Department of Pediatrics, Yale University, New Haven, Connecticut 06520
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ABSTRACT |
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We recently
demonstrated that -opioid receptor (DOR) activation protects
cortical neurons against glutamate-induced injury. Because glutamate is
a mediator of hypoxic injury in neurons, we hypothesized that DOR is
involved in neuroprotection during O2 deprivation and that
its activation/inhibition may alter neuronal susceptibility to hypoxic
stress. In this work, we tested the effect of opioid receptor
activation and inhibition on cultured cortical neurons in hypoxia (1%
O2). Cell injury was assessed by lactate dehydrogenase
release, morphology-based quantification, and live/dead staining. Our
results show that 1) immature neurons (days 4 and
6) were not significantly injured by hypoxia until 72 h
of exposure, whereas day 8 neurons were injured after only 24-h hypoxia; 2) DOR inhibition (naltrindole) caused
neuronal injury in both day 4 and day 8 normoxic
cultures and further augmented hypoxic injury in these neurons;
3) DOR activation
([D-Ala2,D-Leu5]enkephalin)
reduced neuronal injury in day 8 cultures after 24 h of
normoxic or hypoxic exposure and attenuated naltrindole-induced injury
with prolonged exposure; and 4) µ- or
-opioid receptor inhibition (
-funaltrexamine or nor-binaltorphimine) had little effect on neurons in either normoxic or hypoxic conditions.
Collectively, these data suggest that DOR plays a crucial role in
neuroprotection in normoxic and hypoxic environments.
cortex; hypoxia; injury; protection; opioids
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INTRODUCTION |
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MAMMALIAN CORTICAL NEURONS are highly sensitive to hypoxic stress. Depending on severity and duration, hypoxia can lead to major alterations in neuronal function, metabolism, and morphology (18, 25). For example, prolonged hypoxic exposure may cause irreversible neuronal injury or even cell death, thus leading to serious neurological disorders (18). However, the cellular and molecular mechanisms of hypoxic injury and adaptation in neurons are not clearly understood.
Our recent studies (50) suggest that activation of the
-opioid receptor (DOR) system protects cortical neurons against glutamate-induced stress, whereas activation of µ- or
-opioid receptors offers no significant neuroprotection. Because glutamate is a
mediator of hypoxic/ischemic injury (8, 16) and
endogenous opioids are released during hypoxia (2, 3, 15,
46), we predicted that DOR might also play a role in
neuroprotection during hypoxic stress. However, there is no direct
evidence showing the effect of DOR on neuronal responses to
O2 deprivation. In whole animal experiments, systemic
administration of DOR agonists prolonged the survival period of mice
during hypoxia (5, 23, 24). Because of the complex effects
of drug administration in the whole body, the targets of DOR-activated
protection are not clearly defined. For example, systemic DOR agonists
may act on cardiac DOR, which is protective against ischemic
stress (1, 32).
On the basis of our previous work, we hypothesized that neuronal DOR is
involved in self-protection and that its inhibition leads to increased
neuronal injury in response to hypoxia. To test this hypothesis, we
exposed cultured neurons to hypoxia and determined the effect of DOR
activation and inhibition on neuronal responses to O2
deprivation. The specificity of DOR was also examined by assessing the
effect of µ- and -opioid receptor antagonists on the same neuronal
cultures. Because our past investigations (44, 45) and
those of others (41) demonstrated a high density of DORs
in the mammalian cortex, we focused the present investigation on
cortical neurons.
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METHODS |
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Animals. Pregnant (embryonic day 16-17) Sprague-Dawley rats were purchased from Charles River Laboratories (Wilmington, MA). All animal procedures were performed in accordance with the guidelines of the Animal Care Committee of Yale University School of Medicine, which is accredited by the American Association for Accreditation of Laboratory Animal Care.
Preparation of neuronal cultures. Primary cultures of cortical neurons from embryonic day 16-17 rats were used as previously described (50). In brief, fetuses were decapitated and cortical tissue was collected under sterile conditions. The tissue was dispersed with a 1-ml pipette and then passed through an 80-µm nylon mesh with a Teflon pestle. Cells were resuspended in neuron-defined, serum-free Neurobasal medium (GIBCO-BRL, Grand Island, NY), supplemented with B-27, glutamine (0.5 mM), glutamate (25 µM), and a combination of penicillin (100 IU/ml) and streptomycin (100 µg/ml). The cells were plated onto poly-D-lysine (100 µg/ml; Sigma, St. Louis, MO)-coated 35-mm culture dishes at 1 × 106 cells/ml. The culture dishes were then kept in a humidified atmosphere of 95% air and 5% CO2 at 37°C.
Hypoxic induction of rat cortical neurons. Neurons were exposed to hypoxia beginning at day 4, 6, or 8 for durations of 0.5, 4, 8, 24, 48, and 72 h. Culture dishes were randomly divided into two groups, one in a normoxic incubator and the other in an incubator maintained at 1% O2, 5% CO2, and 94% N2.
Cell treatment.
Culture dishes were treated with
[D-Ala2,D-Leu5]enkephalin
(DADLE, a highly selective DOR agonist; Refs. 20, 50),
naltrindole (NTI, a highly selective DOR antagonist; Ref.
28), -funaltrexamine (FNA, a selective µ-opioid
receptor antagonist; Refs. 38, 43), or
nor-binaltorphimine (BNI, a selective
-opioid receptor antagonist; Ref. 6). DADLE, NTI, FNA, or BNI was added to cultures at
a final concentration of 10 µM, and then they were immediately
exposed to either normoxia or hypoxia. For long-term exposures (72 h), supplemental doses were added to the culture medium after 36 h. NTI, FNA, and BNI were purchased from RBI (Natick, MA), and DADLE was
purchased from Sigma. After 24-72 h of exposure, neurons were assessed for neuronal injury via assay of lactate dehydrogenase (LDH)
release, morphology-based quantification, and/or viability/cytotoxicity assay (Molecular Probes, Eugene, OR; see Live/dead
assay).
LDH assay. LDH activity in the culture medium was measured with an LDH kit (Sigma Diagnostics Procedure no. 228-UV) and a Beckman DU-640 spectrophotometer system (Beckman Instruments, Fullerton, CA). Culture medium was sampled and centrifuged to remove cellular debris from the supernatant. Subsequently, 100 µl of the sample was added to a polystyrene cuvette containing 1 ml of LDH reagent (50 mM lactate and 7 mM NAD+ in 0.05% sodium azide buffer, pH 8.9). The cuvette was placed immediately into the spectrophotometer, maintained at 30°C. After stabilization for 30 s, absorbance at 340 nm was recorded at 30-s intervals for 2 min. The change in absorbance was then expressed in concentration units (U) per liter and converted to percentage of control levels.
Morphological studies. For the qualitative assessment of viable and injured neurons, a "same-field" quantification method, developed in our laboratory (50), was used with a computer-based image analysis system. In brief, microphotographs of cultured cells were taken before experimental treatment, using a phase-contrast microscope to establish baseline viability. After experimental treatment, the same field of each culture dish was reexamined and photomicrographs were taken again. Viable and injured neurons were compared in the same field before and after experiments to assess cell injury. The criteria for neuronal injury were the same as in our previous work (50).
Live/dead assay. Neuronal survival was quantified using a live/dead viability/cytotoxicity kit (L-3224) from Molecular Probes. In accordance with the manufacturer's protocol, neurons were exposed to cell-permeant calcein AM (3 µM), which is hydrolyzed by intracellular esterases, and to ethidium homodimer-1 (4 µM), which binds to nucleic acids. (The cleavage product of calcein AM, calcein, produces a green fluorescence when exposed to 494-nm light and is used to identify live cells. Bound ethidium homodimer-1 produces a red fluorescence when exposed to 528-nm light, allowing the identification of dead cells.) Culture dishes were dually stained and examined under a fluorescence microscope system (Zeiss Axiovert 25; Sony Progressive 3CCD and Camera Adapter CMA-D2; blue excitation filter: 488/515 nm, green excitation filter: 514/550 nm). Neuronal viability was determined from five random fields per dish, averaged and expressed as percent cell survival, i.e., [live cells/(dead + live cells)] × 100.
Data analysis. The data are expressed as means ± SE and were subjected to statistical analysis via nonpaired, two-tailed Student's t-test with GraphPad Prism 3.0 software (GraphPad Software, San Diego, CA). The level of statistical significance was set at P < 0.05.
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RESULTS |
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Hypoxic susceptibility increases with neuronal maturation and
stress duration.
Because responses to hypoxic stress are dependent on hypoxic duration
and neuronal maturation (18), we determined the effect of
these factors on neuronal injury in our culture system. We exposed the
cortical neurons at day 4, 6, or 8 to
hypoxia (1% oxygen) for 0.5-72 h and assessed hypoxia-induced
injury by measuring LDH release into the medium. The method of LDH
assay provided the possibility of studying dynamic changes in the same
culture dish. In day 4 neurons, no substantial injury was
observed during the first 48 h of hypoxic exposure, although a
significant increase in medium LDH occurred after 72 h of exposure
(30% increase; n = 16; Fig.
1). A slightly different pattern was
observed in day 6 neurons. After the first 24 h of
exposure, a small increase in LDH levels was observed, and by 72 h
LDH activity increased by 50% (n = 14; Fig. 1). In
contrast, day 8 neurons experienced elevated medium LDH
levels after only 8 h of hypoxia, and these levels rose
significantly after 24 h (30% increase; n = 9;
Fig. 1). The greatest increase in LDH leakage in these neurons was observed after 72-h hypoxia (100% increase; n = 15),
which was 2.5-fold greater than that of day 4 neurons
exposed to the same hypoxic duration. Furthermore, our same-field
morphological data (see Figs. 4C-C' and 6C-C')
demonstrated more observable injury due to hypoxic stress in day
8 cultures than in day 4 cultures. These
observations confirm our LDH results in that day 8 neurons are more susceptible to hypoxic insult than younger neurons.
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DOR activation reduces neuronal injury in hypoxic conditions.
Our previous work (50) showed that DOR activation greatly
reduces glutamate-induced injury in day 8 cortical neurons.
Because hypoxia induced significant injury in day 8 neurons
after 24 h of exposure, as shown in Hypoxic susceptibility
increases with neuronal maturation and stress duration, we used
the same method of DOR activation (50) to test whether DOR
protects these neurons from hypoxic stress. First, we determined the
effect of DOR activation on neuronal survival under hypoxia with a
live/dead staining assay. Because of the accumulation of cell death
beginning with neuronal preparation and plating through day
9 in culture, an average base level of 67% neuronal viability was
seen in a given field in control cultures (day 9, Figs.
2A-A' and
3A). Neuronal cultures
exposed to hypoxia (1% O2) for 24 h significantly
decreased survival to 59% (Figs. 2B-B' and 3A).
DADLE treatment before hypoxic exposure completely abolished the
hypoxia-induced neuronal injury (Figs. 2C-C' and
3A; P < 0.05 compared with hypoxia alone).
To further ascertain the protective effects of DOR activation, we
measured LDH release in day 8 cultures exposed to 24-h
hypoxia and 24-h hypoxia plus DADLE. As shown in Fig. 3B,
LDH release increased significantly after 24-h hypoxia. This hypoxic
elevation of LDH release was abolished by concurrent treatment with
DADLE. These data strongly suggest that DOR activation protects
cortical neurons against hypoxic stress. It is interesting to note that
addition of DADLE to the same neuronal cultures exposed to prolonged
hypoxia (48-72 h) provided no appreciable neuroprotection.
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DOR inactivation increases hypoxia-induced neuronal injury.
Because DOR is neuroprotective against hypoxia, we further asked
whether its inhibition leads to an increase in neuronal injury during
hypoxia. We also questioned whether DOR protection is dependent on
neuronal age, because DOR density is much greater in day 8 than in day 4 cortical neurons (49). Therefore,
we compared day 4 and day 8 neurons in terms of
their responses to DOR inhibition. Day 4 neurons
demonstrated observable morphological changes with same-field
microscopy after 72 h of normoxia with NTI administration (Fig.
4). Furthermore, hypoxia-induced neuronal
injury was amplified with NTI treatment. These findings were confirmed
by LDH release measurements. As depicted in Fig. 5
A, medium LDH levels increased significantly in normoxic day 4 cultures after NTI
administration for 72 h (50% increase; n = 15;
P < 0.0001). In addition, exposure of NTI-treated
day 4 neurons to 72-h hypoxia further raised medium LDH
levels by 75% (n = 14; P < 0.0001;
Fig. 5B), a 1.5-fold greater increase than in normoxia.
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DOR agonists reduce neuronal injury induced by DOR inactivation.
To further clarify the role of DOR in neuroprotection, we questioned
whether the NTI effect could be attenuated by an increase in DOR
agonists in culture medium. Again, we took advantage of the LDH assay,
i.e., dynamic studies of neuronal injury with hypoxic time from the
same neuronal culture. As shown in Fig. 8
A, DOR inhibition in normoxic
conditions induced a significant rise in medium LDH levels after only
24 h (15% increase; n = 12; P < 0.05). Although continued exposure for 48 h resulted in
considerably more LDH release (40% increase; n = 9;
P < 0.05), prolonged exposure for 72 h showed the
most dramatic rise in LDH. In hypoxic neurons, exposure to NTI
greatly elevated medium LDH levels (in comparison to hypoxic controls)
after only 24 h of treatment (30% increase; n = 10; P < 0.05) and further increased levels with more
prolonged exposure durations (Fig. 8B).
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µ- and -opioid receptor inactivation causes minor neuronal
injury.
To test the specificity of DOR with respect to its role in neuronal
responses to hypoxia, µ- and
-opioid receptor antagonists (FNA and
BNI, respectively) were also used in the treatment of cultured neurons
in both normoxic and hypoxic conditions for 72 h. LDH measurements
revealed that treatment with either FNA or BNI at 10 µM induced a
very slight increase in medium LDH levels in both normoxic day 4 (Fig. 5) and day 8 (Fig. 7) neurons. Treatment with FNA
yielded a 9% and 13% increase in LDH levels in day 4 and
day 8 neurons, respectively, whereas BNI exposure resulted in somewhat smaller rises in LDH for both groups. Furthermore, a
comparable pattern was seen under hypoxic conditions. As demonstrated in Figs. 5 and 7, exposure to FNA produced a 25-30% increase in LDH release over hypoxia alone in day 4 and day 8 neurons and BNI yielded a 15-25% increase in these neurons. All
changes were significantly smaller than those seen in NTI-treated neurons.
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DISCUSSION |
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The major findings of the present study are 1)
hypoxia-induced injury is dependent on hypoxic duration and neuronal
age in cultured cortical neurons; 2) DOR activation is
neuroprotective in both normoxic and hypoxic neurons; and 3)
DOR inhibition, but not µ- or -opioid receptor inhibition, induces
major neuronal injury in normoxic conditions and further increases
injury during hypoxia, which is attenuated by DOR agonist.
We used three independent approaches to determine neuronal injury in this work, i.e., LDH assay, morphology-based same-field quantification, and live/dead staining. Among these approaches, LDH assay has a unique advantage, i.e., the use of a small sample for dynamic study of the same culture dish, which provides relatively more accurate time course data for either hypoxic duration- or DOR treatment-based experiments. The release of intracellular LDH into extracellular fluids has been documented as a reliable indicator of neuronal injury (19, 22, 50). In past investigations (50), we observed that glutamate injury in cultured cortical neurons was determined equally by assaying LDH release or same-field morphology. Other works have also demonstrated that the results of LDH release assays parallel those of phase-contrast microscopy, trypan blue staining, and fluorescent staining (19, 22). However, the use of LDH release as an index of neuronal injury does not provide direct assessment of the percentage of neurons injured and/or dead in culture. Therefore, we conducted additional experiments, including morphological assessment and live/dead staining, to assess neuronal injury more directly. The consistency of our results for all assays further confirms the reliability of our data in terms of hypoxia and the effect of opioid receptor activation/inhibition on neuronal injury.
The present study draws a detailed time course for hypoxia-induced injury in cultured cortical neurons based on neuronal age and hypoxic duration, demonstrating that cultured cortical neurons become less tolerant to hypoxic stress with further maturation. This is similar to what is seen in newborn and adult brain tissues (9, 17, 18). Interestingly, our results indicate a major increase in hypoxic sensitivity between culture day 6 and day 8 in cortical neurons. This is also true in cultured hippocampal neurons, which exhibit increased hypoxic susceptibility after 7 days in vitro (13). It seems, therefore, that a critical developmental transition occurs in cultured neurons after 7 days in terms of neuronal vulnerability to stress. A variety of factors are believed to be related to this phenomenon (18, 25), including membrane protein expression, metabolism, and release of excitatory amino acids. One of the major factors that influence neuronal susceptibility may be attributed to glutamate release and receptor expression. During hypoxic exposure, glutamate is expelled from neurons (8, 47), resulting in the overstimulation of glutamate receptors and subsequent cell injury or death (7, 18). Because glutamate receptor expression increases during development (31) and sensitivity to glutamate excitotoxicity increases with neuronal maturation (50), the observed differences in hypoxic susceptibility between neuronal ages may be associated with the developmental increase in glutamate toxicity.
In this work, we demonstrated that stimulation of DOR reduces neuronal
injury after 24-h treatments in either normoxic or hypoxic conditions
but does not yield any substantial benefit with prolonged exposure
durations. A possible explanation for this phenomenon is that prolonged
hypoxia may cause a significant accumulation of opioid release, which
saturates DORs in these neurons. Past investigations showed that
opioids are present at significant levels in bovine glial cultures and
cerebral spinal fluid (2, 3). In response to short-term
hypoxia, the level of enkephalins, which are endogenous agonists for
- and µ-opioid receptors, sharply increases. Other works have
shown similar findings with different models (15, 46).
From these findings, it appears that cortical neurons may release
opioids during normal function and in response to hypoxic stress as a
mechanism of self-protection against injury. Because high levels of
endogenous opioids may already have been present in the culture medium
after prolonged hypoxia, adding more DOR agonist may not have further
increased DOR protection. On the other hand, desensitization of DOR may also have occurred because of the prolonged treatment with DOR agonist in conjunction with endogenous opioid release during chronic hypoxia. Indeed, past investigations showed that chronic DOR agonist treatment of NG108 and C6 gliomal cell lines results in DOR
downregulation via expression and activity (29, 48).
On the basis of the above discussion, it is reasonable to predict that cortical neurons are deeply susceptible to DOR inhibition, which causes serious neuronal injury, especially during hypoxic stress. In fact, our results strongly support this prediction. Interestingly, greater injury was observed in day 8 neurons than in day 4 neurons in both normoxic and hypoxic conditions, which suggests that maturational differences exist between these age groups. We previously observed (44, 49) in the brain and in cultured neurons that DOR expression increases significantly with development. Because DOR density of day 8 neurons is more than twofold that of day 4 neurons (49), these more mature neurons may have a greater dependence on this pathway to maintain neuronal function and, therefore, may be more susceptible to neuronal injury with DOR inhibition. Another noteworthy point is that DADLE did not increase protection on neurons subjected to prolonged hypoxia but significantly attenuated NTI-induced injury in the same neurons. This observation suggests that the increase in DOR agonist may compete with DOR antagonist in terms of DOR binding and thus reduce neuronal injury induced by DOR inhibition during prolonged hypoxia.
The question arises of why -, but not µ- or
-, opioid receptors
are involved in neuroprotection. In addressing this issue, the
inhibitory efficacies of the three antagonists used should first be
considered. Previous reports showed that NTI, FNA, and BNI have
extremely high binding affinities, with dissociation constant
(Kd) and IC50 values in the
nanomolar to picomolar range, to their respective receptors (6,
14, 38, 43). Because a final concentration of 10 µM was used
for each antagonist, it may be assumed that each type of opioid
receptor was sufficiently saturated by antagonist, suggesting that
differences in neuroprotective effects are less likely associated with
inhibition efficacy. However, we could not rule out the possibility
that FNA or BNI binds to DOR at the concentration used in this study.
In fact, it seems possible that these µ- and
-specific antagonists
may have yielded a slight inhibition of DOR, an occurrence that has
been observed in other works (37, 43). If these
antagonists did partially block DOR, it would account for the minor
neuronal injury that was observed in FNA- and BNI-treated neurons. A
second issue to address is whether differences in expression levels of
the various opioid receptors account for the observed phenomenon in
cortical neurons. Past studies demonstrated that µ-opioid receptors
are present at similar or even higher densities than DOR in the
mammalian cortex, although
-opioid receptor density is slightly
lower (33, 41, 44, 45). This implies that the relative
distribution/expression levels of opioid receptor subtypes within the
cortex, as a whole, may not be a key factor in the observations shown
in this work. In light of our previous work (45, 50), we
are confident that DOR, but not µ- or
-opioid receptors, plays an
important role in neuronal survival in normoxic conditions and during
environmental stress.
The mechanism(s) of DOR-mediated neuroprotection is unknown at present. It may involve the regulation of specific G proteins, ion channels (e.g., Ca2+ and K+ channels), and excitatory neurotransmitter release. For instance, studies at the cellular level showed that DOR activation inhibits calcium currents by restricting N-type Ca2+ channels in NG108-15 cells (40) and can also inhibit voltage-gated L-type Ca2+ channels in GH3 pituitary cells (27). Because intracellular Ca2+ levels are elevated during hypoxic exposure, leading to irreversible cell injury (18, 34), the inhibition of Ca2+ currents by DOR stimulation may serve as a neuroprotective mechanism in prevention of Ca2+ overload. In addition, DOR regulation of glutamate signaling may also be involved in normal function and protection of neurons. Immunolabeling studies have shown that DOR is localized at presynaptic and postsynaptic terminals in a variety of neurons, including those of the mammalian cortex (4, 10, 36, 42). At the axon terminal, it has been proposed that blockage of Ca2+ currents by DOR stimulation prevents the release of glutamate from presynaptic vesicles, thereby reducing glutamate excitability (26, 39). Alternatively, past studies also found that DOR may interact with glutamate receptors on the postsynaptic membrane and suppress neuronal signaling (35, 39). In either situation, DOR apparently has the ability to reduce neuronal overstimulation via inhibition of glutamate excitation. Such regulation may be utilized during normal cell functioning and in response to environmental stresses. Therefore, one of the reasons why DOR inhibition induced substantial injury in normoxic neurons may be the loss of inhibitory regulation of excitatory neurotransmitter release and/or receptor excitation.
In general, -, µ- and
-opioid receptors have many similarities,
such as consisting of seven transmembrane domains, existing as ~60%
identical sequences, and being coupled to G proteins (20). Similarities also exist in several of their regulatory targets, including adenylyl cyclases, protein kinases, and certain
Ca2+ and K+ channels. However, DOR seems to
play a unique role in neuroprotection. One possible explanation for the
difference in neuroprotective capabilities is that individual opioid
receptors regulate different effectors, thereby eliciting different
responses. Because of the common features of opioid receptors, it was
proposed by Connor and Christie (11) that the selectivity
of these receptors for eliciting specific pathways does not lie in
differences between each opioid receptor subtype but in their
association with divergent types of G proteins. Included in this
statement are observations that demonstrate that each opioid receptor
subtype preferentially couples to specific G proteins, such as where
DOR is more efficiently coupled to the G16 protein than
either µ- or
-opioid receptors (21). Whatever the
mechanistic pathway is, it apparently yields specialized differences
between receptor subtypes and the regulation of effectors. For example,
it was shown that µ-opioid receptors, but not
- or
-opioid
receptors, are involved in the regulation of Ca2+ channels
in neurons isolated from the nucleus tractus solitarii of rats and in
mouse periaqueductal gray neurons (12, 30).
In conclusion, the present data clearly show that DOR modulation, but
not that of µ- and -opioid receptors, plays a major role in
neuroprotection in both normoxic and hypoxic environments. Although the
related mechanisms are unknown at present, we speculate that this
phenomenon may be linked to the role of DOR in selective regulation of
G proteins, excitatory neurotransmitter release, glutamate receptor
stimulation, and Ca2+ homeostasis. Because the activation
of DOR may have a wide range of clinical applications in treating acute
hypoxia-related impairments, further research is warranted to develop a
better understanding of opioid receptors and the pathways involved in neuroprotection.
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ACKNOWLEDGEMENTS |
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This work was supported by March of Dimes (FY00-722) and National Institute of Child Health and Human Development (R01-HD-34852) grants to Y. Xia.
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FOOTNOTES |
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* J. Zhang and G. T. Gibney contributed equally to this work.
Address for reprint requests and other correspondence: Y. Xia, Yale Univ. School of Medicine, Dept. of Pediatrics, 333 Cedar St., LMP 3107, New Haven, CT 06520 (E-mail: ying.xia{at}yale.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
10.1152/ajpcell.00226.2001
Received 18 May 2001; accepted in final form 4 January 2002.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Aitchison, KA,
Baxter GF,
Awan MM,
Smith RM,
Yellon DM,
and
Opie LH.
Opposing effects on infarction of delta and kappa opioid receptor activation in the isolated rat heart: implications for ischemic preconditioning.
Basic Res Cardiol
95:
1-10,
2000[ISI][Medline].
2.
Armstead, WM.
The contribution of 1- and
2-opioid receptors to hypoxia-induced pial artery dilation in the newborn pig.
J Cereb Blood Flow Metab
15:
539-546,
1995[ISI][Medline].
3.
Armstead, WM.
Nitric oxide contributes to opioid release from glia during hypoxia.
Brain Res
813:
398-401,
1998[ISI][Medline].
4.
Bausch, SB,
Patterson TA,
Appleyard SM,
and
Chavkin C.
Immunocytochemical localization of opioid receptors in mouse brain.
J Chem Neuroanat
8:
175-189,
1995[ISI][Medline].
5.
Bofetiado, DM,
Mayfield KP,
and
D'Alecy LG.
Alkaloid agonist BW373U86 increases hypoxic tolerance.
Anesth Analg
82:
1237-1241,
1996[Abstract].
6.
Chang, A,
Takemori AE,
Ojala WH,
Gleason WB,
and
Portoghese PS.
Opioid receptor selective affinity labels: electrophilic benzeneacetamides as
-selective opioid antagonists.
J Med Chem
37:
4490-4498,
1994[ISI][Medline].
7.
Choi, DW.
Ionic dependence of glutamate neurotoxicity in cortical cell culture.
J Neurosci
7:
369-379,
1987[Abstract].
8.
Choi, DW,
and
Rothman SM.
The role of glutamate neurotoxicity in hypoxic-ischemic neuronal death.
Annu Rev Neurosci
13:
171-182,
1990[ISI][Medline].
9.
Chow, E,
and
Haddad GG.
Differential effects of anoxia and glutamate on cultured neocortical neurons.
Exp Neurol
150:
52-59,
1998[ISI][Medline].
10.
Commons, KG,
and
Milner TA.
Cellular and subcellular localization of opioid receptor immunoreactivity in the rat dentate gyrus.
Brain Res
738:
181-195,
1996[ISI][Medline].
11.
Connor, M,
and
Christie MJ.
Opioid receptor signalling mechanisms.
Clin Exp Pharmacol Physiol
26:
493-499,
1999[ISI][Medline].
12.
Connor, M,
Schuller A,
Pintar JE,
and
Christie MJ.
µ-Opioid receptor modulation of calcium channel current in periaqueductal grey neurons from C57B16/J mice and mutant mice lacking MOR-1.
Br J Pharmacol
126:
1553-1558,
1999
13.
Di Loreto, S,
and
Balestrino M.
Development of vulnerability to hypoxic damage in in vitro hippocampal neurons.
Int J Dev Neurosci
15:
225-230,
1997[ISI][Medline].
14.
Fang, L,
Knapp RJ,
Horvath R,
Matsunaga TO,
Haaseth RC,
Hruby VJ,
Porreca F,
and
Yamamura HI.
Characterization of [3H]naltrindole binding to delta opioid receptors in mouse brain and mouse vas deferens: evidence for delta opioid receptor heterogeneity.
J Pharmacol Exp Ther
268:
836-846,
1994[Abstract].
15.
Foutz, AS,
Dauthier C,
and
Kerdelhue B.
-Endorphin plasma levels during neuromuscular blockade in unanesthetized cat.
Brain Res
263:
119-123,
1983[ISI][Medline].
16.
Goldberg, MP,
Monyer H,
and
Choi DW.
Hypoxic neuronal injury in vitro depends on extracellular glutamine.
Neurosci Lett
94:
52-57,
1988[ISI][Medline].
17.
Grafe, MR.
Developmental changes in the sensitivity of the neonatal rat brain to hypoxic/ischemic injury.
Brain Res
653:
161-166,
1994[ISI][Medline].
18.
Haddad, GG,
and
Jiang C.
O2 deprivation in the central nervous system: on mechanisms of neuronal response, differential sensitivity and injury.
Prog Neurobiol
40:
277-318,
1993[ISI][Medline].
19.
Hirashima, Y,
Kurimoto M,
Nogami K,
Endo S,
Saitoh M,
Ohtani O,
Nagata T,
Muraguchi A,
and
Takaku A.
Correlation on glutamate-induced apoptosis with caspase activities in cultured rat cerebral cortical neurons.
Brain Res
849:
109-118,
1999[ISI][Medline].
20.
Law, PY,
Wong YH,
and
Loh HH.
Molecular mechanisms and regulation of opioid receptor signaling.
Annu Rev Pharmacol Toxicol
40:
389-430,
2000[ISI][Medline].
21.
Lee, JW,
Joshi S,
Chan JS,
and
Wong YH.
Differential coupling of µ-, -, and
-opioid receptors to G
16-mediated stimulation of phospholipase C.
J Neurochem
70:
2203-2211,
1998[ISI][Medline].
22.
Lobner, D.
Comparison of the LDH and MTT assays for quantifying cell death: validity for neuronal apoptosis?
J Neurosci Methods
96:
147-152,
2000[ISI][Medline].
23.
Mayfield, KP,
and
D'Alecy LG.
Role of endogenous opioid peptides in the acute adaptation to hypoxia.
Brain Res
582:
226-231,
1992[ISI][Medline].
24.
Mayfield, KP,
and
D'Alecy LG.
Delta-1 opioid agonist acutely increases hypoxic tolerance.
J Pharmacol Exp Ther
268:
683-688,
1994[Abstract].
25.
Nyakas, C,
Buwalda B,
and
Luiten PGM
Hypoxia and brain development.
Prog Neurobiol
49:
1-51,
1996[ISI][Medline].
26.
Ostermeier, AM,
Schlosser B,
Schwender D,
and
Sutor B.
Activation of µ- and -opioid receptors causes presynaptic inhibition of glutamatergic excitation in neocortical neurons.
Anesthesiology
93:
1053-1063,
2000[ISI][Medline].
27.
Piros, ET,
Prather PL,
Law PY,
Evan CJ,
and
Hales TG.
Voltage-dependent inhibition of L-type Ca2+ channels by cloned µ- and -opioid receptors.
Mol Pharmacol
50:
947-956,
1996[Abstract].
28.
Portoghese, PS,
Sultana M,
and
Takemori AE.
Naltrindole, a highly selective and potent non-peptide opioid receptor antagonist.
Eur J Pharmacol
146:
185-186,
1988[ISI][Medline].
29.
Remmers, AE,
Clark MJ,
Liu XY,
and
Medzihradsky F.
Delta opioid receptor down-regulation is independent of functional G protein yet is dependent on agonist efficacy.
J Pharmacol Exp Ther
287:
625-632,
1998
30.
Rhim, H,
and
Miller RJ.
Opioid receptors modulate diverse types of calcium channels in the nucleus tractus solitarius of the rat.
J Neurosci
14:
7608-7615,
1994[Abstract].
31.
Riva, MA,
Tascedda F,
Molteni R,
and
Racagni G.
Regulation of NMDA receptor subunit mRNA expression in the rat brain during postnatal development.
Mol Brain Res
25:
209-216,
1994[Medline].
32.
Schultz, JJ,
Hsu AK,
and
Gross GJ.
Ischemic preconditioning and morphine-induced cardioprotection involve the delta ()-opioid receptor in the intact rat heart.
J Mol Cell Cardiol
29:
2187-2195,
1997[ISI][Medline].
33.
Sharif, NA,
Durie E,
Michel AD,
and
Whiting RL.
Dog cerebral cortex contains µ-, - and
-opioid receptors at different densities: apparent lack of evidence for subtypes of the
-receptor using selective radioligands.
Brain Res
510:
108-114,
1990[ISI][Medline].
34.
Stys, PK,
Ransom BR,
Waxman SG,
and
Davis PK.
Role of extracellular calcium in anoxic injury of mammalian central white matter.
Proc Natl Acad Sci USA
87:
4212-4216,
1990[Abstract].
35.
Sutor, B,
and
Zieglgansberger W.
Actions of D-Ala2-D-Leu5-enkephalin and dynorphin A (1-17) on neocortical neurons in vitro.
Neuropeptides
5:
241-244,
1984[ISI][Medline].
36.
Svingos, AL,
Cheng PY,
Clarke CL,
and
Pickel VM.
Ultrastructural localization of -opioid receptor and Met5-enkephalin immunoreactivity in rat insular cortex.
Brain Res
700:
25-39,
1995[ISI][Medline].
37.
Takemori, AE,
Ho BY,
Naeseth JS,
and
Portoghese PS.
Nor-binaltorphimine, a highly selective kappa-opioid antagonist in analgesic and receptor binding assays.
J Pharmacol Exp Ther
246:
255-258,
1988[Abstract].
38.
Takemori, AE,
Larson DL,
and
Portoghese PS.
The irreversible narcotic antagonistic and reversible agonistic properties of the fumaramate methyl ester derivative of naltrexone.
Eur J Pharmacol
70:
445-451,
1981[ISI][Medline].
39.
Tanaka, E,
and
North RA.
Opioid actions on rat anterior cingulate cortex neurons in vitro.
J Neurosci
14:
1106-1113,
1994[Abstract].
40.
Taussig, R,
Sanchez S,
Rifo M,
Gilman AG,
and
Belardetti F.
Inhibition of the -conotoxin-sensitive calcium current by distinct G proteins.
Neuron
8:
799-809,
1992[ISI][Medline].
41.
Tempel, A,
and
Zukin RS.
Neuroanatomical patterns of the µ, , and
opioid receptors of rat brain as determined by quantitative in vitro autoradiography.
Proc Natl Acad Sci USA
84:
4308-4312,
1987[Abstract].
42.
Vogt, BA,
Wiley RG,
and
Jensen EL.
Localization of mu and delta opioid receptors to anterior cingulate afferents and projection neurons and input/output model of mu regulation.
Exp Neurol
135:
83-92,
1995[ISI][Medline].
43.
Ward, SJ,
Fries DS,
Larson DL,
Portoghese PS,
and
Takemori AE.
Opioid receptor binding characteristics of the non-equilibrium µ antagonist, -funaltrexamine (
-FNA).
Eur J Pharmacol
107:
323-330,
1985[ISI][Medline].
44.
Xia, Y,
and
Haddad GG.
Ontogeny and distribution of opioid receptors in the rat brainstem.
Brain Res
549:
181-193,
1991[ISI][Medline].
45.
Xia, Y,
and
Haddad GG.
Major difference in the expression of µ- and -opioid receptors between turtle and rat brain.
J Comp Neurol
436:
202-210,
2001[ISI][Medline].
46.
Yanagida, H,
and
Corssen G.
Respiratory distress and beta-endorphin-like immunoreactivity in humans.
Anesthesiology
55:
515-519,
1981[ISI][Medline].
47.
Young, RS,
During MJ,
Donnelly DF,
Aquila WJ,
Perry VL,
and
Haddad GG.
Effect of anoxia on excitatory amino acids in brain slices of rats and turtles: in vitro microdialysis.
Am J Physiol Regulatory Integrative Comp Physiol
264:
R716-R719,
1993
48.
Zadina, JE,
Kastin AJ,
Harrison LM,
Ge LJ,
and
Chang SL.
Opiate receptor changes after chronic exposure to agonists and antagonists.
Ann NY Acad Sci
757:
353-361,
1995[ISI][Medline].
49.
Zhang, JH,
Haddad GG,
and
Xia Y.
-Opioid receptor expression is differentially regulated in cortical neurons depending on hypoxic duration (Abstract).
Soc Neurosci Abstr
26:
772,
2000.
50.
Zhang, JH,
Haddad GG,
and
Xia Y.
-, But not µ- and
-, opioid receptor activation protects neocortical neurons from glutamate-induced excitotoxic injury.
Brain Res
885:
143-153,
2000[ISI][Medline].