The impact of anesthetic drugs on the developing central nervous system (CNS) has been a topic of rigorous investigation. Preclinical reports have unequivocally established that both intravenous and volatile anesthetic agents induce neuroapoptosis.1 These concerns have also fueled retrospective reports demonstrating an association between exposure to surgery and anesthesia at infancy with subsequent learning disabilities,2–4 which in turn compelled the Food and Drug Administration and the SmartTots collaborative to heighten public awareness on the neurological impact of anesthetics in infants and children and advocate for additional preclinical and clinical research.5 However, the recent interim report from the General Anaesthesia compared to Spinal anaesthesia (GAS) collaboration noted equivalence between spinal and general anesthesia in infants undergoing a brief surgical procedure.6 Given that these are preliminary findings in a very specific group of surgical patients, a comprehensive evaluation of other anesthetic and sedative drugs needs to be continued both on the bench and bedside. Because avoidance of general anesthesia and sedation is not a viable option, the design of a nontoxic anesthetic and sedative drug regimen is an inevitable task.
Dexmedetomidine (DEX) is a selective α2-adrenergic agonist with sympatholytic, sedative, amnestic, and analgesic properties. When administered as an adjuvant to volatile anesthetics, DEX reduces minimum alveolar concentration7 and has been shown to decrease isoflurane-induced and ketamine (KET)-induced neurotoxicity in neonatal rats.8–11 DEX is also neuroprotective in preclinical models of stroke and traumatic brain injury.12,13 The intrinsic neuroprotective properties of DEX have been attributed to increased expression of the prosurvival kinases, phosphorylated extracellular signal-regulated protein kinase 1 and 2 (pERK1/2), and protein kinase B (AKT)-glycogen synthase kinase-3β (GSK-3β).13–15
These reports demonstrate the protective effect of DEX in injury models and concurrent exposure to anesthetic drugs. However, the impact of increasing doses of DEX on naive rat pups and neuronal cell cultures is not known. Human neutrophils incubated in increasing concentrations of DEX underwent apoptosis and loss of mitochondrial transmembrane potentials.16 It is unclear whether these inconsistencies stem from the type of target cells or cumulative dose of DEX used in these studies. One possible explanation is that high-dose DEX results in concentration-dependent activation of the α1-adrenergic receptor, which may attenuate its α2-adrenergic effect.17 Given the use of DEX as a sedative and an adjuvant during general anesthesia, determining its threshold effect on neuronal viability is critical. In the present study, we tested the hypothesis that the neuroprotective profile of DEX has a ceiling effect and high cumulative doses will induce neuroapoptosis and neuronal cell death by modulating phosphorylation of the prosurvival kinases, ERK1/2 and AKT-GSK-3β. Using the same experimental paradigm that we used to demonstrate KET-induced neuroapoptosis,18–20 we evaluated the impact of increasing the cumulative dose and concentration of DEX on neonatal rat pups and primary neuronal cell cultures, respectively.
Animals and Reagents
The following experimental protocols were approved by the Boston Children’s Hospital Investigational Review Board and adhered to the Guide for the Care and Use of Laboratory Animals.21 Pathogen-free Sprague-Dawley (SD) postnatal day 7 (P7) rat pups and embryonic day 18 fetuses (E18) from pregnant SD dams were used for all the experimental procedures (Charles River Laboratories, Wilmington, MA). DEX (Precedex; Hospira, Lake Forest, IL) and KET (Ketalar, Bedford Labs, Bedford, OH) were obtained from commercial sources. The rat pups were kept from their dam and visually monitored for respiratory effort and activity. The treatments were conducted in a temperature-controlled acrylic container maintained at 36.7°C.
Drug Treatment Groups and Determination of Loss of Righting Reflex and Physiological Parameters
Rat pups were randomly assigned to 6 groups by an investigator blinded to the drug treatment. The sex of the pups was not determined because a previous report demonstrated that sex does not have an impact on acute histological assessments of neuronal death.22 However, sex has a significant impact on subsequent neurobehavioral assessments. Each rat pup received 5 intraperitoneal injections of vehicle (saline), DEX (10, 25, or 50 μg/kg/injection), KET (20 mg/kg/injection), or DEX (50 μg/kg/injection) with KET (20 mg/kg/injection) at 90-minute intervals over 6 hours. The corresponding total cumulative doses administered during the 6-hour treatment period were DEX (50, 125, and 250 μg/kg), KET (100 mg/kg), or DEX (250 μg/kg) with KET (100 mg/kg). The DEX dosing schedules were derived from published reports on rodents.8,10,23 A single 25 μg/kg intraperitoneal dose of DEX has been reported to yield plasma concentrations of 5.23 ± 1.22 μg/mL in P7 rat pups.23 The drug dosages and resultant target-site concentrations utilized in this and other published preclinical reports significantly exceed the clinically relevant plasma concentrations obtained in pediatric patients receiving DEX and exemplify divergent species-dependent responses to anesthetic drugs.24,25 The KET dosing regimen is similar to that used in previous investigations using a similar experimental paradigm that yields a KET plasma concentration of 5.80 ± 3.10 μg/mL and a brain concentration of 2.65 ± 1.60 μg/g, which also exceeds clinically relevant plasma concentrations in humans.26 Because a previous report noted that 25 μg/kg of DEX given after KET attenuated the neuroapoptotic response,10 we elected to administer DEX (250 μg/kg) with KET (100 mg/kg) over 6 hours to determine the effect of a higher dose of DEX. The rat pups were kept from their dam and visually monitored for respiratory effort and activity. The effect of the different drug treatments was measured by the loss of righting reflex (LORR), which is the time between the administration of the drug and when the rat pup righted itself. Heart rate and oxygen saturation were also recorded immediately after each injection period with a noninvasive rodent physiologic monitor (MouseOx; Starr Life Sciences Corp., Oakmont, PA) (5 pups per treatment group).
Six hours after the initial treatment, the rat pups were euthanized with intraperitoneal pentobarbital (75 mg/kg). The brains from each group (n = 6 per group) were rapidly isolated and frozen in liquid nitrogen and processed for protein analysis. A second cohort of rat pups (n = 6 per group) was anesthetized with intraperitoneal pentobarbital (75 mg/kg) and immediately perfused with saline followed by 4% paraformaldehyde. The brains were subsequently embedded in paraffin for histological processing.
Immunohistochemical Staining for Cleaved-Caspase-3
Expression of cleaved-caspase-3 was determined in the brain sections by immunohistochemistry.19 Sections of rat brain tissue were deparaffinized and rehydrated with distilled water. Endogenous peroxidases were inactivated by immersing the samples in hydrogen peroxide for 10 minutes and then incubated for 10 minutes with 10% normal goat serum to block nonspecific binding. The sections were incubated at 4°C overnight with a rabbit anti-cleaved-caspase-3 antibody (1:1000; Cell Signaling Technology, Beverly, MA) and incubated with a biotinylated anti-rabbit IgG and peroxidase-conjugated streptavidin (ZYMED Laboratories, Inc, Carlsbad, CA) and followed by a chromogenic reaction by 3,3′-diaminobenzidine.27 All the sections were counterstained with hematoxylin. The same protocol was applied to the control slides with the omission of the primary antibody.
Immunofluorescence Staining for Cleaved-Caspase-3
To determine the cell type that expressed cleaved-caspase-3, a cohort of brain sections was incubated with anti-cleaved-caspase-3 (1:1000; Cell Signaling Technology) and neuron-specific mouse anti-NeuN (1:100; Abcam, Cambridge, MA) antibodies overnight. The brain sections were then incubated with the following secondary antibodies: Cy3-conjugate donkey anti-rabbit (Jackson ImmunoResearch, West Grove, PA) and streptavidin-Alexa Fluor 488 conjugate (Invitrogen, Carlsbad, CA) for 90 minutes. The secondary antisera were diluted 1:100. Both primary and secondary antisera were diluted in phosphate-buffered solution (PBS), 0.3% Triton X-100 (Sigma-Aldrich, St. Louis, MO), 0.04% bovine serum albumin, and 0.1% sodium azide. The processed tissue was rinsed in PBS before mounting on slides. After drying, mounted sections were covered with 90% glycerol. Finally, the slides were imaged with fluorescent microscope (Olympus IX81; Olympus, Tokyo, Japan).
Apoptosis was also determined using the terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL) assay (Millipore; Serologicals Corporation, Norcross, GA). Brain sections were deparaffinized and rehydrated. After treatment with proteinase K (20 μg/mL; Roche Applied Science, Indianapolis, IN) and quenching with 3.0% hydrogen peroxide, the sections were incubated in a terminal deoxynucleotidyl transferase (TdT) reaction mix for 1 hour at 37°C. The sections were washed in stop/wash buffer, then incubated for 30 minutes in a solution of antidigoxigenin conjugate, and colorized with 3,3'-Diaminobenzidine (DAB, Sigma-Aldrich). These sections were counterstained with hematoxylin. The TdT reaction mix was omitted for the control sections.
The number of cleaved-caspase-3 and TUNEL-positive cells was counted by an investigator blinded to the treatment group under ×400 magnification in 7 representative fields in the somatosensory cortex from 6 rat cortices chosen from each group.20 We focused our cell counts in the somatosensory cortex because this region was most vulnerable to the neurotoxic effects of KET in P7 rat pups.18,28
Western Blotting Assay
To measure the effect of the treatment conditions on intrinsic apoptotic cell death (cleaved-caspase-3) and cell survival proteins, total protein was extracted from flash-frozen brain tissue, as described in our previous study.19 Primary antibodies for cleaved-caspase-3, AKT, pAKT (Ser 473), GSK-3β, pGSK-3β (Ser 9), pERK1/2 (Thr202/Tyr204), total ERK1/2, and β-actin (Cell Signaling Technology) were used for incubation with the membrane overnight at 4°C with slow shaking. Horseradish peroxidase–conjugated anti-rabbit secondary antibody was added at room temperature for an additional 2 hours. The protein bands in membranes were visualized by enhanced chemiluminescence (Thermo Fisher Scientific, Rockford, IL). The densities of the protein bands were quantified with ImageJ 1.42 (NIH, Bethesda, MD).
Primary Neuronal Cell Viability
Primary neuronal cells were isolated from cortices of embryonic day 18 SD rat fetuses as previously described.18 The fetuses were removed quickly from the pregnant dams euthanized with CO2. The cortices of brain were dissected in ice-cold Hank’s balance salt solution (HBSS) under a microscope (Leica, Buffalo Grove, IL). The cortices were dissociated with 0.25% trypsin-EDTA in HBSS for 30 minutes. Cortical neurons were isolated and cultured in 96-well plates precoated with poly-D-lysine and Neurobasal medium (Invitrogen) containing 2% B27 supplement, 500 μM glutamine, and 10 μg/mL gentamycin (Invitrogen). After a 3-day incubation period, E18 cells were treated with different concentrations of DEX (0.001, 0.01, 0.05, 0.1, and 0.2 μM) or KET (50 μM) for 6 hours. The concentration of KET was derived from published work on KET and neuronal cell cultures.29 Cell viability was measured by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay (Sigma-Aldrich). To determine whether blockade of the α1-adrenergic receptor mitigated the toxic effect of DEX, E18 cells were treated with DEX 0.2 μM with and without prazosin hydrochloride 0.5 μM or the α2-adrenergic receptor antagonist yohimbine hydrochloride (0.3 μM; Sigma-Aldrich).
In line with our and other laboratories’ practices, a minimum of 6 rat pups for each experimental group was utilized to detect a 40% difference in the mean values with 80% power at a significance level of .05.30,31 Changes in cleaved-caspase-3 levels were presented as percentage of control value. The cell viability was presented as a ratio of control. AKT, GSK-3β, and ERK ratios of the phosphorylated and total forms were calculated. Cleaved-caspase-3, TUNEL, and merged cleaved-caspase-3 and NeuN-positive cells were presented as absolute values per microscopic field. Data were expressed as mean and standard error of the mean (SEM).
Normal distribution of the residuals was assessed by the Shapiro-Wilk test. Cleaved-caspase-3, AKT, GSK-3β, and ERK1/2 protein levels and E18 cell viability assays had a normal distribution, were analyzed with 1-way analysis of variance (ANOVA), and were followed by a post hoc analyses derived by the Tukey and its 95% confidence intervals on focused paired comparisons on the Western blots and the Dunnett test (2-tailed) on the in vitro cell viability assessments.
The cell counts from the brain slices stained for cleaved-caspase-3, TUNEL, double-stained images for cleaved-caspase-3, and NeuN had skewed distributions, and a Kruskal-Wallis test was applied to analyze differences. Post hoc analyses on focused paired comparisons were derived by determining the Wilcoxon-Mann-Whitney Odds (WMWodds) and its 95% confidence intervals. These values were determined by binary logistic regression, with the dependent variable being the treatment group (control, DEX 50, etc) and the independent variable being the cell counts per field followed by receiver operating characteristic (ROC) analysis. WMWodds and its Dunn corrected 95% confidence interval were calculated directly from the ROC model area under the curve and the confidence interval.a
The impact of increasing doses of DEX over time on LORR, heart rate, and oxygen saturation was analyzed using 2-way ANOVA with dose and time as independent variables. Data analyses were generated using IBM SPSS Statistics version 23.0 (IBM, Armonk, NY). Statistical significance was set at P < .05.
Cumulative Dose of DEX Affects the LORR and Physiological Parameters
There was no mortality during the experimental period for all groups. However, increasing the cumulative dose of DEX prolonged the LORR, with the highest dose of DEX producing similar times as the KET and DEX + KET groups (Figure 1A). The LORR persisted throughout the 6-hour treatment period in the high-dose DEX, KET, and DEX + KET groups. Rats from the 3 DEX groups had significantly lower heart rates (Figure 1B). Oxygen saturation was maintained in the DEX groups, but decreased in the KET and DEX + KET groups (Figure 1C). Two-way ANOVA revealed significant main effects of time and drug treatment on LORR, heart rate, and oxygen saturation.
Escalating the Cumulative Dose of DEX Increased Neuroapoptosis
Increasing the cumulative dose of DEX resulted in elevated cleaved-caspase-3 levels. Although DEX 125 and 250 were statistically higher, the difference between DEX 50 and control was not statistically significant. Coadministration of DEX with KET did not significantly attenuate cleaved-caspase-3 expression when compared with the KET group (Figure 2).
Significant differences in apoptotic cell counts from brain sections stained with either cleaved-caspase-3 or TUNEL technique were noted between the treatment groups (Figures 3 and 4). In contrast with the Western blots, the post hoc analysis revealed that the apoptotic cell counts from the cleaved-caspase-3 and TUNEL-stained brain slices of the DEX 50 group were higher than control. The KET group also had higher counts of cleaved-caspase-3 and TUNEL-positive cells than the DEX and DEX + KET groups (Figures 3 and 4). Immunofluorescence microscopy of brain sections stained with antibodies to cleaved-caspase-3 (red) and neuron-specific nuclear protein (NeuN, green cells) confirmed that most of the cleaved-caspase-3-positive cells were neurons (Figure 5).
DEX Maintained Phosphorylation Levels of AKT and GSK-3β
The effect of DEX on the ratio of phosphorylated and total levels of AKT and GSK-3β is shown in Figure 6. As we previously reported, KET reduced the phosphorylation of both kinases.20 When DEX was administered as the sole drug, pAKT at DEX 50 and DEX 125 μg/kg were maintained at the same levels as the control group and decreased at DEX 250 μg/kg. pGSK-3β levels were maintained at control levels and peaked at DEX (125 μg/kg). However, the KET group and the combined DEX and KET group had a significant reduction in both pAKT and pGSK-3β levels.
Low-Dose DEX Enhanced pERK1/2 Expression
The cumulative dose of DEX had a differential effect on pERK1 and pERK2. DEX at cumulative doses of 50 and 125 μg/kg significantly increased the expression of pERK1 and pERK2 (Figure 7). DEX 250 μg/kg or KET 100 mg/kg significantly decreased both the expression of pERK1 and pERK2. Of note, coadministration of DEX and KET resulted in a significant reduction when compared with the effect of either drug alone. This finding implies that the combination of high-dose DEX and KET synergistically reduced pERK1 and pERK2 levels.
DEX Decreased Neuronal Cell Viability in a Concentration-Dependent Manner
Increasing the concentration of DEX reduced E18 neuronal cell viability (Figure 8A). The addition of prazosin 0.5 μM or yohimbine 0.3 μM to E18 cell cultured in DEX 0.02 μM preserved cell viability (Figure 8B). These data suggest that increasing DEX concentrations has a reciprocal effect on neuronal cell viability. Pharmacological blockade of α1-adrenergic receptors with prazosin or yohimbine mitigated this response.
In the present study, increasing the cumulative dose of DEX escalated neuroapoptosis as detected by cleaved-caspase-3 expression and TUNEL staining. High-dose DEX, which equates to a cumulative dose of 250 μg/kg over 6 hours, resulted in a similar neuroapoptotic profile as KET alone, thus demonstrating that DEX administered solely over 6 hours induced neuroapoptosis in a cumulative dose-dependent manner. DEX (50 μg/kg) at least maintained and in some cases enhanced phosphorylation of ERK1/2, AKT, and GSK-3β, which favors neuronal survival. Increasing the cumulative dose of DEX significantly prolonged the LORR, with the highest doses having a similar effect as the KET and DEX + KET groups. DEX decreased heart rate but maintained oxygen saturation. DEX reduced neuronal cell viability in a concentration-dependent manner in vitro, but coincubation with an α1-adrenergic receptor antagonist mitigated this response.
The key differences between our findings and these published reports are that we used higher cumulative dose concentrations of DEX on naive rat pups and E18 cells. Previously published reports mainly examined the impact of DEX on various models of CNS injury. These experimental paradigms include isoflurane-induced neurotoxicity, traumatic brain injury, and stroke.8,12,13 These studies also used doses of DEX that were at the lower range utilized in our study. Specifically, the cumulative doses of DEX used in these published reports range from 75 to 225 μg/kg over 4 hours, 75 to 225 μg/kg over 3 days, and 75 μg/kg over 6 hours.8,10,11 We used doses up to 250 μg/kg over 6 hours. Collectively, we show that neuroapoptosis and cell death can be attributed to the larger cumulative dose of DEX. Our findings support the premise that DEX administered at low doses was neuroprotective, whereas high doses were neurotoxic.
We attribute the neurotoxic effect of DEX to the off-target α1-adrenergic activation17 because pharmacological blockade of the α1-adrenergic receptor by prazosin or yohimbine preserved cell viability in the presence of high concentrations of DEX in vitro. Although yohimbine is touted as being an α2-adrenergic receptor selective antagonist, it has a selectivity value (α2/α1) of 45.32 Therefore, it is likely that the neuroprotective effect of yohimbine is because of off-target antagonism of the α1-adrenergic receptor.
The AKT-GSK-3β signaling pathway is a key cell signaling pathway that plays a major role in neuronal development and cell survival, primarily by regulating neuronal progenitor cell proliferation and neuronal polarity.33 GSK-3β is a unique protein kinase that is inactivated by phosphorylation by AKT. Activation of GSK-3β increases neuroapoptosis and is associated with neurodegenerative disorders.20,34 Prolonged exposure to KET or sevoflurane results in decreased phosphorylation of AKT and increased GSK-3β activity.20,35,36 DEX (75 μg/kg over 4 hours) has been shown to inhibit isoflurane-induced neuroapoptosis in rat pups by increasing in pAKT expression.11 DEX similarly has been shown to increase pAKT and pGSK-3β in rodents undergoing ischemic injury.13,37 In the present study, low-dose DEX slightly increased pGSK-3β, instead of decreasing levels as seen in the KET-treated rat pups. In contrast to the impact of KET and sevoflurane on decreasing phosphorylation of GSK-3β, our data confirm that DEX maintains pGSK-3β levels at all dosing regimens. These findings confirm that the neuroprotective properties of DEX may be because of its ability to attenuate GSK-3β activity.8,10
Extracellular signal-regulated kinases 1 and 2 are protein kinases that play a central role in neuronal transcriptional events that promote neuronal development and synaptic plasticity.38 Several reports attribute the neuroprotective effect of DEX to its ability to enhance phosphorylation of ERK1/2 in a variety of neuronal injury models.9,13,14 KET and propofol have been shown to attenuate this process.36 Our data demonstrate that cumulative doses of 50 and 125 μg/kg increase pERK1/2, which may account for the reported neuroprotective properties.8,10 However, the highest cumulative dose (250 μg/kg) significantly decreased pERK1/2 expression and increased cleaved-caspase-3 expression, thereby exceeding the neuroprotective threshold of DEX.
There are conflicting reports on the impact of DEX on cell viability. DEX induced neutrophil apoptosis in a concentration-dependent fashion after a 24-hour exposure.16 Neutrophils incubated in 10 or 100 ng/mL, which corresponds to 0.05 and 0.5 μM, respectively, had a significantly higher apoptosis index than neutrophils in the control and 0.005 μM groups. Furthermore, neutrophils treated with DEX at 0.5 μM lost mitochondrial transmembrane potentials, which is an indicator of mitochondrial dysfunction that leads to apoptotic cell death. In contrast, DEX at concentrations between 1 and 100 μM prevented staurosporine- or wortmannin-induced cell death in primary neuronal cell cultures.9 DEX (1 μM) also attenuated cell death in the CA1 region of organotypic hippocampal slices deprived of oxygen and glucose.15 This putative protection was attributed to activation of the prosurvival AKT and ERK pathways. In an in vitro model of traumatic brain injury, delayed administration of DEX protected organotypic hippocampal slices.12 This report included a concentration response study that showed maximal protection at DEX 1 μM, while 10 and 100 μM had no effect. Coincubation with an ERK inhibitor mitigated the protective properties of DEX. Another report on isolated spinal cord neurons pretreated with DEX 1 μM before oxygen and glucose deprivation had increased levels of pAKT, which resulted in increased cell viability and decreased apoptosis.37 Together, these in vitro reports demonstrate that DEX boosts cell survival pathways in the setting of experimental models of cell injury and stress. In naive primary neuronal cells, we found that DEX produces concentration-dependent decreases in cell viability, which suggest that DEX has a cytotoxic effect at high concentrations. After a 6-hour incubation period, exposure to DEX at concentrations of 0.05 μM and higher decreased E18 viability. These DEX concentrations are consistent with levels reported to induce neutrophil apoptosis.16 The difference between the published reports on DEX neuroprotection and our observations is that the former examines the impact of DEX in injury models, whereas our experiments examine the effect of DEX on naive neurons. The reported range of plasma levels measured in sedated pediatric patients receiving DEX infusions is 0.4 to 0.8 μg/L, which corresponds to 0.002 to 0.004 μM.24 Therefore, it should be noted that the DEX concentrations utilized in this rodent study is 10-fold higher than the plasma concentrations used to provide sedation in humans. The disparate dosing schedules utilized in preclinical studies and clinical practice in human highlight species-dependent variability in response to anesthetic drugs.25
There are several limitations to this report. The primary aim of this investigation is to determine the acute impact of cumulative doses on neuroapoptosis and cell signaling kinases at an early developmental stage (P7) and not at later developmental stages. Anesthetic exposure at later developmental stages will result in distinct histological and behavioral derangements.39–41 It is also known that exposure to anesthetics at this early critical period of development has a detrimental impact on subsequent neurobehavioral assessments.42 The cell counting performed in our study was based on an acquired image from the microscope rather than the stereological technique. These counts derived from a single plane, which does not correct for cell morphology.
Because published hemodynamic profiling of DEX revealed bradycardia and hypertension in rats,43 a secondary aim of this report is to measure the physiological effects of DEX on P7 rat pups. We demonstrated a significant effect of duration and dose on LORR and heart rate. Given the limitations of our model, it is difficult to determine the impact of these physiological changes on neuroapoptosis and cell signaling pathways. However, the in vitro experiments clearly demonstrate a concentration-dependent decrease in neuronal cell viability, which is consistent with published reports in other cell types.16 Last, we show that DEX increased pERK1/2 and pAKT, which have intrinsic neuroprotective properties. However, our experiments did not investigate the role of the imidazoline 1 receptor-protein kinase C pathway, which modulates the phosphorylation of ERK1/2 and has been implicated in the DEX-mediated pre- and postconditioning neuroprotection.14,15 Future investigations should interrogate the impact of DEX on neurogenesis, differentiation, dendritic development, synaptic maintenance, and neurobehavioral assessments throughout the life span.
In summary, our findings demonstrate that DEX induces neuroapoptosis in P7 rat pups in cumulative dose-dependent manner. The combination DEX and KET did not attenuate neuroapoptosis in comparison with KET alone. Our findings also showed that the total cumulative dose of DEX regulates phosphorylation of ERK1/2 and AKT. Similar to published reports, low cumulative doses enhanced, whereas high cumulative doses decreased pERK1/2 expression. DEX combined with KET reduced phosphorylation of AKT, GSK-3β, and ERK1/2 and increased neuronal apoptosis. The interaction of DEX with KET in the setting of developmental neuroapoptosis is still unclear. Our observations suggest that, although DEX is neuroprotective in low doses, high cumulative doses can induce neuroapoptosis. Increasing concentrations of DEX also diminished direct cytotoxic effect. Because the current dosing schedules used in humans yield plasma levels that are substantially below concentrations that induced neuroapoptosis in vivo and cell death in vitro,24,44 current human dosing schedules for DEX should not be neurotoxic and DEX has the potential to be a neuroprotective adjuvant.
Name: Jia-Ren Liu, MD, PhD.
Contribution: This author helped design the study, perform the experiments and data analysis, and prepare the manuscript.
Name: Koichi Yuki, MD.
Contribution: This author helped design the study and data analysis and prepare the manuscript.
Name: Chongwha Baek, MD.
Contribution: This author helped perform the experiments and data analysis.
Name: Xiao-Hui Han, RN.
Contribution: This author helped perform the experiments.
Name: Sulpicio G. Soriano, MD.
Contribution: This author helped design the study, perform the experiments and data analysis, and prepare the manuscript.
This manuscript was handled by: Gregory Crosby, MD.
a Dexter F. Wilcoxon-Mann-Whitney test used for data that are not normally distributed. Anesth Analg. 2013;117:537–538.; and Devine G, Norton HJ, Hunt R, Dienemann J. Statistical grand rounds: a review of analysis and sample size calculation considerations for Wilcoxon tests. Anesth Analg. 2013;117:699–710.
1. Stratmann G. Review article: neurotoxicity of anesthetic drugs in the developing brain. Anesth Analg. 2011;113:1170–1179.
2. Wilder RT, Flick RP, Sprung J, et al. Early exposure to anesthesia and learning disabilities in a population-based birth cohort. Anesthesiology. 2009;110:796–804.
3. Ing C, DiMaggio C, Whitehouse A, et al. Long-term differences in language and cognitive function after childhood exposure to anesthesia. Pediatrics. 2012;130:e476–e485.
4. Block RI, Thomas JJ, Bayman EO, Choi JY, Kimble KK, Todd MM. Are anesthesia and surgery during infancy associated with altered academic performance during childhood? Anesthesiology. 2012;117:494–503.
5. Rappaport BA, Suresh S, Hertz S, Evers AS, Orser BA. Anesthetic neurotoxicity—clinical implications of animal models. N Engl J Med. 2015;372:796–797.
6. Davidson AJ, Disma N, de Graaff JC, et al; GAS Consortium. Neurodevelopmental outcome at 2 years of age after general anaesthesia and awake-regional anaesthesia in infancy (GAS): an international multicentre, randomised controlled trial. Lancet. 2016;387:239–250.
7. Segal IS, Vickery RG, Walton JK, Doze VA, Maze M. Dexmedetomidine diminishes halothane anesthetic requirements in rats through a postsynaptic alpha 2 adrenergic receptor. Anesthesiology. 1988;69:818–823.
8. Sanders RD, Xu J, Shu Y, et al. Dexmedetomidine attenuates isoflurane-induced neurocognitive impairment in neonatal rats. Anesthesiology. 2009;110:1077–1085.
9. Sanders RD, Sun P, Patel S, Li M, Maze M, Ma D. Dexmedetomidine provides cortical neuroprotection: impact on anaesthetic-induced neuroapoptosis in the rat developing brain. Acta Anaesthesiol Scand. 2010;54:710–716.
10. Duan X, Li Y, Zhou C, Huang L, Dong Z. Dexmedetomidine provides neuroprotection: impact on ketamine-induced neuroapoptosis in the developing rat brain. Acta Anaesthesiol Scand. 2014;58:1121–1126.
11. Li Y, Zeng M, Chen W, et al. Dexmedetomidine reduces isoflurane-induced neuroapoptosis partly by preserving PI3K/Akt pathway in the hippocampus of neonatal rats. PLoS One. 2014;9:e93639.
12. Schoeler M, Loetscher PD, Rossaint R, et al. Dexmedetomidine is neuroprotective in an in vitro
model for traumatic brain injury. BMC Neurol. 2012;12:20.
13. Zhu YM, Wang CC, Chen L, et al. Both PI3K/Akt and ERK1/2 pathways participate in the protection by dexmedetomidine against transient focal cerebral ischemia/reperfusion injury in rats. Brain Res. 2013;1494:1–8.
14. Dahmani S, Paris A, Jannier V, et al. Dexmedetomidine increases hippocampal phosphorylated extracellular signal-regulated protein kinase 1 and 2 content by an alpha 2-adrenoceptor-independent mechanism: evidence for the involvement of imidazoline I1 receptors. Anesthesiology. 2008;108:457–466.
15. Dahmani S, Rouelle D, Gressens P, Mantz J. Characterization of the postconditioning effect of dexmedetomidine in mouse organotypic hippocampal slice cultures exposed to oxygen and glucose deprivation. Anesthesiology. 2010;112:373–383.
16. Kishikawa H, Kobayashi K, Takemori K, Okabe T, Ito K, Sakamoto A. The effects of dexmedetomidine on human neutrophil apoptosis. Biomed Res. 2008;29:189–194.
17. Schwinn DA, Correa-Sales C, Page SO, Maze M. Functional effects of activation of alpha-1 adrenoceptors by dexmedetomidine: in vivo
and in vitro
studies. J Pharmacol Exp Ther. 1991;259:1147–1152.
18. Soriano SG, Liu Q, Li J, et al. Ketamine activates cell cycle signaling and apoptosis in the neonatal rat brain. Anesthesiology. 2010;112:1155–1163.
19. Liu JR, Liu Q, Li J, et al. Noxious stimulation attenuates ketamine-induced neuroapoptosis in the developing rat brain. Anesthesiology. 2012;117:64–71.
20. Liu JR, Baek C, Han XH, Shoureshi P, Soriano SG. Role of glycogen synthase kinase-3β in ketamine-induced developmental neuroapoptosis in rats. Br J Anaesth. 2013;110(suppl 1):i3–i9.
21. Institute of Laboratory Animal Research Commission on Life Sciences NRC: Guide for the Care and Use of Laboratory Animals
. 1996Washington, DC: The National Academy Press.
22. Lee BH, Chan JT, Kraeva E, Peterson K, Sall JW. Isoflurane exposure in newborn rats induces long-term cognitive dysfunction in males but not females. Neuropharmacology. 2014;83:9–17.
23. McAdams RM, McPherson RJ, Kapur R, Phillips B, Shen DD, Juul SE. Dexmedetomidine reduces cranial temperature in hypothermic neonatal rats. Pediatr Res. 2015;77:772–778.
24. Potts AL, Anderson BJ, Warman GR, Lerman J, Diaz SM, Vilo S. Dexmedetomidine pharmacokinetics in pediatric intensive care—a pooled analysis. Paediatr Anaesth. 2009;19:1119–1129.
25. Berde C, Cairns B. Developmental pharmacology across species: promise and problems. Anesth Analg. 2000;91:1–5.
26. Zou X, Patterson TA, Sadovova N, et al. Potential neurotoxicity of ketamine in the developing rat brain. Toxicol Sci. 2009;108:149–158.
27. Slack GW, Wizniak J, Dabbagh L, Shi X, Gelebart P, Lai R. Flow cytometric detection of ZAP-70 in chronic lymphocytic leukemia: correlation with immunocytochemistry and Western blot analysis. Arch Pathol Lab Med. 2007;131:50–56.
28. Ikonomidou C, Bosch F, Miksa M, et al. Blockade of NMDA receptors and apoptotic neurodegeneration in the developing brain. Science. 1999;283:70–74.
29. Wang C, Sadovova N, Hotchkiss C, et al. Blockade of N-methyl-D-aspartate receptors by ketamine produces loss of postnatal day 3 monkey frontal cortical neurons in culture. Toxicol Sci. 2006;91:192–201.
30. Dell RB, Holleran S, Ramakrishnan R. Sample size determination. ILAR J. 2002;43:207–213.
31. Stratmann G, Sall JW, May LD, et al. Isoflurane differentially affects neurogenesis and long-term neurocognitive function in 60-day-old and 7-day-old rats. Anesthesiology. 2009;110:834–848.
32. Doxey JC, Lane AC, Roach AG, Virdee NK. Comparison of the alpha-adrenoceptor antagonist profiles of idazoxan (RX 781094), yohimbine, rauwolscine and corynanthine. Naunyn Schmiedebergs Arch Pharmacol. 1984;325:136–144.
33. Hur EM, Zhou FQ. GSK3 signalling in neural development. Nat Rev Neurosci. 2010;11:539–551.
34. King TD, Bijur GN, Jope RS. Caspase-3 activation induced by inhibition of mitochondrial complex I is facilitated by glycogen synthase kinase-3beta and attenuated by lithium. Brain Res. 2001;919:106–114.
35. Tao G, Zhang J, Zhang L, et al. Sevoflurane induces tau phosphorylation and glycogen synthase kinase 3β activation in young mice. Anesthesiology. 2014;121:510–527.
36. Straiko MM, Young C, Cattano D, et al. Lithium protects against anesthesia-induced developmental neuroapoptosis. Anesthesiology. 2009;110:862–868.
37. Freeman KA, Puskas F, Bell MT, et al. Alpha-2 agonist attenuates ischemic injury in spinal cord neurons. J Surg Res. 2015;195:21–28.
38. Thomas GM, Huganir RL. MAPK cascade signalling and synaptic plasticity. Nat Rev Neurosci. 2004;5:173–183.
39. Vutskits L, Gascon E, Tassonyi E, Kiss JZ. Effect of ketamine on dendritic arbor development and survival of immature GABAergic neurons in vitro
. Toxicol Sci. 2006;91:540–549.
40. De Roo M, Klauser P, Briner A, et al. Anesthetics rapidly promote synaptogenesis during a critical period of brain development. PLoS One. 2009;4:e7043.
41. Qiu L, Zhu C, Bodogan T, et al. Acute and long-term effects of brief sevoflurane anesthesia during the early postnatal period in rats. Toxicol Sci. 2016;149:121–133.
42. Vutskits L. General anesthesia: a gateway to modulate synapse formation and neural plasticity? Anesth Analg. 2012;115:1174–1182.
43. Bol CJ, Vogelaar JP, Mandema JW. Anesthetic profile of dexmedetomidine identified by stimulus-response and continuous measurements in rats. J Pharmacol Exp Ther. 1999;291:153–160.
© 2016 International Anesthesia Research Society
44. Fujita Y, Inoue K, Sakamoto T, et al. A comparison between dosages and plasma concentrations of dexmedetomidine in clinically ill patients: a prospective, observational, cohort study in Japan. J Intensive Care. 2013;1:15.