Content uploaded by Jenny Felth
Author content
All content in this area was uploaded by Jenny Felth
Content may be subject to copyright.
Cyclooxygenase enzymes (COX enzymes) catalyse the
production of prostaglandins, which are important mediators
in the inflammatory process. To date, two isoforms of COX
have been identified; a constitutively expressed enzyme,
COX-1 and an inducible enzyme, COX-2,1,2) of which the lat-
ter is induced by inflammatory stimuli. An important group
of anti-inflammatory drugs is the Non-Steroid Anti Inflam-
matory Drugs (NSAIDs), of which aspirin and indomethacin
are representatives. These compounds act by inhibiting the
COX enzymes. The substrate for the prostaglandin produc-
tion is arachidonic acid (Fig. 1),1) an eicosanoid, which is
produced on demand by phospholipase A2from arachido-
nate, which is stored in the lipid bilayers of the cell wall.3) In
recent years, COX-2 overexpression has been associated with
colon cancer development, and COX enzyme inhibition is
studied as a potential target for cancer chemoprevention.4,5)
Other compounds in the eicosanoid group are the endo-
cannabinoids. These endogenous compounds bind to cellular
receptors, including the cannabinoid receptors, which are the
molecular targets of the active principle in Cannabis sativa.
The biological function of the endocannabinoids involves
several regulatory agents, forming the endocannabinoid sys-
tem (ECS).6) It has been reported that endocannabinoids also
can function as substrates for the COX enzymes resulting in
production of prostaglandin ethanolamides and prostaglandin
glycerol esters.7,8) Recently, the endogenous cannabinoid
anandamide was shown to induce COX-2 dependent cell
death in colon cancer cells.9)
There are several structural (Fig. 1) and physiological sim-
ilarities between human endocannabinoids and cannabinoids
occurring in plant material. Structurally the 5-carbon side
chain in cannabinoids is present in the endocannabinoids as
the last five carbons of the fatty acid chain, and the C-3 OH
might correspond to the polar hydroxyl end of the endo-
cannabinoids. Furthermore, the relative distances between
774 Vol. 34, No. 5Note
Evaluation of the Cyclooxygenase Inhibiting Effects of Six Major
Cannabinoids Isolated from Cannabis sativa
Lucia Renee RUHAAK,*,a,b,† Jenny FELTH,a,# Pernilla Christina KARLSSON,cJoseph James RAFTER,c
Robert VERPOORTE,band Lars BOHLINa
aDivision of Pharmacognosy, Department of Medicinal Chemistry, Biomedical Centre, Uppsala University; Box 574, SE-
751 23 Uppsala, Sweden: bDivision of Pharmacognosy, Section Metabolomics, Institute of Biology, Leiden University;
2300 RA, Leiden, the Netherlands: and cDepartment of Biosciences and Nutrition, Karolinska Institutet; Novum, S-141 86
Huddinge, Sweden. Received December 22, 2010; accepted February 19, 2011; published online February 28, 2011
Cyclooxygenase enzymes (COX-1 and COX-2) catalyse the production of prostaglandins from arachidonic
acid. Prostaglandins are important mediators in the inflammatory process and their production can be reduced
by COX-inhibitors. Endocannabinoids, endogenous analogues of the plant derived cannabinoids, occur normally
in the human body. The Endocannabinoids are structurally similar to arachidonic acid and have been suggested
to interfere with the inflammatory process. They have also been shown to inhibit cancer cell proliferation. Anti-
inflammatory effects of cannabinoids and endocannabinoids have been observed, however the mode of action is
not yet clarified. Anti-inflammatory activity (i.e., inhibition of COX-2) is proposed to play an important role in
the development of colon cancer, which makes this subject interesting to study further. In the present work, the
six cannabinoids tetrahydrocannabinol (DD9-THC), tetrahydrocannabinolic acid (DD9-THC-A), cannabidiol (CBD),
cannabidiolic acid (CBDA), cannabigerol (CBG) and cannabigerolic acid (CBGA), isolated from Cannabis sativa,
were evaluated for their effects on prostaglandin production. For this purpose an in vitro enzyme based COX-
1/COX-2 inhibition assay and a cell based prostaglandin production radioimmunoassay were used. Cannabinoids
inhibited cyclooxygenase enzyme activity with IC50 values ranging from 1.7· 10
3to 2.0 · 104M.
Key words cannabinoid; cyclooxygenase inhibition; prostaglandin production
Biol. Pharm. Bull. 34(5) 774—778 (2011)
© 2011 Pharmaceutical Society of Japan
∗To whom correspondence should be addressed. e-mail: Lruhaak@ucdavis.edu
#Equal contribution with the first author.
†
Present address: Department of Chemistry, University of California at Davis,
Davis, CA 95616, U.S.A.
Fig. 1. Structural Formulas of the Endocannabinoid Anandamide (1) and
the Endocannabinoid Precursor Arachidonic Acid (2) together with the Six
Cannabinoids; D9-THC (3), D9-THCA-A (4), CBD (5), CBDA (6), CBG (7),
CBGA (8)
the groups are comparable due to the ring system in cannabi-
noids, which can be mimicked by the U-shaped endocannabi-
noids and their four double bonds.10) Also physiologically
there are similarities, since both cannabinoids and endo-
cannabinoids bind to the cannabinoid receptors.11) Endo-
cannabinoids, such as anandamide, are derived from arachi-
donic acid and are structurally similar to this compound (Fig.
1).
Altogether, these similarities gave rise to the hypothesis
that cannabinoids can affect the COX enzyme activity.
Several studies have demonstrated anti-inflammatory activi-
ties in vivo and in vitro for various cannabinoid com-
pounds,12—18) which makes this hypothesis very plausible. In-
hibiting effects on COX enzyme activity have also previously
been observed for cannabidiol and cannabidiolic acid,17,19)
and cannabinoids have potential to affect the potency of
NSAIDs.20,21) Furthermore, in recent years, it has been shown
that the ECS can protect against colonic inflammation,6,22)
which is of interest in prevention of bowel disease and colo-
rectal cancer. The cannabinoid receptors are suggested to be
involved in the control of colonic inflammation,6,22) however,
the mode of action for the anti-inflammatory effects of
cannabinoids is not yet clarified.
In the present study we evaluated the COX-mediated anti-
inflammatory properties of six different naturally occuring
cannabinoids; tetrahydrocannabinol (D9-THC), tetrahydro-
cannabinolic acid-A (THCA-A), cannabidiol (CBD), canna-
bidiolic acid (CBDA), cannabigerol (CBG) and cannabigero-
lic acid (CBGA) (Fig. 1). An enzyme-based in vitro COX
inhibition assay was used to evaluate the effects on both
COX-1 and COX-2 on enzyme-level, while a cell-based
prostaglandin production assay was used to evaluate the
effects on COX-2 at cellular level.
MATERIALS AND METHODS
Materials All solvents were purchased from Lab-Scan,
Dublin, Ireland, and were of analytical grade. Scientific sam-
ples of cannabinoids (D9-THC, THCA-A, CBD, CBDA,
CBG and CBGA) were provided by Prof. Robert Verpoorte
and Dr. Arno Hazekamp, Leiden University, The Nether-
lands. The cannabinoids were isolated from Cannabis sativa
and characterized and quantified using the chromatography
and 1H-NMR methods as described by Hazekamp et al.23,24)
All cannabinoid samples were at least 92% pure.
COX-1 enzyme, purified from ram seminal vesicles and
COX-2 enzyme, purified from sheep placental cotyledons,
and the reference compound NS-398 (N-[2-(cyclo-hexyl-
oxy)-4-nitrophenyl]methanesulphonamide) were purchased
from Cayman Chemical Co., Ann Arbor, MI, U.S.A.
Hematin was obtained from ICN biomedicals Inc., Aurora,
Ohio, U.S.A. Adrenalin was purchased from Apoteket AB,
Göteborg, Sweden. Reduced gluthatione, indomethacine, un-
labeled arachidonic acid, anti-prostaglandin E2, prostaglandin
E2standard, Bovine Serum Albumin, tumor necrosis factor
(TNF)-
a
and charcoal were obtained from Sigma-Aldrich,
St. Louis, MO, U.S.A. 14C-Arachidonic acid, [5,6,8,11,12,-
14,15(n)-3H] Prostaglandin E2and dextran molecular weight
(mw) 70000 was purchased from Amersham Pharmacia,
Stockholm, Sweden, while silica gel 60, particle size 0.063—
2 mm was obtained from Merck, Darmstadt, Germany. Dul-
becco’s modified Eagle’s medium (DMEM)-high glucose and
trypsin-ethylenediaminetetraacetic acid (EDTA) were ob-
tained from Invitrogen, Taastrup, Denmark.
Cell Culture The human colon adenocarcinoma cell line
HT29, was cultured in monolayer in DMEM (Dulbecco’s
modified Eagle medium supplemented with 10% fetal bovine
serum (FBS), 2 mML-glutamine, and 1% penicillin/strepto-
mycin) at 37 °C and 5% CO2. All experiments were per-
formed with 60—80% confluent cells and 0.1% DMEM
(0.1% FBS). Pure compounds were dissolved in ethanol and
diluted in 0.1% DMEM (with the final concentration in the
cell cultures being maximum 0.25% ethanol).
Enzyme-Based Inhibition Assay The assay followed
the original method described by White and Glassman,25)
with modifications as described by Noreen et al.26) The assay
described below was used for both COX-1 and COX-2 en-
zymes. In short, 20
m
l of each sample was dispensed in a 96-
well plate. All samples were dissolved in 20% dimethyl sul-
foxide (DMSO) in TRIS buffer. To determine minimal and
maximal activity of the enzyme, 20% DMSO in TRIS buffer
was used as the sample. Total inhibition of the enzyme in the
minimum wells was reached by addition of 10
m
l of 2 MHCl
to the wells before the enzyme was added. Cofactors were
dissolved in TRIS buffer to concentrations of 1.27 mg/ml
hematin, 6.50 mg/ml adrenalin and 1.50 mg/ml gluthatione,
giving final concentrations in the wells of 1.3
m
g/ml, 1.3
mg/ml and 0.3 mg/ml respectively. COX enzyme was mixed
with the co-factors, pre-incubated and activated on ice for
5 min. Sixty microliters of enzyme-cofactor solution was
added to the sample in the wells, and the plate was incubated
for 10 min on ice. The activity of the enzyme in the wells was
6U (COX-1) or 3U (COX-2). Twenty microliters of 14C-
arachidonic acid (14C-AA) solution was dispensed in each
well and to start the enzymatic reaction, the plate was incu-
bated in a 37 °C waterbath for 15 min (COX-1) or 3min
(COX-2). The reaction was stopped by addition of 10
m
l
of HCl (2 M). To separate the non-converted 14C-AA from
the 14C-labeled prostaglandins, column chromatography
(Silica gel 60, particle size 0.063—2 mm) was used. The
columns were equilibrated using 2 ml of eluent, consisting of
heptane : ethyl acetate : acetic acid (70 : 30 : 1), thereafter the
samples were applied, and the non-converted AA was eluted
using 4 ml of the same eluent. The prostaglandins were
then eluted using 3 ml of a second eluent, consisting of
dioxane : methanol (85 : 15). Scintillation fluid was added
to the samples, and the amount of radioactively labeled
prostaglandin in the samples was determined using a Packard
scintillation spectrometer. Percent inhibition values were cal-
culated and IC50-values were obtained by applying the non-
linear regression analysis tool of Graph Pad Prism (Graph-
Pad Software Inc., CA, U.S.A.).
Prostaglandin E2(PGE2) Production in HT29 Cells
PGE2is a major product produced by COX from arachidonic
acid and is often used to estimate COX activity in cells. The
method used is a standard procedure for measuring PGE2
production in cells, and has previously been described in de-
tail.27—29) In brief, HT29 cells were seeded out at a concen-
tration of 3.30105cells/well. At day 2, 100
m
Maspirin was
added to the wells to prevent activation of COX-1. At day 3,
the cells were incubated with TNF-
a
(50 ng/ml) and cannabi-
noid samples (12.5 or 25
m
M) for 5 h, thereafter the test solu-
May 2011 775
tion was replaced with medium containing 100
m
mol/l
arachidonic acid (Sigma) and the cells were incubated for 1h.
The concentration of released prostaglandin E2(PGE2) was
quantified using radio immuno-assay (RIA), according to the
protocol supplied by Sigma Chemical Co., using [3H]PGE2
and polyclonal antiserum to PGE2(Sigma). The amount of
prostaglandins in each sample was detected using a scintilla-
tion counter, and expressed as the percentage inhibition of
the TNF-
a
treated cells. Each cannabinoid was tested at least
twice in the cell system and later analyzed in duplicate in the
RIA. The results were expressed as the percentage inhibition
of the TNF-
a
treated cells. In all experiments untreated cells
were included as controls, and the selective COX-2 inhibitor
NS398 was used as a reference compound for comparison of
inhibiting activity.
Prior to the PGE2 experiments, all cannabinoid samples
were tested for cytotoxicity in the AlamarBlueTM assay to en-
sure that potential COX-2 inhibitory effects were not due to
cell death.30,31) A cell survival of approximately 70% was
considered as acceptable for studying the prostaglandin pro-
duction. Cannabinoid concentrations causing cell death (i.e.,
cell survival 70%) were excluded from the PGE2produc-
tion experiments.
RESULTS
Enzyme-Based Inhibition Assay The inhibitory effects
of six cannabinoids on the cyclooxygenase enzyme activity
was evaluated by an in vitro COX enzyme inhibition assay.
D9-THC, D9-THCA-A, CBD, CBDA, CBG and CBGA were
screened for their ability to inhibit COX-1 and COX-2 at a
concentration of 100 mg/ml (approximately 3·104M), since
higher concentrations were assumed to be irrelevant. In this
screening, an enzyme inhibition of 30% was considered as
sufficient to be relevant, and was set as a cutoff limit for
compounds to investigate further. D9-THCA-A, CBDA, CBG
and CBGA showed more than 30% inhibition on COX-1
(Fig. 2). The concentration-dependent activity (i.e. inhibition
of COX-1) for these compounds was further evaluated at
concentrations ranging from 3.18 · 103to 2.78 · 105M, as
presented by concentration–effect graphs (Fig. 3A). The
IC50-values are presented in Table 1. The IC50-value of the
reference compound indomethacin was within acceptable
limits of the value reported previously for this COX-1 assay
(1.4 · 106M),26) confirming that the assay was successful.
When screened for COX-2 enzyme inhibiting activity D9-
THCA-A, CBG and CBGA showed more than 30% inhibi-
tion. Interestingly, CBDA, which was recently reported to se-
lectively inhibit COX-2,19) did not reach the 30% inhibition
threshold (Fig. 2), and was therefore not considered in our
further COX-2 inhibition studies. The inhibition of D9-
THCA-A, CBG and CBGA was measured at concentrations
ranging from 3.18 · 103to 2.78 · 105M, as represented by
the concentration–effect graphs (Fig. 3B) with IC50-values
presented in Table 1. The IC50 value of the reference com-
pound indomethacin was within acceptable limits of the
value previously reported for this COX-2 assay (1.64·106
M),26) confirming that the assay results were reliable.
Complementary to the enzyme-inhibition assay, the effects
776 Vol. 34, No. 5
Fig. 2. Screening of Six Cannabinoids for Their Potential to Inhibit COX-
1 and COX-2 Enzymes
All cannabinoids were screened at concentrations of 100
m
g/ml. To justify further
analysis, a cut off value of at least 30% inhibition was used, represented by the black
dotted line.
Fig. 3. (A) Graphs Representing the COX-1 Inhibition of D9-THCA-A,
CBDA, CBGA and Indomethacin in the Enzyme Based Assay
For each datapoint n3.
(B) Graphs Representing the COX-2 Inhibiton of D9-THCA-A, CBG,
CBGA and Indomethacin in the Enzyme Based Assay
For each datapoint n3.
Table 1. COX Inhibition IC50-Values Determined for D9-THCA-A, CBG,
CBGA and Indomethacin Using an Enzyme Based in Vitro Assay
IC50 (M)
Compound
COX-1 COX-2
D9-THCA-A 1.7 · 1036.3·10
4
CBDA 4.7 · 104N.D.a)
CBG N.D.a)2.7·10
4
CBGA 4.6 · 1042.0·10
4
Indomethacin 3.1 · 1069.3·10
5
a) N.D., not determined.
of cannabinoids on prostaglandin production were examined
in a cell based assay. Six different cannabinoids were tested
for their ability to decrease prostaglandin production in TNF-
a
stimulated HT29 cells. Prior to measuring the prosta-
glandin production, the effects of cannabinoids on cell sur-
vival were investigated, to make sure that the effects were not
due to cell death. A cell survival of approximately 70% was
considered as acceptable for studying the prostaglandin pro-
duction, and the observed effects on the PGE2production are
very unlikely to be explained by cell death. Both apoptosis
and necrosis make the cells detach from the plate surface. No
such signs were observed. D9-THC, CBD, CBDA and CBG
were tested at concentrations of 2.5·105M, whereas D9-
THCA-A and CBGA were tested at a concentration of
6.25 · 105M. However, higher concentrations of cannabi-
noids caused a high cytotoxicity and could not be used in the
experiments. The results, as presented in Fig. 4, showed that
D9-THC, D9-THCA-A, CBD, CBG and CBGA inhibited
prostaglandin production, however the level of inhibition was
low (10%). CBDA, on the other hand seemed to stimulate
the prostaglandin production (Fig. 4).
DISCUSSION
Cannabinoids have been shown to possess anti-inflamma-
tory effects,12—18) but the mechanism of action is not yet
known. COX enzyme inhibiting activity has previously been
observed for CBD and CBDA.17,19) Overexpression of COX-
2 has in recent years also been associated with colon cancer
development,5) and COX-2 enzyme inhibition is regarded as
a potential target for cancer chemoprevention.4) Interestingly,
endocannabinoid levels are elevated in colon cancer tissue,
and they also inhibit cancer cell proliferation by acting at
cannabinoid receptors.32) Recently, it has also been shown
that the ECS can protect against colonic inflammation,6,22)
which is of interest in prevention of bowel disease and col-
orectal cancer. Additionally, cannabinoids have been shown
to affect the potency of NSAIDs,20,21) potentially via modula-
tion of the COX pathway.
In the present study, six major cannabinoids isolated from
plant material modulated the activity of COX enzymes, with
IC50 values ranging from 1.7·103to 2.0 · 104M. None of
the cannabinoids showed high COX selectivity except from
CBDA, which only inhibited COX-1. This finding is contra-
dictory to previously reported results by Takeda et al., where
CBDA was found to be a selective COX-2 inhibitor in an en-
zyme inhibition assay using purified COX enzymes.19) These
inconsistencies might be caused by differences in the detec-
tion method. In the present study radioactively labeled
prostaglandin was measured, while Takeda et al. measured
the oxidation of TMPD spectrophotometrically. Alternatively,
as the cannabinoids used in the studies were purified from
plant material, different impurities in the samples could
cause different results. Further studies, preferably in human
cell lines, are needed to validate the COX inhibition by
cannabinoids.
In the screening, it was observed that D9-THC showed
stimulation in a dose-related matter (between 3.18·104and
3.18 · 105M) both in the COX-1 inhibition assay and the
COX-2 inhibition assay (data not shown). However, the
COX-inhibition assay is not designed to quantify COX
enzyme activation, and hence no definitive conclusions can
be drawn from these findings.
Interestingly, CBD and D9-THC showed low activity in the
in vitro assay of the COX-enzymes in comparison with the
other cannabinoids tested. The COX inhibition assay is an in
vitro assay where purified COX enzyme (from ram seminal
vesicles and sheep placental cotyledons, respectively) is
used. This assay is far from the human in vivo conditions.
Therefore, we complementarily used human colon cancer
cells to investigate if the prostaglandin production would be
inhibited also in living cells. The inhibition of prostaglandin
production in cancer cells is of great interest, since the in-
flammatory process is believed to be of importance for colon
carcinogenesis.33) As shown in Fig. 4, the results (e.g. inhibi-
tion of PGE2production) from the cell-based assay were sim-
ilar for all cannabinoids. All compounds tested inhibited the
production of PGE2only slightly. An experiment with higher
concentrations might give more clear results. However,
higher concentrations of the cannabinoids were cytotoxic,
causing detachment of cells and signs of cell death, and such
experiments were not possible to perform using this cell-
based assay.
The cannabinoids are known to be involved in the immune
system via the CB2receptor. The binding constants Kifor
D9-THC interacting with the CB1and CB2receptors are
8.0·10
5Mand 3.2 · 105Mrespectively.34) These binding
constants are in the same range as the IC50 values we found
for the COX-inhibition by cannabinoids. This might indicate
a possibility of physiologically important effects of the COX-
inhibiting cannabinoids via interaction with the COX-en-
zymes. Further in vitro studies are required to prove such ef-
fects, but the present study shows that several of the major
cannabinoids may also affect other receptors than CB1and
CB2. Interestingly, a recent report, linking COX-2 inhibition
to increased endocannabinoid levels, suggests the ECS and
the COX-mediated prostaglandin pathway to be closely con-
nected.35)
In conclusion, it is clear that cannabinoids inhibit COX-
enzymes, but in a higher concentration range, as compared
to anti-inflammatory drugs (i.e. indomethacin). The obvious
contradiction regarding the selectivity for CBDA, as com-
pared to the previous report by Takeda et al.,19) is interesting
May 2011 777
Fig. 4. Decrease in Prostaglandin Production in TNF-
a
Stimulated HT29
Cells
The prostaglandin production inhibitor NS398 was used as a reference compound.
Error bars represent S.D.
and should be object for further investigation. Additional
studies will also be needed to conclude the relevance of the
COX-inhibitory effects in relation to other anti-inflammatory
activities mediated by cannabinoids. As evident from recent
reports, the ECS plays an important role in the human body.
Interestingly, colonic inflammation can be controlled via the
ECS, and plant-derived cannabinoids may have a potential to
be used as future therapeutic agents.
Acknowledgements This work was financially support-
ed by Grants from the Agricultural Sciences and Spatial
Planning (FORMAS) and European University Consortium
for Pharmaceutical Research (ULLA). The authors also want
to thank Dr. Ulrika Huss Melin for helpful discussion and Dr.
Arno Hazekamp for providing purified cannabinoid samples.
REFERENCES
1) Chandrasekharan N. V., Simmons D. L., Genome Biol., 5, 241 (2004).
2) Simmons D. L., Botting R. M., Hla T., Pharmacol. Rev., 56, 387—437
(2004).
3) Leslie C. C., Biochem. Cell Biol., 82, 1—17 (2004).
4) Ruegg C., Zaric J., Stupp R., Ann. Med., 35, 476—487 (2003).
5) Zha S., Yegnasubramanian V., Nelson W. G., Isaacs W. B., De Marzo
A. M., Cancer Lett., 215, 1—20 (2004).
6) Maccarrone M., Gasperi V., Catani M. V., Diep T. A., Dainese E.,
Hansen H. S., Avigliano L., Annu. Rev. Nutr., 30, 423—440 (2010).
7) Kozak K. R., Rowlinson S. W., Marnett L. J., J. Biol. Chem., 275,
33744—33749 (2000).
8) Yu M., Ives D., Ramesha C. S., J. Biol. Chem., 272, 21181—21186
(1997).
9) Patsos H. A., Greenhough A., Hicks D. J., Al Kharusi M., Collard T. J.,
Lane J. D., Paraskeva C., Williams A. C., Int. J. Oncol., 37, 187—193
(2010).
10) Tong W., Collantes E. R., Welsh W. J., Berglund B. A., Howlett A. C.,
J. Med. Chem., 41, 4207—4215 (1998).
11) Martin B. R., Jefferson R. G., Winckler R., Wiley J. L., Thomas B. F.,
Crocker P. J., Williams W., Razdan R. K., Eur. J. Pharmacol., 435,
35—42 (2002).
12) Malfait A. M., Gallily R., Sumariwalla P. F., Malik A. S., Andreakos
E., Mechoulam R., Feldmann M., Proc. Natl. Acad. Sci. U.S.A., 97,
9561—9566 (2000).
13) Sofia R. D., Knobloch L. C., Vassar H. B., Res. Commun. Chem.
Pathol. Pharmacol., 6, 909—918 (1973).
14) Sofia R. D., Nalepa S. D., Harakal J. J., Vassar H. B., J. Pharmacol.
Exp. Ther., 186, 646—655 (1973).
15) Sofia R. D., Nalepa S. D., Vassar H. B., Knobloch L. C., Life Sci., 15,
251—260 (1974).
16) Wirth P. W., Watson E. S., ElSohly M., Turner C. E., Murphy J. C., Life
Sci., 26, 1991—1995 (1980).
17) Costa B., Colleoni M., Conti S., Parolaro D., Franke C., Trovato A. E.,
Giagnoni G., Naunyn Schmiedebergs Arch. Pharmacol., 369, 294—
299 (2004).
18) Burstein S. H., Zurier R. B., AAPS J., 11, 109—119 (2009).
19) Takeda S., Misawa K., Yamamoto I., Watanabe K., Drug Metab. Dis-
pos., 36, 1917—1921 (2008).
20) Anikwue R., Huffman J. W., Martin Z. L., Welch S. P., J. Pharmacol.
Exp. Ther., 303, 340—346 (2002).
21) Guindon J., De Lean A., Beaulieu P., Pain, 121, 85—93 (2006).
22) Massa F., Marsicano G., Hermann H., Cannich A., Monory K., Cravatt
B. F., Ferri G. L., Sibaev A., Storr M., Lutz B., J. Clin. Invest., 113,
1202—1209 (2004).
23) Hazekamp A., Simons, R., Peltenburg-Looman, A., Sengers, M., Van
Zweden, P., Verpoorte, R., J. Liq. Chromatogr. Relat. Technol., 27,
2421—2439 (2004).
24) Hazekamp A., Choi Y. H., Verpoorte R., Chem. Pharm. Bull., 52,
718—721 (2004).
25) White H. L., Glassman A. T., Prostaglandins, 7, 123—129 (1974).
26) Noreen Y., Ringbom T., Perera P., Danielson H., Bohlin L., J. Nat.
Prod., 61, 2—7 (1998).
27) Calviello G., Di Nicuolo F., Gragnoli S., Piccioni E., Serini S., Mag-
giano N., Tringali G., Navarra P., Ranelletti F. O., Palozza P., Carcino-
genesis, 25, 2303—2310 (2004).
28) Pettersson J., Karlsson P. C., Goransson U., Rafter J. J., Bohlin L., Biol.
Pharm. Bull., 31, 534—537 (2008).
29) Yamada M., Niki H., Yamashita M., Mue S., Ohuchi K., J. Pharmacol.
Exp. Ther., 281, 1005—1012 (1997).
30) Klinder A., Gietl E., Hughes R., Jonkers N., Karlsson P., McGlyn H.,
Pistoli S., Tuohy K., Rafter J., Rowland I. R., van Loo J., L. P.-Z. B.,
Int. J. Cancer Prev., 1, 19—32 (2004).
31) O’Brien J., Wilson I., Orton T., Pognan F., Eur. J. Biochem., 267,
5421—5426 (2000).
32) Ligresti A., Bisogno T., Matias I., De Petrocellis L., Cascio M. G.,
Cosenza V., D’Argenio G., Scaglione G., Bifulco M., Sorrentini I., Di
Marzo V., Gastroenterology, 125, 677—687 (2003).
33) Terzic J., Grivennikov S., Karin E., Karin M., Gastroenterology, 138,
2101—2114 e2105 (2010).
34) Rhee M. H., Vogel Z., Barg J., Bayewitch M., Levy R., Hanus L.,
Breuer A., Mechoulam R., J. Med. Chem., 40, 3228—3233 (1997).
35) Telleria-Diaz A., Schmidt M., Kreusch S., Neubert A. K., Schache F.,
Vazquez E., Vanegas H., Schaible H. G., Ebersberger A., Pain, 148,
26—35 (2010).
778 Vol. 34, No. 5
