.; Missale, C. Regulation of dopamine D1 receptor trafficking and desensitization by oligomerization with glutamate N-methyl-d-aspartate receptors. J. Biol. Chem. 2003, 278, 20196–20202. [Google Scholar] [CrossRef] [PubMed]
  146. Perroy, J.; Raynaud, F.; Homburger, V.; Rousset, M.C.; Telley, L.; Bockaert, J.; Fagni, L. Direct interaction enables cross-talk between ionotropic and group I metabotropic glutamate receptors. J. Biol. Chem. 2008, 283, 6799–6805. [Google Scholar] [CrossRef] [PubMed]
  147. Rodríguez-Muñoz, M.; Sánchez-Blázquez, P.; Vicente-Sánchez, A.; Berrocoso, E.; Garzón, J. The mu-opioid receptor and the NMDA receptor associate in pag neurons: Implications in pain control. Neuropsychopharmacology 2012, 37, 338–349. [Google Scholar] [CrossRef] [PubMed]
  148. Sánchez-Blázquez, P.; Rodríguez-Muñoz, M.; Herrero-Labrador, R.; Burgueño, J.; Zamanillo, D.; Garzón, J. The calcium-sensitive sigma-1 receptor prevents cannabinoids from provoking glutamate NMDA receptor hypofunction: Implications in antinociception and psychotic diseases. Int. J. Neuropsychopharmacol. 2014, 17, 1943–1955. [Google Scholar] [CrossRef] [PubMed]
  149. Rodríguez-Muñoz, M.; Cortés-Montero, E.; Pozo-Rodrigálvarez, A.; Sánchez-Blázquez, P.; Garzón-Niño, J. The ON:Off switch, σ1R-HINT1 protein, controls GPCR-NMDA receptor cross-regulation: Implications in neurological disorders. Oncotarget 2015, 6, 35458–35477. [Google Scholar] [PubMed]
  150. Rodríguez-Muñoz, M.; Sánchez-Blázquez, P.; Herrero-Labrador, R.; Martínez-Murillo, R.; Merlos, M.; Vela, J.M.; Garzón, J. The σ1 receptor engages the redox-regulated HINT1 protein to bring opioid analgesia under NMDA receptor negative control. Antioxid. Redox Signal 2015, 22, 799–818. [Google Scholar] [CrossRef] [PubMed]
  151. Khaspekov, L.G.; Brenz Verca, M.S.; Frumkina, L.E.; Hermann, H.; Marsicano, G.; Lutz, B. Involvement of brain-derived neurotrophic factor in cannabinoid receptor-dependent protection against excitotoxicity. Eur. J. Neurosci. 2004, 19, 1691–1698. [Google Scholar] [CrossRef] [PubMed]
  152. Liu, Q.; Bhat, M.; Bowen, W.D.; Cheng, J. Signaling pathways from cannabinoid receptor-1 activation to inhibition of N-methyl-d-aspartic acid mediated calcium influx and neurotoxicity in dorsal root ganglion neurons. J. Pharmacol. Exp. Ther. 2009, 331, 1062–1070. [Google Scholar] [CrossRef] [PubMed]
  153. Vicente-Sánchez, A.; Sánchez-Blázquez, P.; Rodríguez-Muñoz, M.; Garzón, J. HINT1 protein cooperates with cannabinoid 1 receptor to negatively regulate glutamate NMDA receptor activity. Mol. Brain 2013, 6, 42. [Google Scholar] [CrossRef] [PubMed]
  154. Fan, N.; Yang, H.; Zhang, J.; Chen, C. Reduced expression of glutamate receptors and phosphorylation of CREB are responsible for in vivo Delta9-THC exposure-impaired hippocampal synaptic plasticity. J. Neurochem. 2010, 112, 691–702. [Google Scholar] [CrossRef] [PubMed]
  155. Ghasemi, M.; Schachter, S.C. The NMDA receptor complex as a therapeutic target in epilepsy: A review. Epilepsy Behav. 2011, 22, 617–640. [Google Scholar] [CrossRef] [PubMed]
  156. Degenhardt, L.; Hall, W.; Lynskey, M. Testing hypotheses about the relationship between cannabis use and psychosis. Drug Alcohol Depend. 2003, 71, 37–48. [Google Scholar] [CrossRef]
  157. Fernandez-Espejo, E.; Viveros, M.P.; Núñez, L.; Ellenbroek, B.A.; Rodriguez de Fonseca, F. Role of cannabis and endocannabinoids in the genesis of schizophrenia. Psychopharmacology 2009, 206, 531–549. [Google Scholar] [CrossRef] [PubMed]
  158. Altar, C.A.; Vawter, M.P.; Ginsberg, S.D. Target identification for CNS diseases by transcriptional profiling. Neuropsychopharmacology 2009, 34, 18–54. [Google Scholar] [CrossRef] [PubMed]
  159. Chen, Q.; Wang, X.; O’Neill, F.A.; Walsh, D.; Kendler, K.S.; Chen, X. Is the histidine triad nucleotide-binding protein 1 (HINT1) gene a candidate for schizophrenia? Schizophr. Res. 2008, 106, 200–207. [Google Scholar] [CrossRef] [PubMed]
  160. Watanabe, Y.; Nunokawa, A.; Kaneko, N.; Shibuya, M.; Egawa, J.; Someya, T. Supportive evidence for the association between the Gln2Pro polymorphism in the SIGMAR1 gene and schizophrenia in the Japanese population: A case-control study and an updated meta-analysis. Schizophr. Res. 2012, 141, 279–280. [Google Scholar] [CrossRef] [PubMed]
  161. Abadias, M.; Escriche, M.; Vaqué, A.; Sust, M.; Encina, G. Safety, tolerability and pharmacokinetics of single and multiple doses of a novel sigma-1 receptor antagonist in three randomized phase I studies. Br. J. Clin. Pharmacol. 2013, 75, 103–117. [Google Scholar] [CrossRef] [PubMed]
  162. Ritsner, M.S.; Gibel, A.; Shleifer, T.; Boguslavsky, I.; Zayed, A.; Maayan, R.; Weizman, A.; Lerner, V. Pregnenolone and dehydroepiandrosterone as an adjunctive treatment in schizophrenia and schizoaffective disorder: An 8-week, double-blind, randomized, controlled, 2-center, parallel-group trial. J. Clin. Psychiatry 2010, 71, 1351–1362. [Google Scholar] [CrossRef] [PubMed]
  163. Horvath, G.; Kekesi, G.; Tuboly, G.; Benedek, G. Antinociceptive interactions of triple and quadruple combinations of endogenous ligands at the spinal level. Brain Res. 2007, 1155, 42–48. [Google Scholar] [CrossRef] [PubMed]
  164. Kekesi, G.; Joo, G.; Csullog, E.; Dobos, I.; Klimscha, W.; Toth, K.; Benedek, G.; Horvath, G. The antinociceptive effect of intrathecal kynurenic acid and its interaction with endomorphin-1 in rats. Eur. J. Pharmacol. 2002, 445, 93–96. [Google Scholar] [CrossRef]
  165. Marek, P.; Ben-Eliyahu, S.; Gold, M.; Liebeskind, J.C. Excitatory amino acid antagonists (kynurenic acid and MK-801) attenuate the development of morphine tolerance in the rat. Brain Res. 1991, 547, 77–81. [Google Scholar] [CrossRef]
  166. Morgan, M.M.; Bobeck, E.N.; Ingram, S.L. Glutamate modulation of antinociception, but not tolerance, produced by morphine microinjection into the periaqueductal gray of the rat. Brain Res. 2009, 1295, 59–66. [Google Scholar] [CrossRef] [PubMed]
  167. Safrany-Fark, A.; Petrovszki, Z.; Kekesi, G.; Keresztes, C.; Benedek, G.; Horvath, G. Telemetry monitoring for non-invasive assessment of changes in core temperature after spinal drug administration in freely moving rats. J. Pharmacol. Toxicol. Methods 2015, 72, 19–25. [Google Scholar] [CrossRef] [PubMed]
  168. Zador, F.; Samavati, R.; Szlavicz, E.; Tuka, B.; Bojnik, E.; Fulop, F.; Toldi, J.; Vecsei, L.; Borsodi, A. Inhibition of opioid receptor mediated g-protein activity after chronic administration of kynurenic acid and its derivative without direct binding to opioid receptors. CNS Neurol. Disord. Drug Targets 2014, 13, 1520–1529. [Google Scholar] [CrossRef] [PubMed]
  169. Samavati, R.; Zádor, F.; Szűcs, E.; Tuka, B.; Martos, D.; Veres, G.; Gáspár, R.; Mándity, I.M.; Fülöp, F.; Vécsei, L.; et al. Kynurenic acid and its analogue can alter the opioid receptor g-protein signaling after acute treatment via NMDA receptor in rat cortex and striatum. J. Neurol. Sci. 2017, 376, 63–70. [Google Scholar] [CrossRef] [PubMed]
  170. Cichewicz, D.L. Synergistic interactions between cannabinoid and opioid analgesics. Life Sci. 2004, 74, 1317–1324. [Google Scholar] [CrossRef] [PubMed]
  171. Desroches, J.; Beaulieu, P. Opioids and cannabinoids interactions: Involvement in pain management. Curr. Drug Targets 2010, 11, 462–473. [Google Scholar] [CrossRef] [PubMed]
  172. Maldonado, R.; Valverde, O. Participation of the opioid system in cannabinoid-induced antinociception and emotional-like responses. Eur. Neuropsychopharmacol. 2003, 13, 401–410. [Google Scholar] [CrossRef] [PubMed]
  173. Mao, J. NMDA and opioid receptors: Their interactions in antinociception, tolerance and neuroplasticity. Brain Res. Brain Res. Rev. 1999, 30, 289–304. [Google Scholar] [CrossRef]
  174. Robledo, P.; Berrendero, F.; Ozaita, A.; Maldonado, R. Advances in the field of cannabinoid—Opioid cross-talk. Addict. Biol. 2008, 13, 213–224. [Google Scholar] [CrossRef] [PubMed]
  175. Scavone, J.L.; Sterling, R.C.; Van Bockstaele, E.J. Cannabinoid and opioid interactions: Implications for opiate dependence and withdrawal. Neuroscience 2013, 248, 637–654. [Google Scholar] [CrossRef] [PubMed]
  176. Zádor, F.; Wollemann, M. Receptome: Interactions between three pain-related receptors or the “triumvirate” of cannabinoid, opioid and TRPV1 receptors. Pharmacol. Res. 2015, 102, 254–263. [Google Scholar] [CrossRef] [PubMed]
  177. Scherma, M.; Muntoni, A.L.; Melis, M.; Fattore, L.; Fadda, P.; Fratta, W.; Pistis, M. Interactions between the endocannabinoid and nicotinic cholinergic systems: Preclinical evidence and therapeutic perspectives. Psychopharmacology 2016, 233, 1765–1777. [Google Scholar] [CrossRef] [PubMed]
  178. Hilmas, C.; Pereira, E.F.; Alkondon, M.; Rassoulpour, A.; Schwarcz, R.; Albuquerque, E.X. The brain metabolite kynurenic acid inhibits alpha7 nicotinic receptor activity and increases non-alpha7 nicotinic receptor expression: Physiopathological implications. J. Neurosci. 2001, 21, 7463–7473. [Google Scholar] [PubMed]
  179. Albuquerque, E.X.; Schwarcz, R. Kynurenic acid as an antagonist of α7 nicotinic acetylcholine receptors in the brain: Facts and challenges. Biochem. Pharmacol. 2013, 85, 1027–1032. [Google Scholar] [CrossRef] [PubMed]
  180. Dobelis, P.; Staley, K.J.; Cooper, D.C. Lack of modulation of nicotinic acetylcholine alpha-7 receptor currents by kynurenic acid in adult hippocampal interneurons. PLoS ONE 2012, 7, e41108. [Google Scholar] [CrossRef] [PubMed]
  181. Mok, M.H.; Fricker, A.C.; Weil, A.; Kew, J.N. Electrophysiological characterisation of the actions of kynurenic acid at ligand-gated ion channels. Neuropharmacology 2009, 57, 242–249. [Google Scholar] [CrossRef] [PubMed]
  182. Dani, J.A.; Bertrand, D. Nicotinic acetylcholine receptors and nicotinic cholinergic mechanisms of the central nervous system. Annu. Rev. Pharmacol. Toxicol. 2007, 47, 699–729. [Google Scholar] [CrossRef] [PubMed]
  183. Gotti, C.; Clementi, F.; Fornari, A.; Gaimarri, A.; Guiducci, S.; Manfredi, I.; Moretti, M.; Pedrazzi, P.; Pucci, L.; Zoli, M. Structural and functional diversity of native brain neuronal nicotinic receptors. Biochem. Pharmacol. 2009, 78, 703–711. [Google Scholar] [CrossRef] [PubMed]
  184. Freund, T.F.; Katona, I.; Piomelli, D. Role of endogenous cannabinoids in synaptic signaling. Physiol. Rev. 2003, 83, 1017–1066. [Google Scholar] [CrossRef] [PubMed]
  185. Wonnacott, S. Presynaptic nicotinic ACh receptors. Trends Neurosci. 1997, 20, 92–98. [Google Scholar] [CrossRef]
  186. Solinas, M.; Scherma, M.; Fattore, L.; Stroik, J.; Wertheim, C.; Tanda, G.; Fratta, W.; Goldberg, S.R. Nicotinic alpha 7 receptors as a new target for treatment of cannabis abuse. J. Neurosci. 2007, 27, 5615–5620. [Google Scholar] [CrossRef] [PubMed]
  187. Jenkins, L.; Brea, J.; Smith, N.J.; Hudson, B.D.; Reilly, G.; Bryant, N.J.; Castro, M.; Loza, M.I.; Milligan, G. Identification of novel species-selective agonists of the G-protein-coupled receptor GPR35 that promote recruitment of β-arrestin-2 and activate Gα13. Biochem. J. 2010, 432, 451–459. [Google Scholar] [CrossRef] [PubMed]
  188. Jenkins, L.; Alvarez-Curto, E.; Campbell, K.; de Munnik, S.; Canals, M.; Schlyer, S.; Milligan, G. Agonist activation of the g protein-coupled receptor GPR35 involves transmembrane domain III and is transduced via Gα₁₃ and β-arrestin-2. Br. J. Pharmacol. 2011, 162, 733–748. [Google Scholar] [CrossRef] [PubMed]
  189. Fallarini, S.; Magliulo, L.; Paoletti, T.; de Lalla, C.; Lombardi, G. Expression of functional GPR35 in human iNKT cells. Biochem. Biophys. Res. Commun. 2010, 398, 420–425. [Google Scholar] [CrossRef] [PubMed]
  190. Mackenzie, A.E.; Lappin, J.E.; Taylor, D.L.; Nicklin, S.A.; Milligan, G. GPR35 as a novel therapeutic target. Front. Endocrinol. (Lausanne) 2011, 2, 68. [Google Scholar] [CrossRef] [PubMed]
  191. Shore, D.M.; Reggio, P.H. The therapeutic potential of orphan GPCRS, GPR35 and GPR55. Front. Pharmacol. 2015, 6, 69. [Google Scholar] [CrossRef] [PubMed]
  192. Oka, S.; Ota, R.; Shima, M.; Yamashita, A.; Sugiura, T. GPR35 is a novel lysophosphatidic acid receptor. Biochem. Biophys. Res. Commun. 2010, 395, 232–237. [Google Scholar] [CrossRef] [PubMed]
  193. Walker, J.M.; Hohmann, A.G.; Martin, W.J.; Strangman, N.M.; Huang, S.M.; Tsou, K. The neurobiology of cannabinoid analgesia. Life Sci. 1999, 65, 665–673. [Google Scholar] [CrossRef]
  194. Taniguchi, Y.; Tonai-Kachi, H.; Shinjo, K. Zaprinast, a well-known cyclic guanosine monophosphate-specific phosphodiesterase inhibitor, is an agonist for GPR35. FEBS Lett. 2006, 580, 5003–5008. [Google Scholar] [CrossRef] [PubMed]
  195. Naftali, T.; Bar-Lev Schleider, L.; Dotan, I.; Lansky, E.P.; Sklerovsky Benjaminov, F.; Konikoff, F.M. Cannabis induces a clinical response in patients with Crohn’s disease: A prospective placebo-controlled study. Clin. Gastroenterol. Hepatol. 2013, 11, 1276–1280.e1. [Google Scholar] [CrossRef] [PubMed]
  196. Imielinski, M.; Baldassano, R.N.; Griffiths, A.; Russell, R.K.; Annese, V.; Dubinsky, M.; Kugathasan, S.; Bradfield, J.P.; Walters, T.D.; Sleiman, P.; et al. Common variants at five new loci associated with early-onset inflammatory bowel disease. Nat. Genet. 2009, 41, 1335–1340. [Google Scholar] [CrossRef] [PubMed][Green Version]
  197. Barth, M.C.; Ahluwalia, N.; Anderson, T.J.; Hardy, G.J.; Sinha, S.; Alvarez-Cardona, J.A.; Pruitt, I.E.; Rhee, E.P.; Colvin, R.A.; Gerszten, R.E. Kynurenic acid triggers firm arrest of leukocytes to vascular endothelium under flow conditions. J. Biol. Chem. 2009, 284, 19189–19195. [Google Scholar] [CrossRef] [PubMed]
  198. Gasperi, V.; Evangelista, D.; Chiurchiù, V.; Florenzano, F.; Savini, I.; Oddi, S.; Avigliano, L.; Catani, M.V.; Maccarrone, M. 2-arachidonoylglycerol modulates human endothelial cell/leukocyte interactions by controlling selectin expression through CB1 and CB2 receptors. Int. J. Biochem. Cell Biol. 2014, 51, 79–88. [Google Scholar] [CrossRef] [PubMed]
  199. Haustein, M.; Ramer, R.; Linnebacher, M.; Manda, K.; Hinz, B. Cannabinoids increase lung cancer cell lysis by lymphokine-activated killer cells via upregulation of ICAM-1. Biochem. Pharmacol. 2014, 92, 312–325. [Google Scholar] [CrossRef] [PubMed]
  200. Kianian, M.; Al-Banna, N.A.; Kelly, M.E.; Lehmann, C. Inhibition of endocannabinoid degradation in experimental endotoxemia reduces leukocyte adhesion and improves capillary perfusion in the gut. J. Basic Clin. Physiol. Pharmacol. 2013, 24, 27–33. [Google Scholar] [CrossRef] [PubMed]
  201. Lehmann, C.; Kianian, M.; Zhou, J.; Küster, I.; Kuschnereit, R.; Whynot, S.; Hung, O.; Shukla, R.; Johnston, B.; Cerny, V.; et al. Cannabinoid receptor 2 activation reduces intestinal leukocyte recruitment and systemic inflammatory mediator release in acute experimental sepsis. Crit. Care 2012, 16, R47. [Google Scholar] [CrossRef] [PubMed]
  202. Lunn, C.A.; Fine, J.S.; Rojas-Triana, A.; Jackson, J.V.; Fan, X.; Kung, T.T.; Gonsiorek, W.; Schwarz, M.A.; Lavey, B.; Kozlowski, J.A.; et al. A novel cannabinoid peripheral cannabinoid receptor-selective inverse agonist blocks leukocyte recruitment in vivo. J. Pharmacol. Exp. Ther. 2006, 316, 780–788. [Google Scholar] [CrossRef] [PubMed]
  203. Montecucco, F.; Burger, F.; Mach, F.; Steffens, S. CB2 cannabinoid receptor agonist JWH-015 modulates human monocyte migration through defined intracellular signaling pathways. Am. J. Physiol. Heart Circ. Physiol. 2008, 294, H1145–H1155. [Google Scholar] [CrossRef] [PubMed]
  204. Murikinati, S.; Jüttler, E.; Keinert, T.; Ridder, D.A.; Muhammad, S.; Waibler, Z.; Ledent, C.; Zimmer, A.; Kalinke, U.; Schwaninger, M. Activation of cannabinoid 2 receptors protects against cerebral ischemia by inhibiting neutrophil recruitment. FASEB J. 2010, 24, 788–798. [Google Scholar] [CrossRef] [PubMed]
Figure 1. The kynurenine pathway. The most relevant metabolites of the pathway are highlighted in different colors, and the key enzymes are also represented on the arrows. Dashed arrows indicate the indirect linkage between the kynurenine and serotonin pathway. The two sequentially arrows symbolizes the multiple (not indicated) steps in quinolinic acid metabolization to nicotinamide adenine dinucleotide (NAD+). Abbreviations: IDO (indoleamine 2,3-dioxygenase); KATs (kynurenine aminotransferases); KMO (l-kynurenine 3-monooxygenase); KYNU (l-kynurenine hydrolase); NAD+ (nicotinamide adenine dinucleotid), TDO (tryptophan 2,3-dioxygenase).
Figure 1. The kynurenine pathway. The most relevant metabolites of the pathway are highlighted in different colors, and the key enzymes are also represented on the arrows. Dashed arrows indicate the indirect linkage between the kynurenine and serotonin pathway. The two sequentially arrows symbolizes the multiple (not indicated) steps in quinolinic acid metabolization to nicotinamide adenine dinucleotide (NAD+). Abbreviations: IDO (indoleamine 2,3-dioxygenase); KATs (kynurenine aminotransferases); KMO (l-kynurenine 3-monooxygenase); KYNU (l-kynurenine hydrolase); NAD+ (nicotinamide adenine dinucleotid), TDO (tryptophan 2,3-dioxygenase).
Ijms 18 01617 g001
Figure 2. The type 1 cannabinoid receptor-N-methyl-d-aspartate (CB1-NMDA) receptor complex and potential pharmacological targets to reduce N-methyl-d-aspartate (NMDA) receptor hyperactivity, which is one of the main pathomechanism of migraine. The pharmacological inhibition (red arrow) of endocannabinoid metabolizing enzymes, monoacylglycerol lipase (MAGL) or fatty acid amide hydrolase (FAAH) (1) increases (upward green arrow) endocannabinoid levels (e.g., 2-arachidonoylglycerol (2-AG), anandamide (AEA)) (2) [113], thus enhancing the agonist-mediated type 1 cannabinoid receptor (CB1) receptor activity (3) (in this step the black arrows indicate ligand binding). This mechanism will overall reduce (red arrow) the activity of the NMDA receptor, hence the risk of excitotoxicity via CB1 receptor and the σ1R-HINT1 protein tandem (4) [124,153,154]. Exogenous ligands can also induce CB1-receptor mediated NMDA receptor inhibition more effectively (3,4) [124]. Additionally, exogenous cannabinoids such as cannabidiol or Δ9-tetrahydrocannabinol are known to stimulate (blue arrow) the indoleamine 2,3-dioxygenase (IDO) enzyme activity in dependence of cannabinoid receptor activation (5) [120]. This stimulation, together with the pharmacological inhibition (red arrow) of the kynurenine 3-monooxygenase (KMO) enzyme (6) [123] may enhance (yellow upward arrow) endogenous KYNA levels indirectly (indicated by dashed lines), through the kynurenine pathway (7) (Figure 1), which will result an enhanced reduction in NMDA receptor activity via the antagonizing effect (black arrow) of kynurenic acid (KYNA) (8).The figure shows a simplified, hypothetical scenario of the indicate elements and mechanisms of the endocannabinoid and kynurenine system within the CB1-NMDA receptor complex, which has been individually reported previously in other circumstances (cited accordingly). The figure also indicates the sigma 1 receptors- histidine triad nucleotide-binding protein 1 (σ1R-HINT1) protein tandem, which associates the two receptors and it is based on Rodriguez-Munoz and co-worker’s review [124]. NR1 and NR2 indicate the two types of subunit of the NMDA receptor and the C terminus of the CB1 receptor is also highlighted. The green color indicates endocannabinoid, while the yellow color indicates kynurenine system related ligands, receptors, enzymes or mechanisms. The shapes of the indicated ligands, receptors or enzymes are schematic or overly simplified representations of their structures.
Figure 2. The type 1 cannabinoid receptor-N-methyl-d-aspartate (CB1-NMDA) receptor complex and potential pharmacological targets to reduce N-methyl-d-aspartate (NMDA) receptor hyperactivity, which is one of the main pathomechanism of migraine. The pharmacological inhibition (red arrow) of endocannabinoid metabolizing enzymes, monoacylglycerol lipase (MAGL) or fatty acid amide hydrolase (FAAH) (1) increases (upward green arrow) endocannabinoid levels (e.g., 2-arachidonoylglycerol (2-AG), anandamide (AEA)) (2) [113], thus enhancing the agonist-mediated type 1 cannabinoid receptor (CB1) receptor activity (3) (in this step the black arrows indicate ligand binding). This mechanism will overall reduce (red arrow) the activity of the NMDA receptor, hence the risk of excitotoxicity via CB1 receptor and the σ1R-HINT1 protein tandem (4) [124,153,154]. Exogenous ligands can also induce CB1-receptor mediated NMDA receptor inhibition more effectively (3,4) [124]. Additionally, exogenous cannabinoids such as cannabidiol or Δ9-tetrahydrocannabinol are known to stimulate (blue arrow) the indoleamine 2,3-dioxygenase (IDO) enzyme activity in dependence of cannabinoid receptor activation (5) [120]. This stimulation, together with the pharmacological inhibition (red arrow) of the kynurenine 3-monooxygenase (KMO) enzyme (6) [123] may enhance (yellow upward arrow) endogenous KYNA levels indirectly (indicated by dashed lines), through the kynurenine pathway (7) (Figure 1), which will result an enhanced reduction in NMDA receptor activity via the antagonizing effect (black arrow) of kynurenic acid (KYNA) (8).The figure shows a simplified, hypothetical scenario of the indicate elements and mechanisms of the endocannabinoid and kynurenine system within the CB1-NMDA receptor complex, which has been individually reported previously in other circumstances (cited accordingly). The figure also indicates the sigma 1 receptors- histidine triad nucleotide-binding protein 1 (σ1R-HINT1) protein tandem, which associates the two receptors and it is based on Rodriguez-Munoz and co-worker’s review [124]. NR1 and NR2 indicate the two types of subunit of the NMDA receptor and the C terminus of the CB1 receptor is also highlighted. The green color indicates endocannabinoid, while the yellow color indicates kynurenine system related ligands, receptors, enzymes or mechanisms. The shapes of the indicated ligands, receptors or enzymes are schematic or overly simplified representations of their structures.
Ijms 18 01617 g002

Share and Cite

MDPI and ACS Style

Nagy-Grócz, G.; Zádor, F.; Dvorácskó, S.; Bohár, Z.; Benyhe, S.; Tömböly, C.; Párdutz, Á.; Vécsei, L. Interactions between the Kynurenine and the Endocannabinoid System with Special Emphasis on Migraine. Int. J. Mol. Sci. 2017, 18, 1617.

AMA Style

Nagy-Grócz G, Zádor F, Dvorácskó S, Bohár Z, Benyhe S, Tömböly C, Párdutz Á, Vécsei L. Interactions between the Kynurenine and the Endocannabinoid System with Special Emphasis on Migraine. International Journal of Molecular Sciences. 2017; 18(8):1617.

Chicago/Turabian Style

Nagy-Grócz, Gábor, Ferenc Zádor, Szabolcs Dvorácskó, Zsuzsanna Bohár, Sándor Benyhe, Csaba Tömböly, Árpád Párdutz, and László Vécsei. 2017. "Interactions between the Kynurenine and the Endocannabinoid System with Special Emphasis on Migraine" International Journal of Molecular Sciences 18, no. 8: 1617.

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop