|
Abstract
OBJECTIVES : Given the presence of morphine, its metabolites
and precursors in mammalian and invertebrate tissues, it became
important to determine if exposing tissues to an opiate alkaloid
precursor, reticuline, would result in increasing endogenous
morphine levels.
METHOD : Endogenous morphine levels were determined
by high pressure liquid chromatography coupled to electrochemical
detection and radioimmunoassay following incubation of Mytilus
edulis pedal ganglia with reticuline. Nitric oxide (NO)
release was determined in real-time via an amperometric probe.
Mu opiate receptor affinity for opiate alkaloid precursors was
determined by a receptor displacement assay.
RESULTS : Morphine is present in the pedal ganglia of
Mytilus edulis (1.43 ± 0.41 ng/mg ± SEM ganglionic
wet weight). Ganglia incubated with 50 ng of reticuline, a morphine
precursor in plants, for 1 hour exhibited a statistical increase
in their endogenous morphine levels (6.7 ± 0.7 ng/mg tissue
wet weight ; P<0.01). This phenomenon is concentration dependent.
The increase in ganglionic morphine levels occurs gradually
over the 60 min incubation period, beginning 10 minutes post
reticuline addition. We show that reticuline (10-6 M) does not
stimulate ganglionic NO release in a manner resembling that
of morphine (10-6 M), which releases NO seconds after its exposure
to the ganglia and lasts for 5 minutes. With reticuline, there
is a 3 minute delay, which is followed by an extended release
period. Furthermore, in binding displacement experiments both
reticuline and salutaridine (another morphine precursor) exhibit
no binding affinity for the pedal ganglia mu opiate receptor
subtype. This finding is further substantiated using the positive
control of human monocytes where the mu3 opiate receptor subtype
has been cloned.
CONCLUSION : Taken together, we surmise that the morphine's
precursors are being converted to morphine. The experiments
strongly indicate that pedal ganglia can synthesize morphine
from reticuline.
Abbreviations : NO - Nitric oxide ; HPLC - High pressure
liquid chromatography ; THP - tetrahydropapoverine ; PBS - phosphate
buffered saline ; RIA - radioimmunoassay
Introduction
There is a body of evidence indicating that opiate alkaloids
such as morphine, morphine-3- and 6- glucuronide, as well as
the morphine putative precursor molecules (thebaine, salutaridine,
norcocolarine, reticuline, tetrahydropapoverine (THP) and codeine)
exist in vertebrates [12, 22,
14, 55]. In invertebrates,
specifically Mytilus edulis, the presence of morphine,
morphine-6-glucuronide, morphine-3-glucuronide, codeine, THP
and reticuline also have been reported [40,
17, 41, 53,
56]. Taken together, these data provide important
evidence supporting the hypothesis that animals have the ability
to synthesize opiate alkaloids.
Based on the above reports, it is important to determine if
a morphine precursor found in animal tissues has the ability
to augment endogenous morphine levels, indicating that morphine
synthesis is occurring. In the present report, we demonstrate
that exposing Mytilus edulis pedal ganglia to low levels
of reticuline results in enhancing ganglionic morphine levels,
strongly suggesting, as in plants, this molecule leads to morphine
biosynthesis [6].
Material and Methods
Mytilus edulis collected from the local waters of
Long Island Sound were maintained under laboratory conditions
for at least 14 days prior using in experiments. Mussels were
kept in artificial seawater (Instant Ocean, Aquarium Systems,
Mentor, Ohio) at a salinity of 30 PSU and at a temperature of
18°C as previously described [47].
Biochemical Analysis
For reticuline exposure, 400 animals were placed and maintained
in artificial seawater at 24°C whereas control animals (100)
were exposed to vehicle (PBS). For the biochemical analysis,
groups of 20 animals had their pedal ganglia excised at different
time periods after incubation with reticuline.
Morphine Determination, Solid Phase Extraction
The extraction protocol, using internal or external morphine
standards, was performed in a room where the animals were not
maintained to avoid morphine contamination. Single use siliconized
tubes were used to prevent the loss of morphine. Mytilus
edulis pedal ganglia also were extensively washed (3 times)
with PBS (0.01M NaCl 0.132 mM, NH4HCO3 0.132 mM ; pH 7.2) prior
to extraction (3 times centrifugation at 1000 rpm, 1 min, then
discard the PBS) to avoid exogenous morphine contamination.
Tissues were dissolved in 1N HCl and sonicated using a Fisher
scientific sonic dismembrator 60 (Fisher Scientific, USA). The
resulting homogenates were extracted with 5 ml chloroform/isopropanol
9 :1. After 5 min at room temperature, homogenates were centrifuged
at 3000 rpm for 15 min. The three phases were separated in the
following order :
- 1) The lowest layer corresponding to the organic phase ;
- 2) The intermediate phase corresponding to precipitated
proteins ; and
- 3) The top aqueous supernatant phase containing morphine.
The supernatant was collected and dried with a Centrivap Console
(Labconco, Kansas City, Missouri). The dried extract was then
dissolved in 0.05 % trifluoroacetic acid (TFA) water before
solid phase extraction. Samples were loaded on a Waters Sep-Pak
Plus C-18 cartridge previously activated with 100 % acetonitrile
and washed with 0.05 % TFA-water. Morphine elution was performed
with a 10 % acetonitrile solution (water/acetonitrile/ TFA,
89.5 % : 10 % :0.05 %, v/v/v). The eluted sample was dried with
a Centrivap Console and dissolved in water prior to high pressure
liquid chromatography (HPLC) analysis.
Radioimmuno-assay (RIA) determination
The morphine RIA determination is a solid phase, quantitative
RIA, wherein 125I-labeled morphine competes for a fixed time
with morphine in the test sample for the antibody binding site.
The commercial kit employed is from Diagnostic Products Corporation
(USA). Because the antibody is immobilized on the wall of a
polypropylene tube, simply decanting the liquid phase to terminate
the competition and to isolate the antibody-bound fraction of
radiolabeled morphine is sufficient. The material is then counted
in a Wallac, 3", 1480 gamma counter (Perkin Elmer, USA). Comparison
of the counts to a calibration curve yields a measure of the
morphine present in the test sample, expressed as nanograms
of morphine per milliliter. The calibrators contain, respectively,
0, 2.5, 10, 25, 75 and 250 nanograms of morphine per milliliter
(ng/mL) in PBS. Reticuline and salutaridine do not cross-react
with the antibody. The detection limit was 0.5 ng/ml.
HPLC and electrochemical detection of morphine in the sample
The HPLC analyses were performed with a Waters 626 pump (Waters,
Milford, MA) and a C-18 Unijet microbore column (BAS). A flow
splitter (BAS) was used to provide the low volumetric flow-rates
required for the microbore column. The split ratio was 1/9.
Operating the pump at 0.5 ml/min, yielded a microbore column
flow-rate of 50 µl/min. The injection volume was 5 µl. Morphine
detection was performed with an amperometric detector LC-4C
(BAS, West Lafayette, Indiana). The microbore column was coupled
directly to the detector cell to minimize the dead volume. The
electrochemical detection system used a glassy carbon-working
electrode (3mm) and a 0.02 Hz filter (500mV ; range 10 nA).
The cell volume was reduced by a 16-µm gasket. The chromatographic
system was controlled by Waters Millennium32 Chromatography
Manager V3.2 software and the chromatograms were integrated
with Chromatograph software (Waters).
Morphine was quantified in the tissues by the method described
by Zhu and Stefano [52]. This method was carried out in the
following manner : The mobile phases were: Buffer A : 10 mM
sodium chloride, 0.5 mM EDTA, 100 mM sodium Acetate, pH 5.0
; Buffer B : 10 mM sodium chloride, 0.5 mM EDTA, 100 mM sodium
Acetate, 50 % acetonitrile, pH 5.0. The injection volume was
5 ul. The running conditions were : from 0 min 0 % buffer B
; 10 min, 5 % buffer B ; at 25 min 50 % buffer B ; at 30 min
100 % buffer B. Both buffers A and B were filtered through a
Waters 0.22 µm filter and the temperature of the whole system
was maintained at 25°C. Several HPLC purifications were performed
between each sample to prevent residual morphine contamination
remaining on the column. Furthermore, mantle tissue was run
as a negative control, demonstrating a lack of contamination
[55].
Nitric oxide assay
Ten pedal ganglia (per determination) dissected from M.
edulis were bathed in 1 mL sterile phosphate buffered saline
(PBS). Experiments used morphine at a final concentration of
10-6 M, naloxone at 10-6 M, and 1 ug of reticuline. For the
opiate receptor antagonist experiments, ganglia were pretreated
with naloxone for 10 min prior to reticuline addition. NO release
was monitored with an NO-selective microprobe manufactured by
World Precision Instruments (Sarasota, FL). The sensor was positioned
approximately 100 µm above the respective tissue
surface. Calibration of the electrochemical sensor was performed
by use of different concentrations of a nitrosothiol donor S-nitroso-N-acetyl-DL-penicillamine
(SNAP), as previously described by [29]. The
NO detection system was calibrated daily. The probe was allowed
to equilibrate for 10 min in the incubation medium free of tissue
before being transferred to vials containing the ganglia for
another 5 min. Manipulation and handling of the ganglia was
only performed with glass instruments. Data was acquired using
the Apollo-4000 free radical analyzer (World Precision Instruments,
Sarasota, FL). The experimental values were then transferred
to Sigma-Plot and -Stat (Jandel, CA) for graphic representation
and evaluation.
Binding experiments
Human monocytes served as a positive control since the mu3
opiate receptor subtype, which is coupled to NO release, has
been cloned from these cells [7]. The monocytes
were obtained from the Long Island Blood Center (Melville, Long
Island) and processed as previously described in detail [40,
5, 30]. An additional 100
excised pedal ganglia were washed and homogenized
in 50 volumes of 0.32 M sucrose, pH 7.4, at 4°C, by the
use of a Brinkmann polytron (30 s, setting no. 5) as were the
human monocytes. The crude homogenate is centrifuged at 900
x g for 10 min at 4 °C, and the supernatant is reserved
on ice. The whitish crude pellet is resuspended by homogenization
(15 s, setting no. 5) in 30 volumes of 0.32 M sucrose/Tris-HCl
buffer, pH 7.4, and centrifuged at 900 x g for 10 min. The extraction
procedure is repeated one more time, and the combined supernatants
were centrifuged at 900 x g for 10 min. The resulting supernatants
(S1') are used immediately. Prior to the binding experiment,
the S1' supernatant is centrifuged at 30,000 x g for 15 min.
and the pellet (P2) is washed once by centrifugation in 50 volumes
of the sucrose/Tris-HCl. The P2 pellet is then re-suspended
with a Dounce hand-held homogenizer (10 strokes) in 100 volumes
of buffer. Binding analysis is then performed on the cell membrane
suspensions. Aliquots of membrane suspension (0.2 ml, 0.12 mg
of membrane protein) are incubated in triplicate at 22 oC for
40 min with the appropriate radiolabeled ligand in the presence
of dextrorphan (10 mM) or levorphanol (10 mM) in 10 mM Tris-HCl
buffer, pH 7.4, containing 0.1 % BSA and 150 mM KCl. Free ligand
is separated from membrane-bound labeled ligand by filtration
under reduced pressure through GF/B glass fiber filters (Whatman)
; filters were presoaked (45 min, 4 °C)) in buffer containing
0.5 % BSA. The filters are rapidly washed with 2.5 ml aliquots
of the incubation buffer (4 °C), containing 2 % polyethylene
glycol 6000 (Baker). They are assayed by liquid scintillation
spectrometry (Packard 460). Stereospecific binding is defined
as binding in the presence of 10 mM dextrorphan minus binding
in the presence of 10 mM levorphanol. Protein concentration
is determined in membrane suspensions (prepared in the absence
of BSA). For IC50 determination (defined as the concentration
of drug which elicits half-maximal inhibition of specific 3H-dihydromorphine
binding (for mu3), an aliquot of the respective tissue-membrane
suspension is incubated with non-radioactive opioid compounds
at 6 different concentrations for 10 min at 22ºC and then with
3H-dihydromorphine for 60 min at 4ºC as previously noted in
detail [40]. The mean +/- S.E.M. for three experiments is recorded
for each compound. All agents Tyr-D-Pen-Gly-Phe-D-Pen (DPDPE)
and naltrexone are from Sigma Chemical Co. (St.Louis, MO).
Results
Morphine was identified in the ganglionic extraction by reverse
phase HPLC using a gradient of acetonitrile following liquid
and solid extraction, and was compared to an authentic standard
(Figure 1 *).
The material exhibited the same retention time as authentic
morphine, confirming earlier studies that also identified this
material via mass spectrometry (Figure 1 ; [52]).
The electrochemical detection sensitivity of morphine is 80
picograms. The concentration of morphine was 1.43 ± 0.41 ng/mg
± SEM ganglionic wet weight as determined by the Chromatogram
Manager 3.2 commercial software (Millemmium32, Waters, Milford,
MA) extrapolated from the peak-area calculated for the external
standard. Ganglia incubated with 50 ng of reticuline for 1 hour
exhibited a statistical increase in their endogenous morphine
levels (6.7 ± 0.7 ng/mg tissue wet weight ;
P<0.01) (Figure 1 *).
The electrochemical results are compatible with the RIA quantification
(Figure 2 *),
which yields a control ganglionic level of morphine at 1.33
± 0.61 ng/mg tissue wet weight ± SEM. Incubation
with various concentrations of reticuline increases ganglionic
morphine levels after one hour in a concentration dependent
manner (Figure 2 *).
Exposure of excised ganglia to 100 ng of reticuline yields about
14.53 ± 4.6 ng/mg morphine (Figure 2 ; P< 0.001). The increase
in ganglionic morphine levels occurs gradually over the 60 min
incubation period, beginning 10 minutes post reticuline addition
(Figure 3 *).
From these studies, we estimate that approximately 24 % of the
reticuline gets converted to morphine. Blank runs between morphine
HPLC determinations did not show a morphine residue with RIA,
nor did any signs of its presence occur with mantle tissue.
Incubation of 50 ng of reticuline with mantle tissues did not
produce detectable morphine (data not shown).
Previously, we demonstrated that pedal ganglia, which contain
mu opiate receptors, respond to morphine exposure by releasing
constitutive nitric oxide synthase derived NO in a naloxone
and L-NAME sensitive manner [45, 8].
In an attempt to substantiate the identity of newly synthesized
morphine further, ganglia were examined for their ability to
release NO in response to reticuline exposure (Figure 4 *;
Table 1 - *).
We show that reticuline (10-7 M)) does not stimulate ganglionic
NO release in a manner resembling that of morphine (10-6 M),
which releases NO seconds after its application to the ganglia
and lasts for 5 minutes (Figure 4 ; Table 1). Instead, with
reticuline, there is a 3 minute delay, which is followed by
an extended release period occurring over 17 minutes (Figure
4*). We
surmise that this reticuline-stimulated release occurs because
it is being converted to morphine, which is actually responsible
for the release, as indicated by the time course of the morphine
increase, following reticuline exposure (Figure 3 *).
Table 2 *
demonstrates that both reticuline and salutaridine, another
putative morphine precursor [55], do not exhibit
binding affinity for the pedal ganglia mu opiate receptor subtype,
i.e., mu3 [8]. This finding is further substantiated
using the positive control of human monocytes where the mu3
opiate receptor subtype has been cloned [7].
This result strongly suggests that the pre-treatment of the
ganglia with naloxone (10-6 M) blocking the reticuline (10-7
M) stimulated release of NO (Figure 4*)
occurs by way of this precursor being converted to morphine
since reticuline does not have an affinity for the mu opiate
receptor (Table 2*).
Discussion
The present study demonstrates the following :
- 1) Morphine is present in Mytilus pedal ganglia
[40, 43, 53],
confirming earlier reports, including those using mass spectrometry
;
- 2) Exposing pedal ganglia to a putative morphine precursor,
reticuline [46, 55], results
in significant increases in ganglionic morphine levels in
a concentration and time dependent manner ;
- 3) Reticuline stimulates ganglionic NO production, following
a latency period, in a manner consistent with it being converted
to morphine ; and
- 4) Reticuline does not exhibit an affinity for the mu3 opiate
receptor, again suggesting that NO release occurs because
of its conversion into morphine. Taken together, it is apparent
that animals have the ability to synthesize morphine from
reticuline.
Morphine's synthesis, including enzymes and precursors, has
been demonstrated in the plant Papaver somniferum [23,
24, 26, 25].
Morphine, codeine and thebaine have been identified in mammalian
tissues [12, 15, 21,
20, 18], suggesting the
ability for mammals to synthesize the morphine skeleton [50].
This evidence suggests that mammals synthesize morphine in a
manner similar to that of the poppy plant, see [55,
34]. Supporting this view are the findings
demonstrating morphine in various invertebrates, including parasites
and free-living invertebrates identified by
HPLC-coupled to electrochemical detection and Q-TOF-Mass Spec
[17, 54, 16,
27]. We have also found reticuline, tetrahydropapoverine
and codeine in invertebrate tissues along with the morphine
metabolites, morphine-6-glucuronide and -3-glucuronide, providing
additional evidence for this ability in evolutionary diverse
animals, as well as suggesting that this pathway has been conserved
during evolution in both plants and animals [40,
39, 41, 17,
53, 54]. Furthermore, the
newly cloned mu3 opiate receptor subtype, found in human and
invertebrate tissues, only responds to opiate alkaloids as opposed
to opioid peptides, providing the means for endogenous opiate
alkaloid signaling [40], adding additional
support for endogenous morphine signaling. Invertebrate tissues
express mu opiate receptors that are also coupled to NO release
[8]. The lack of mu affinity for reticuline
and salutaridine suggests that these compounds are indeed being
converted to morphine since they cannot release NO directly.
The argument for a de novo biosynthetic pathway in
animals can be supported with studies demonstrating the ability
of animal enzymes to synthesize, through the same precursors,
morphine in an identical stereo- and regio-specific manner to
that of the poppy plant, see [55]. In particular,
reticuline's conversion to morphine in mammalian and invertebrate
tissues, as suggested by the present data, can occur in pigs,
rats, sheep, and cows where the conversion of reticuline to
salutaridine, the next step in the pathway, can be mediated
by cytochrome P450 [1, 2].
This enzyme catalyzes the phenol oxidative coupling reaction
in the same NADPH and O2 dependent manner as that of the plant.
This enzyme is stereo-, regio-, and substrate specific, which
may further demonstrate the ability of animals to synthesize
morphine endogenously.
Thebaine, following salutaridine in the morphine biosynthetic
pathway, also was found in mammalian brain [20].
This is the first morphian alkaloid formed en route to the formation
of morphine. Rat microsomes demonstrated the ability to transform
thebaine to oripavine through a cytochrome P450 2D1 enzyme [38].
Furthermore, morphine dehydrogenase was discovered in mammals
[51]. This enzyme is NADPH-dependent and catalyzes
the oxidation of morphine to morphinone as well as the reverse
reaction [51] that produces morphine, adding
to the evidence of the capability of animal enzymes to catalyze
the necessary reactions to conclude the pathway.
With regard to reticuline, it is found in a variety of plants
[56, 10, 33,
35, 49]. The fruit and bark
of Annonacea family of plants is used as an analgesic, diuretic,
and cough suppressant [9], properties shared
with morphine. Graviola, which also contains reticuline,
is also used as an anti-hypertensive, anti-arthritic, and
anti-diarrhea remedy [9], further supporting
our hypothesis that this compound may yield morphine in animal
tissues since it shares pharmacological actions associated with
morphine.
At the cellular level, reticuline reduces cytosolic calcium
concentrations, inhibiting uterine contractions [32].
Kimura et al. [19] have isolated tetrahydroisoquinoline
alkaloids from Magnolia and have shown that reticuline
decreases contractions in guinea pig papillary muscle. Martin
et al. [31] also have demonstrated the antispasmodic
activity of isoquinoline alkaloids. These researchers speculated
that the antispasmodic effects were due to an inhibition of
inter- and/or extracellular calcium. In our laboratory, we have
demonstrated that NO via morphine or estrogen stimulation can
inhibit muscle contraction [44] and subsequently
influence calcium mobility, as well as actin degranulation,
accounting for the lack of muscle contraction [44].
Hence, via reticuline synthesis to morphine, these actions can
also be attributable to morphine.
Moreover, other researchers have found that reticuline has
antimicrobial, antiviral and molluscicidal properties [28,
36, 35, 13].
This can also be explained by the synthesis of morphine via
reticuline since morphine via the mu3 opiate receptor subtype
is coupled to NO release in brain, immune, vascular and gut
tissues [42, 30, 37,
11, 48]. Thus, these actions
may also be attributable to NO [3, 4],
following morphine synthesis.
Acknowledgements
This work was supported, in part, by the following grants :
NIMH 47392 and DA 09010. The authors wish to express their gratitude
to Dr. Meinhart H. Zenk and Dr. Trevor Robinson for providing
reticuline.
References
1 Amann T, Roos
PH, Huh H, Zenk MH. Purification and characterization of a cytochrome
P450 enzyme from pig liver, catalyzing the phenol oxidative
coupling of (R)-reticuline to salutaridine, the critical step
in morphine biosynthesis. Heterocycles 1995 ; 40
:425-40.
2 Amann T, Zenk
MH. Formation of the morphine precursor salutaridine is catalyzed
by a cytochrome P-450 enzyme mammalian liver. Tetrahedron Letters
1991 ; 32 :3675-8.
3 Benz D, Cadet
P, Mantione K, Zhu W, Stefano GB. Tonal nitric oxide and health
: A free radical and a scavenger of free radicals. Medical Science
Monitor 2002 ; 8 :1-4.
4 Benz D, Cadet
P, Mantione K, Zhu W, Stefano GB. Tonal nitric oxide and health
: Anti-bacterial and -viral actions and implications for HIV.
Medical Science Monitor 2002 ; 8 :RA27-RA31.
5 Bilfinger TV,
Fricchione GL, Stefano GB. Neuroimmune implications of cardiopulmonary
bypass. Adv Neuroimmunol 1993 ; 3 :277-88.
6 Brochmann-Hanssen
E. Biosynthesis of morphinan alkaloids. In : Phillipson JD,
Roberts MF, Zenk MH, editors. The Chemistry and Biology of Isoquinoline
Alkaloids. Berlin, Heidelberg : Springer-Verlag ; 1985.p. 229-39.
7 Cadet P, Mantione
KJ, Stefano GB. Molecular identification and functional expression
of mu3, a novel alternatively spliced variant of the human mu
opiate receptor gene. Journal of Immunology 2003 ; 170
:5118-23.
8 Cadet P, Stefano
GB. Mytilus edulis pedal ganglia express µ opiate receptor
transcripts exhibiting high sequence identity with human neuronal
µ1. Mol Brain Res 1999 ; 74 :242-6.
9 Carbajal D,
Casaco A, Arruzazabala L, Gonzalez R, Fuentes V. Pharmacological
screening of plant decoctions commonly used in Cuban folk medicine.
J Ethnopharmacol 1991 ; 33 :21-4.
10 Chen IS, Chen
JJ, Duh CY, Tsai IL, Chang CT. New aporphine alkaloids and cytotoxic
constituents of Hernandia nymphaeifolia. Planta Med 1997 ; 63
:154-7.
11 de la Torre
JC, Pappas BA, Prevot V, Emmerling MR, Mantione K, Fortin T
et al. Hippocampal nitric oxide upregulation precedes memory
loss and A beta I-40 accumulation after chronic brain hypoperfusion
in rats. Neurological Research 2003 ; 25 :635-41.
12 Donnerer J,
Oka K, Brossi A, Rice KC, Spector S. Presence and formation
of codeine and morphine in the rat. Proc Natl Acad Sci USA 1986
; 83 :4566-7.
13 dos Santos
AF, Sant'Ana AE. Molluscicidal properties of some species of
Annona. Phytomedicine 2001 ; 8 :115-20.
14 Epple A, Nibbio
B, Spector S, Brinn J. Endogenous codeine : autocrine regulator
of catecholamine release from chromaffin cells. Life Sci 1994
; 54 :695-702.
15 Goldstein
A, Barrett RW, James IF, Lowney LI, Weitz C, Knipmeyer LI et
al. Morphine and other opiates from beef brain and adrenal.
Proc Natl Acad Sci USA 1985 ; 82 :5203-7.
16 Goumon Y,
Casares F, Pryor S, Ferguson L, Brownwell B, Cadet P et al.
Ascaris suum, an internal parasite, produces morphine.
J Immunol 2000 ; 165 :339-43.
17 Goumon Y,
Casares F, Zhu W, Stefano GB. The presence of morphine in ganglioic
tissues of Modiolus deminissus : A highly sensitive
method of quantitation for morphine and its derivatives. Mol
Brain Res 2001 ; 86 :184-8.
18 Goumon Y,
Stefano GB. Identification of Morphine in the Rat Adrenal Gland.
Mol Brain Res 2000 ; 77 :267-9.
19 Kimura I,
Chui LH, Fujitani K, Kikuchi T, Kimura M. Inotropic effects
of (+/-)-higenamine and its chemically related components, (+)-R-coclaurine
and (+)-S-reticuline, contained in the traditional sino-Japanese
medicines "bushi" and "shin-i" in isolated guinea pig papillary
muscle. Jpn J Pharmacol 1989 ; 50 :75-8.
20 Kodaira H,
Listek CA, Jardine I, Arimura A, Spector S. Identification of
the convusant opiate thebaine in the mammalian brain. Proc Natl
Acad Sci,USA 1989 ; 86 :716-9.
21 Kodaira H,
Spector S. Transformation of thebaine to oripavine, codeine,
and morphine by rat liver,kidney, and brain microsomes. Proc
Natl Acad Sci,USA 1988 ; 85 :1267-71.
22 Lee SC, Spector
S. DON'T USE Changes in Endogenous Morphine and Codeine contents
in the fasting rat. Journal of Pharmacology & Experimental Therapeutics
1991 ; 257 :647-52.
23 Lenz R, Zenk
MH. Closure of the oxide bridge in morphine biosynthesis. Tetrahedron
Letters 1994 ; 35 :3897-900.
24 Lenz R, Zenk
MH. Acetyl coenzyme A :salutaridinol-7-O-acetyltransferase from
papaver somniferum plant cell cultures. The enzyme catalyzing
the formation of thebaine in morphine biosynthesis. J Biol Chem
1995 ; 270 :31091-6.
25 Lenz R, Zenk
MH. Purification and properties of codeinone reductase (NADPH)
from Papaver somniferum cell cultures. European Journal
of Biochemistry 1995 ; 233 :132-9.
26 Lenz R, Zenk
MH. Stereospecific reduction of codeinone, the penultimate enzymatic
step during morphine biosynthesis in Papaver somniferum.
Tetrahedron Letters 1995 ; 36 :2449-52.
27 Leung MK,
Dissous C, Capron A, Woldegaber H, Duvaux-Miret O, Pryor SC
et al. Schistosoma mansoni : The presence and potential
use of opiate-like substances. Exp Parasit 1995 ; 81
:208-15.
28 Liao WT, Beal
JL, Wu WN, Doskotch RW. Alkaloids of Thalictrum XXVI. New hypotensive
and other alkaloids from Thalictrum minus race B. Lloydia 1978
; 41 :257-70.
29 Liu Y, Shenouda
D, Bilfinger TV, Stefano ML, Magazine HI, Stefano GB. Morphine
stimulates nitric oxide release from invertebrate microglia.
Brain Res 1996 ; 722 :125-31.
30 Magazine HI,
Liu Y, Bilfinger TV, Fricchione GL, Stefano GB. Morphine-induced
conformational changes in human monocytes,granulocytes, and
endothelial cells and in invertebrate immunocytes and microglia
are mediated by nitric oxide. J Immunol 1996 ; 156
:4845-50.
31 Martin ML,
Diaz MT, Montero MJ, Prieto P, San Roman L, Cortes D. Antispasmodic
activity of benzylisoquinoline alkaloids analogous to papaverine.
Planta Med 1993 ; 59 :63-7.
32 Martin ML,
Sagredo JA, Morais JM, Montero MJ, Sanchez MT, San Roman L.
Uterine inhibitory effect of reticuline. J Pharm Pharmacol 1988
; 40 :801-2.
33 Morais LC,
Barbosa-Filho JM, Almeida RN. Central depressant effects of
reticuline extracted from Ocotea duckei in rats and mice. J
Ethnopharmacol 1998 ; 62 :57-61.
34 Neri C, Guarna
M, Bianchi E, Sonetti D, Matteucci G, Stefano GB. Endogenous
morphine and codeine in the brain of non-human primate. Medical
Science Monitor 2004 ; 10 :MS1-MS5.
35 Padma P, Pramod
NP, Thyagarajan SP, Khosa RL. Effect of the extract of Annona
muricata and Petunia nyctaginiflora on Herpes simplex virus.
J Ethnopharmacol 1998 ; 61 :81-3.
36 Paulo MQ,
Barbosa-Filho JM, Lima EO, Maia RF, Barbosa RC, Kaplan MA. Antimicrobial
activity of benzylisoquinoline alkaloids from Annona salzmanii
D.C. J Ethnopharmacol 1992 ; 36 :39-41.
37 Prevot V,
Rialas C, Croix D, Salzet M, Dupouy J-P, Puolain P et al. Morphine
and anandamide coupling to nitric oxide stimulated GnRH and
CRF release from rat median eminence : neurovascular regulation.
Brain Res 1998 ; 790 :236-44.
38 Sindrup SH,
Poulsen L, Brosen K, Arendt-Nielsen L, Gram LF. Are poor metabolisers
of sparteine/debrisoquine less pain tolerant than extensive
metabolisers? Pain 1993 ; 53 :335-9.
39 Sonetti D,
Mola L, Casares F, Bianchi E, Guarna M, Stefano GB. Endogenous
morphine levels increase in molluscan neural and immune tissues
after physical trauma. Brain Res 1999 ; 835
:137-47.
40 Stefano GB,
Digenis A, Spector S, Leung MK, Bilfinger TV, Makman MH et al.
Opiatelike substances in an invertebrate, a novel opiate receptor
on invertebrate and human immunocytes, and a role in immunosuppression.
Proc Natl Acad Sci USA 1993 ; 90 :11099-103.
41 Stefano GB,
Goumon Y, Casares F, Cadet P, Fricchione GL, Rialas C et al.
Endogenous morphine. Trends in Neurosciences 2000 ; 9
:436-42.
42 Stefano GB,
Hartman A, Bilfinger TV, Magazine HI, Liu Y, Casares F et al.
Presence of the mu3 opiate receptor in endothelial cells : Coupling
to nitric oxide production and vasodilation. J Biol Chem 1995
; 270 :30290-3.
43 Stefano GB,
Leung MK, Bilfinger TV, Scharrer B. Effect of prolonged exposure
to morphine on responsiveness of human and invertebrate immunocytes
to stimulatory molecules. J Neuroimmunol 1995 ; 63
:175-81.
44 Stefano GB,
Murga J, Benson H, Zhu W. Nitric oxide inhibits norepinephrine
stimulated contraction of human internal thoracic artery and
rat aorta. Pharmacol Res 2001 ; 43 :199-203.
45 Stefano GB,
Salzet B, Rialas CM, Pope M, Kustka A, Neenan K et al. Morphine
and anandamide stimulated nitric oxide production inhibits presynaptic
dopamine release. Brain Res 1997 ; 763 :63-8.
46 Stefano GB,
Scharrer B. Endogenous morphine and related opiates, a new class
of chemical messengers. Adv Neuroimmunol 1994 ; 4
:57-68.
47 Stefano GB,
Teoh M, Grant A, Reid C, Teoh H, Hughes TK. In vitro effects
of electromagnetic fields on immunocytes. Electro-Magnetobiol
1994 ; 13 :123-36.
48 Stefano GB,
Zhu W, Cadet P, Mantione K. Morphine enhances nitric oxide release
in the mammalian gastrointestinal tract via the m3 opiate receptor
subtype : A hormonal role for endogenous morphine. Journal of
Physiology and Pharmacology 2004 ; 55 :279-88.
49 Watanabe H,
Ikeda M, Watanabe K, Kikuchi T. Effects on central dopaminergic
systems of d-coclaurine and d-reticuline, extracted from Magnolia
salicifolia. Planta Med 1981 ; 42 :213-22.
50 Weitz CJ,
Lowney LI, Faull KF, Feister G, Goldstein A. Morphine and codeine
from mammalian brain. Proc Natl Acad Sci,USA 1986 ; 83
:9784-8.
51 Yamano S,
Kageura E, Ishida T, Toki S. Purification and charcterization
of guinea pig liver morphine 6-dehydrogenase. Journal of Biological
Chemistry 1985 ; 260 :5259-64.
52 Zhu W, Baggerman
G, Goumon Y, Casares F, Brownawell B, Stefano GB. Presence of
morphine and morphine-6-glucuronide in the marine mollusk Mytilus
edulis ganglia determined by GC/MS and Q-TOF-MS. Starvation
increases opiate alkaloid levels. Brain Res Mol Brain Res 2001
; 88 :155-60.
53 Zhu W, Baggerman
G, Goumon Y, Casares F, Brownwell B, Stefano GB. Presence of
morphine and morphine-6-glucuronide in the marine mollusk Mytilus
edulis ganglia determined by GC/MS and Q-TOF-MS. Starvation
increases opiate alkaloid levels. Mol Brain Res 2001 ; 88
:155-60.
54 Zhu W, Baggerman
G, Secor WE, Casares F, Pryor SC, Fricchione GL et al. Dracunculus
medinensis and Schistosoma mansoni contain opiate
alkaloids. Annals of Tropical Medicine and Parasitology 2002
; 96 :309-16.
55 Zhu W, Ma
Y, Cadet P, Yu D, Bilfinger TV, Bianchi E et al. Presence of
reticuline in rat brain : A pathway for morphine biosynthesis.
Mol Brain Res 2003 ; 117 :83-90.
56 Zhu W, Ma
Y, Stefano GB. Presence of isoquinoline alkaloids in molluscan
ganglia. Neuroendocrinol Lett 2002 ; 23 :329-34.
* in
process - Publisher's comment
|
