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The mechanisms leading to delayed neuronal death after asphyxial cardiac arrest (ACA) in the developing brain are unknown. This study aimed at investigating the possible role of microglial activation in neuronal death in developing brain after ACA. Postnatal day-17 rats were subjected to 9mins of ACA followed by resuscitation. Rats were randomized to treatment with minocycline, (90mg/kg, intraperitoneally (i.p.)) or vehicle (saline, i.p.) at 1h after return of spontaneous circulation. Thereafter, minocycline (22.5mg/kg, i.p.) was administrated every 12h until sacrifice. Microglial activation (evaluated by immunohistochemistry using ionized calcium-binding adapter molecule-1 (Iba1) antibody) coincided with DNA fragmentation and neurodegeneration in CA1 hippocampus and cortex (assessed by deoxynucleotidyltransferase-mediated dUTP nick-end labeling (TUNEL), Fluoro-Jade-B and Nissl stain). Minocycline significantly decreased both the microglial response and neuronal degeneration compared with the vehicle. Asphyxial CA significantly enhanced proinflammatory cytokine and chemokine levels in hippocampus versus control (assessed by multiplex bead array assay), specifically tumor necrosis factor-α (TNF-α), macrophage inflammatory protein-1α (MIP-1α), regulated upon activation, normal T-cell expressed and secreted (RANTES), and growth-related oncogene (GRO-KC) (P<0.05). Minocycline attenuated ACA-induced increases in MIP-1α and RANTES (P<0.05). These data show that microglial activation and cytokine production are increased in immature brain after ACA. The beneficial effect of minocycline suggests an important role for microglia in selective neuronal death after pediatric ACA, and a possible therapeutic target.
Experimental models, which mimic the pathophysiology of cerebral ischemia resulting from cardiac arrest (CA) in adults (Liu et al, 2001; Stoll et al, 1998; Walski and Borowicz, 1995; Walski and Gajkowska, 1998) and hypoxia–ischemia in neonates (Carty et al, 2008; Chew et al, 2006; McRae et al, 1995; Rothstein and Levison, 2002; Sheldon et al, 1996), have been widely used to evaluate the role of inflammation and neuronal death. Compared with adults and neonates, there has been no study evaluating inflammatory response after ischemia in the juvenile brain resulting from asphyxial CA (ACA). While a cardiac etiology is the principal cause of cardiopulmonary arrest in adults, asphyxia is the principal cause of cardiopulmonary arrest in infants and children, resulting in systemic hypoxemia, hypercapnia, acidosis, and ultimately hypotension and bradycardia progressing to complete cardiovascular collapse (Young and Seidel, 1999). Some insight into the developmental aspects of the inflammatory response to hypoxic–ischemic encephalopathy can be extrapolated from the model of carotid artery ligation followed by hypoxemia in immature rodents (Rice et al, 1981); however, this model does not mimic global and systemic hypoxemia/ischemia seen in the ACA victims. Therefore, we sought to examine the possible role of inflammation because it relates to microglial response and cytokine production in neuronal death in the developing brain after ACA, a significant cause of morbidity and mortality in children (Berg et al, 2008). At present there are no interventions to reverse the cellular consequences of ACA and treatment remains in the form of supportive intensive care.
Microglia constitute approximately 10% of the overall number of cells in the brain during most of development and mediate the intrinsic inflammatory response to injury (Chew et al, 2006; Danton and Dietrich, 2003; Zhang et al, 2006). Studies of rodents suggest that microglial activation begins early and can last for days after global cerebral ischemia (Jorgensen et al, 1993; Liu et al, 2001; Morioka et al, 1991; Walski and Borowicz, 1995; Walski and Gajkowska, 1998). Sub-lethal ischemia results in microglial proliferation and activation in the cortex and striatum (Liu et al, 2001). During lethal global ischemia, however, microglial activation occurs throughout the brain, with accumulation in the vulnerable CA1 hippocampus (De Simone et al, 2003). Studies in experimental models have suggested both deleterious and protective effects of the microglial activation after cerebral ischemia. Microglia are capable of producing potentially toxic molecules such as reactive oxygen species, arachidonic acid derivatives, and cytokines (Danton and Dietrich, 2003). Conversely, microglia produce neurotrophins as well as transforming growth factor-β1, which are involved in tissue repair (Danton and Dietrich, 2003; Yrjanheikki et al, 1998). Several lines of evidence suggest that activated microglia are detrimental in cerebral ischemia. Administration of minocycline and other tetracycline antibiotics, which attenuate microglial activation and proinflammatory cytokine production, reduces neuronal death in focal and global cerebral ischemia models (Jantzie et al, 2005; Wang et al, 2002; Yrjanheikki et al, 1998). In contrast, other studies have shown a neuroprotective role for microglia. Selective ablation of proliferating microglial cells exacerbated injury after focal cerebral ischemia (Lalancette-Hebert et al, 2007), and intra-arterial injection of microglia protected hippocampal CA1 neurons against global ischemia in rodents (Hayashi et al, 2006; Imai et al, 2007).
We have developed a model of pediatric ACA and resuscitation in postnatal day (PND) 17 rats that produces transient coma, delayed neuronal death, and functional outcome deficits that are proportional to the duration of asphyxia (Fink et al, 2004, 2005). We used this model to examine the possible role of inflammation as it relates to the microglial response and cytokine production after ACA. We found that ACA induced significant microglial response and increases in cytokine levels in the PND17 brain. Administration of minocycline attenuated the microglial response and increases in cytokine levels, and improved neuronal survival. This suggests an important role for inflammation in selective neuronal death after pediatric ACA, and a possible therapeutic avenue.
Studies were approved by the Institutional Animal Care and Use Committee at the University of Pittsburgh. An established pediatric ACA model (Fink et al, 2004) was used with minor modifications. Briefly, male PND16 to 18 Sprague–Dawley rats (30 to 45g) were anesthetized with 3% isofluorane/50% N2O/balance O2 in a plexiglass chamber until unconscious, then trachea was intubated with an 18-gauge catheter and rats were placed on mechanical ventilation with 1% isofluorane/50%N2O/O2 for surgery. Femoral arterial and venous catheters were inserted via cutdown. All procedures were performed using aseptic technique. Rectal temperature was continuously monitored and maintained at 37°C via a heated water blanket during the surgery. Heart rate and mean arterial blood pressure were monitored continuously. Vecuronium (1mg/kg/h, intravenously) was used for immobilization and was administered 10mins before asphyxia. Isofluorane/N2O was discontinued 2mins before asphyxia. All rats were then exposed to 100% O2 for 1mins for anesthetic wash out and then 21% O2 for 1mins before CA, respectively. After this the ventilator was turned off for 9mins and then restarted with FiO2=1.0 to mimic clinical resuscitative efforts. Epinephrine 0.005mg/kg and sodium bicarbonate 1mEq/kg were administered intravenously, followed by rapid manual chest compressions until return of spontaneous circulation (ROSC). Rats then received 20mL/kg 5% dextrose in water with 0.45mEq/L normal saline, i.p., to prevent dehydration, and were removed from the ventilator and extubated. Rats were observed in FiO2=1.0 for 1h to mimic clinical condition, and then returned to their mother and littermates.
Rats were randomized into one of three groups: naïve, ACA plus vehicle, or ACA plus minocycline (n=3 per group). For the minocycline group, rats were treated with 90mg/kg minocycline (Sigma, St Louis, MO, USA) at 60mins after ROSC on the day of surgery, followed by 22.5mg/kg minocycline every 12h beginning the day after surgery until sacrificed. Rats in the ACA plus vehicle group received similar treatment, except that minocycline was replaced with an equal volume of normal saline.
At 72h after asphyxia, rats were anesthetized and transcardially perfused with 250mL ice-cold heparinized saline followed by 50mL 4% paraformaldehyde (in phosphate-buffered saline, pH 7.4). Brains were removed and further post-fixed in 4% paraformaldehyde for 72h. Paraffin-embedded brains were cut into 5-μm coronal sections. Gross ischemic cellular changes were evaluated in cresyl violet-stained sections, with three sections (20μm apart) for each animal. Viable neurons within the dorsal CA1 region of the hippocampus and cerebral cortex were counted and reported as neuronal density (neurons per mm2).
Fluoro-Jade-B staining of brain sections was used to identify degenerating neurons. Briefly, 5-μm coronal brain sections were deparaffinized in a series of xylene, immersed twice in 100% ethanol (EtOH) and 1% sodium hydroxide (in 80% EtOH) for 90secs, and in then 70% EtOH for 30secs. Slides were then placed on a shaker in 0.06% potassium permanganate for 10mins and washed in distilled water before immersion in a 0.006% working solution of Fluoro-Jade-B (Histo-Chem Inc., Jefferson, AR, USA) with 4′,6-diamidino-2-phenylindole (Sigma, St Louis, MO, USA) for 30mins. Fluoro-Jade-B positive neurons within the dorsal CA1 region of the hippocampus and cerebral cortex were counted.
DNA damage was assessed using terminal deoxynucleotidyltransferase-mediated dUTP nick-end labeling (TUNEL) on 5-μm paraffin sections with a commercially available kit (ApopTag; Chemicon, Temecula, CA, USA).
Brain sections were deparaffinized and then rehydrated in automation buffer (Biocare Medical, Concord, CA, USA). Antigen retrieval was conducted in a microwave oven for 5mins using a commercial antigen retrieval buffer (Biocare Medical, Concord, CA, USA). In some cases, tissue sections were treated with 0.1% Triton X-100 for 5mins to permeabilize cell membranes. After rinsing with automation buffer, sections were incubated with goat serum (Vector, Burlingame, CA, USA) for 1h at room temperature. This was followed by overnight incubation with primary antibodies, antiNeuN (1:200 dilution; Invitrogen, Carlsbad, CA, USA) and antiIba1 (ionized calcium-binding adapter molecule-1) antibody (1:200 dilution; Wako, Osaka, Japan). A combination (1:1) of Alexa Fluor secondary antibodies (594 goat antirabbit, 488 goat antimouse, both with 1:250 dilution; Invitrogen, Carlsbad, CA, USA) were applied to the slides the next day. To assess nonspecific staining, primary antibody-deleted controls were performed at the same time. Sections were examined with a Nikon (EclipSE-E600) microscope.
Cytokine levels were quantified using a Luminex kit (Luminex, Austin, TX, USA), which included 24 cytokines (macrophage inflammatory protein-1α (MIP-1α); regulated upon activation, normal T-cell expressed and secreted (RANTES); and growth-related oncogene (GRO-KC), tumor necrosis factor-α (TNF-α), Eotaxin, granulocyte macrophage colony-stimulating factor, granulocyte colony-stimulating factor, monocyte chemotactic protein-1, leptin, vascular endothelial growth factor (VEGF), interleukin (IL)-1α, IL-1β, IL-2, IL-4, IL-5, IL-6, IL-9, IL-10, IL-12, IL-13, IL-17, IL-18, interferon (IFN)-inducible protein-10, and IFN-γ) following the manufacturer's instructions. Briefly, at 24h after asphyxia, rats were anesthetized and then transcardially perfused with 250mL ice-cold heparinized saline. Brains were removed and hippocampi were dissected, then snap frozen in liquid nitrogen. The tissue was homogenized in saline by using Dounce homogenizer for 20 strokes. The homogenate was then sonicated for 10secs for three times with an interval of 20secs, followed by centrifugation at 40,000g for 25mins. The supernatant was used for cytokine analysis. Protein levels in the supernatants were measured using the BCA protein kit (Thermo Fisher Scientific Inc., Rockford, IL, USA).
Statistical analysis was performed using analysis of variance (ANOVA) and the Tukey's test for post hoc comparison between groups. P<0.05 was considered statistically significant.
Body weight on PND17 was similar among the groups, 40.9±3.5g, 39.5±2.3g, and 40.5±2.5g, respectively, for naïve, ACA plus vehicle, and ACA plus minocycline groups. mean arterial blood pressure (mmHg), PaCO2, PaO2, and pH at baseline and after ROSC are shown in Table 1. There were no statistically significant differences in physiological variables between treatment groups.
To evaluate neuronal loss after ACA, we used crystal violet and Fluoro-Jade staining, and TUNEL. ACA resulted in decrease in cresyl violet staining, indicative of neuronal loss in hippocampus, particularly in CA1 of hippocampus at 72h after the insult (Figure 1). Accompanying neuronal loss, an increase in smaller darkly stained nuclei appeared in CA1 hippocampus at 72h after ACA. Minocycline treatment at 60mins after ROSC prevented the loss of cresyl violet-stained neurons in CA1 hippocampus after ACA (Figure 1).
Fluoro-Jade staining showed neuronal degeneration in CA1 and CA2 hippocampus at 72h after ACA. Some degenerating cells were also observed in CA3, hilar region, and dentate gyrus (Figure 2). In cerebral cortex, there were sporadically distributed degenerating neurons after ACA (Figure 2). Treatment with minocycline versus vehicle, beginning 60mins after ROSC, decreased the number of degenerating neurons in CA1 and CA2 hippocampus, and cortex (Figure 2).
TUNEL positivity was observed in the CA1 and CA2 of hippocampus at 72h after ACA (Figure 3), corresponding to the degenerating neurons stained by Fluoro-Jade. In accordance with Fluoro-Jade staining, some DNA fragmentation was also observed in CA3 hippocampus and the hilar area (data not shown). However, in the cortex we did not see TUNEL positivity at this time point. Minocycline treatment attenuated TUNEL positivity in hippocampus compared with vehicle treatment (Figure 3).
Using Iba1 antibody we evaluated the microglial response to ACA. Naïve rats had modest staining with Iba1 in hippocampus and in cortex (Figure 4). Most of the observed microglial cells had a ramified shape, indicating resting status. After ACA, Iba1 positivity increased both in the hippocampus and cortex. These Iba1-positive cells showed large and amoeboid shapes, typical of activated microglia. Interestingly, in addition to the increased distribution of activated microglial cells over the whole hippocampus, there was aggregation of large ameboid microglia along the injured CA1 hippocampus, with the appearance of neuronal engulfment, as shown in Figure 4. The microglial response was less robust in the cortex than in the hippocampus after ACA. Treatment with minocycline attenuated the microglial response both in the hippocampus and cortex (Figure 4).
Next we evaluated the cytokine and chemokine response to ACA in hippocampus in PND17 rats at 24h. As shown in Figure 5, ACA resulted in significant increase in several chemokines, including MIP-1α, RANTES, and GRO-KC. Among the proinflammatory cytokines, TNF-α levels were increased significantly after ACA versus naïve controls. Although the levels of proinflammatory cytokines IL-12 and IL-18 were increased approximately 20% after ACA compared with naïve controls; this difference was not statistically significant (data not shown). We did not find significant changes in other cytokines tested. Minocycline treatment significantly attenuated the increases in MIP-1α and RANTES induced by ACA.
Our results indicate that ACA induces selective neuronal degeneration associated with enhanced microglial response and altered cerebral cytokine levels in PND17 rats. Treatment with minocycline 60mins after ROSC attenuated neuronal death, microglial response, and increases in MIP-1α and RANTES induced by ACA. The beneficial effect of minocycline administered at 60mins after resuscitation is clinically appealing and suggests an important role for inflammation in selective neuronal death after ACA in the developing rat. The ACA in this experiment resulted in the loss of two thirds of hippocampal neurons compared with naïve PND17 animals. Post-treatment with minocycline led to significant protection decreasing neuronal loss by more than 50%. In the current study, the effect of minocycline in lowering TUNEL positivity (five-fold compared with vehicle) in hippocampus was larger compared with its effect on Fluoro-Jade labeling (three-fold compared with vehicle). This difference may have implications in understanding mechanisms of injury and identifying therapeutic targets. Relative to Fluoro-Jade-labeling, TUNEL labeling is more apoptosis-specific (Schmued and Hopkins, 2000). This is in line with the known antiapoptotic effects of minocycline in neurons realized via multiple mechanisms, including inhibition of PARP-1, expression of pro-apoptotic proteins such as Bax, and opening of mitochondrial permeability transition pore (Alano et al, 2006; Garden and Moller, 2006; Gieseler et al, 2009; Li and McCullough, 2009).
In the present study, we found that microglia, as assessed by Iba1 immunoreactivity, is increased in close proximity to the regions where degenerating neurons were observed in PND17 rats at 72h after ACA. Iba1 is a macrophage/microglia-specific, calcium-binding protein and it is involved in the Rac signaling pathway, which regulates the reorganization of the actin cytoskeleton during microglial activation (Imai and Kohsaka, 2002). On activation, the ramified resting microglia migrate toward the lesion site and transform to an amoeboid activated shape (Danton and Dietrich, 2003; Yrjanheikki et al, 1998). The activated microglia would either engulf unhealthy neuronal cells or release chemicals, which might be detrimental to neuronal cells or regulate the recruitment of other immunocytes (Danton and Dietrich, 2003; Yrjanheikki et al, 1998). Despite an increase in Iba-1-positive cells in response to ACA, we did not observe a change in the levels of GM-CSF, one of the most potent microglia mitogenic factors, in vitro (Garden and Moller, 2006; Suh et al, 2005). This suggests a role for microglia recruitment mechanism rather than increase in the local proliferation of microglial cells. This is further supported by the lack of post-asphyxial changes of several cytokines, IL-1β, IL-4, and IFN-γ, known stimulators of microglial proliferation (Garden and Moller, 2006). We recognize, however, that microglial activation, proliferation, and migration are each differentially regulated and that the inflammatory mediators that we measured represent only a portion of important factors governing these three processes. Other mediators such as brain-derived neurotrophic factor and neurotrophin-3 can be released by activated microglia and act in a paracrine manner as microglia mitogens (Elkabes et al, 1996). The expression of brain-derived neurotrophic factor and its receptor tyrosine kinase-B was reported to be increased within the first day after global cerebral ischemia in adult rat (Kiprianova et al, 1999; Lindvall et al, 1992).
The patterns of microglia activation and inflammation associated with brain injury are dependent on the type of insult and age. In adult rats, it has been shown that microglia contribute to delayed neuronal death after forebrain ischemia (Denes et al, 2007; Gehrmann et al, 1992a,1992b; Liu et al, 2001; Stoll et al, 1998; Wang et al, 2008). In neonatal hypoxia–ischemia, intense expression of microglial antigens occurs with an acute or delayed time course depending on the brain region (Beilharz et al, 1995; McRae et al, 1995; Sheldon et al, 1996). Microglial activation and accumulation of macrophages have been documented early (within hours) after injury (Zhang et al, 1997), with a prolonged time course up to months after focal cerebral ischemia in rodents and humans (Gerhard et al, 2000; Schroeter et al, 2001; Tomimoto et al, 1996). However, it remains to be determined whether therapies targeting microglial activation improve neurological outcome. In this study, treatment with minocycline attenuated the microglial response and preserved neural cells in vulnerable regions. suggesting an important role for microglia in ACA-induced neurodegeneration in PND17 rats.
Hypoxic–ischemic insults induce proinflammatory cytokine production in both adult and immature brain (Cowell et al, 2002; Denker et al, 2007; Gregersen et al, 2000; Jander et al, 2002). These cytokines can contribute to secondary neurotoxicity or exacerbate ischemic brain damage by recruiting inflammatory cells such as microglia, macrophages, and neutrophils (Kriz, 2006; Minami and Satoh, 2003; Trendelenburg, 2008). Bona et al (1999) found that common carotid artery ligation resulted in increases of mRNA of several chemokines, such as MIP-1α and RANTES, in PND7 rats. The mRNA for the proinflammatory cytokine TNF-α was also increased after common carotid artery ligation in PND7 rats. Most of the chemokines and cytokines peaked at 12 to 24h after the insult, which is in agreement with our results that ACA results in significant increase of TNF-α, MIP-1α, RANTES, and GRO-KC levels in PND17 rats at 24h. Treatment with minocycline after ACA reduced the MIP-1α levels in the hippocampus at 24h and significantly attenuated the microglial response at 72h. We also found that minocycline treatment reduced RANTES levels in hippocampus induced by ACA.
The effects of minocycline on the post-asphyxial content of chemokines are likely because of its primary protective action on neurons secondarily leading to decreased recruitment of microglial cells (Duan et al, 2009; Garden and Moller, 2006; Kurpius et al, 2007; Rogove et al, 2002). Alternatively, the primary effects of minocycline could be directed toward microglia (Kremlev et al, 2004), whereby the response of different chemokines was unequal. Indeed, we found that not all the inflammatory mediators tested were affected by minocycline treatment at 24h after ACA. Whereas minocycline decreased the induction of MIP-1α, TNF-α induction was not affected. As MIP-1α is one of the important chemokines for microglia recruitment (Cowell et al, 2002; Wang et al, 2008), it is possible that inhibition of recruitment of microglia through MIP-1α-dependent pathways is one of the mechanisms responsible for the neuroprotective effects of minocycline. Decreased microglial response and neuronal death observed at a later time point (72h) after ACA supports this interpretation. It should be noted, however, that several studies documented inhibition of microglial expression of p38, an important target for minocycline, thus favoring a possible direct effect of the drug on microglial cells (Tikka et al, 2001; Tikka and Koistinaho, 2001). Finally, other cell types, in addition to microglia and neurons, may be affected by minocycline. For example, RANTES is primarily regulated and secreted by T cells, which can cross the blood–brain barrier after cerebral ischemia (Terao et al, 2008).
Minocycline has been used to ameliorate microglial activation (Jantzie et al, 2005; Wang et al, 2002; Yrjanheikki et al, 1998). It has very good blood–brain barrier penetration, has rapid absorption and adequate tissue levels within hours after oral or parenteral administration, and has a low side-effect profile making it an attractive agent for clinical use. Minocycline has been shown to have neuroprotective effects by attenuating microglial activation in adult global and focal ischemia models (Wang et al, 2002; Yenari et al, 2006; Yrjanheikki et al, 1998, 1999). A recent clinical trial indicated a protective effect of minocycline in acute stroke (Lampl et al, 2007). Compared with adults, there are limited data available on the use of minocycline in juvenile brain. The inflammatory response in the immature brain is more robust than that in the adult and is characterized by greater elaboration of cytokines (Potts et al, 2006). In addition, the developing brain is less able to detoxify reactive oxygen and nitrogen species generated by inflammatory cells because of inadequate expression of certain antioxidant molecules (Bayır et al, 2006). Thus, inflammatory response may be a rational therapeutic target in the immature brain. In line with this, minocycline treatment has been shown to attenuate neuroinflammation and improve neurobehavioral performance after unilateral carotid artery ligation and hypoxia model in PND3 to PND7 rats (Carty et al, 2008; Fan et al, 2006; Lechpammer et al, 2008; Leonardo et al, 2008). However, the beneficial effects of minocycline likely depend on the type of insult, since its protective effects were reported to be transient after middle cerebral artery occlusion in PND7 rats (Fox et al, 2005). To our knowledge, our investigation is the first study evaluating the effect of minocycline after ACA in the immature brain. Further studies are needed to evaluate whether these beneficial effects of minocycline observed early after ACA in juvenile brain will have long-lasting consequences weeks after the insult by comprehensive functional outcome testing.
In conclusion, microglia activation and cytokine production are increased in immature brain after ACA. Minocycline exerted beneficial effects on neuronal survival associated with decreased microglial and chemokine response after ACA in the immature brain. The beneficial effect of minocycline suggests an important role for inflammation in selective neuronal death after pediatric ACA, and a possible therapeutic avenue. Further research is necessary in this model to elucidate the mechanisms that provoke neuroinflammation and the interactions of inflammatory cascade with cells susceptible to injury. Understanding such mechanisms could aid in the design of new pharmacological strategies aimed at reducing neurological sequela after CA in infants and children. Finally, given that minocycline is an FDA-approved medication, it merits additional study in ACA for possible clinical translation.
This work was supported by grants from American Heart Association (0535365N) and NIH (NS061817, NS30318, HD057587, and HD045968).
Conflict of interest
The authors declare no conflict of interest.