A Paraventricular Nucleus–Rostral Ventrolateral Medulla Pathway Contributes to Myocardial Ischemia/Reperfusion Injury

doi:10.21203/rs.3.rs-2678309/v1

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A Paraventricular Nucleus–Rostral Ventrolateral Medulla Pathway Contributes to Myocardial Ischemia/Reperfusion Injury

https://doi.org/10.21203/rs.3.rs-2678309/v1

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Myocardial ischemia/reperfusion injury (MIRI), the major pathophysiology of cardiovascular disease, is a crucial therapeutic focus. To date, whether MIRI is centrally mediated and its underlying processing hierarchy remain elusive. We show that the electrical activity of the paraventricular nucleus (PVN) and rostral ventrolateral medulla (RVLM) neurons increased after MIRI in a mouse model. We identified a neural circuit involving glutamatergic projections from the paraventricular nucleus (Glu^PVN) to tyrosine hydroxylase–expressing neurons in the rostral ventrolateral medulla (TH^RVLM) that contributes to MIRI. Transneuronal tracing with neurotropic viruses indicated that the TH^RVLM neurons project directly to the spinal preganglionic neurons and then to the stellate ganglion, two critical neural nodes along the brain–heart axis. Chemogenetic inhibition of the Glu^PVN→TH^RVLM circuit or cervical sympathetic blockade reduced the level of norepinephrine in the heart and thereby prevented MIRI. Furthermore, pharmacological blockade of myocardium β-receptors also reduced MIRI. This brain–heart circuit that promotes MIRI represents a potential therapeutic target for MIRI treatment.

Health sciences/Diseases/Cardiovascular diseases/Acute coronary syndromes/Myocardial infarction

Biological sciences/Neuroscience

Biological sciences/Physiology/Cardiovascular biology/Cardiovascular diseases/Acute coronary syndromes/Myocardial infarction

Health sciences/Anatomy/Nervous system

Timely and effective reperfusion after acute myocardial ischemia is a standard therapeutic strategy^1,2. Unfortunately, accumulating evidence has shown that myocardial reperfusion can itself cause further cardiomyocyte death, known as myocardial ischemia/reperfusion injury (MIRI). The cardiac dysfunction during MIRI includes myocardial stunning, the no-reflow phenomenon, reperfusion arrhythmias, and lethal reperfusion injury. Both basic and clinical studies have shown that MIRI accounts for almost 50% of the final myocardial infarction size^1,3. Despite the increasing use and improved methodology of interventional approaches for reperfusion, the lethality of myocardial ischemia remains substantial^4,5. Although various nonpharmacological and pharmacological clinical interventions are used to reduce MIRI, such as ischemic pre-conditioning, cyclosporine-A, and metoprolol⁶, their therapeutic effects have been disappointing. The lack of clear-cut success in clinical trials may be due to a lack of knowledge about the regulatory mechanism for MIRI. Previous studies have revealed several molecular and cellular mechanisms that lead to MIRI, including oxidative stress, energy metabolism dysfunction, and necrosis in the injured myocardium^7,8. However, there seems to be a lack of consideration of MIRI from a holistic perspective, such as the role of the central nervous system in regulating MIRI.

Recently, the cardiac sympathetic nerves (CSNs) have been found to be activated during the process of myocardial ischemia/reperfusion (MIR)^9–12. The development and progression of MIRI are always associated with sympathetic excitation^13,14. Activation of CSNs increases myocardial contractility and the blood supply to the myocardium, which may further aggravate MIRI⁹. Inhibition of CSNs excitability provides a cardioprotective effect against MIRI^15,16. Norepinephrine (NE), the major neurotransmitter released by CSNs, has been shown to cause oxidative stress and induces myocardium apoptosis in rodent models^17,18. Clinical studies have also shown that blocking the β-adrenergic receptor reduces myocardial injury in patients with MIRI^19–23, suggesting the involvement of CSNs in MIRI. The involvement of sympathetic nerves as a vital connection between the central and peripheral organs led us to speculate that the brain–heart axis may play a key role in regulating MIRI.

The brain controls the cardiovascular system through the autonomic nervous system²⁴. The release of neurohormones by the autonomic nervous system has diverse downstream effects on the cardiovascular system, such as platelet activation, proarrhythmic consequences, and vasoconstriction²⁵. Researchers have realized the impact of primary brain disorders that lead to cardiac dysfunction (i.e., the brain–heart axis)^26,27. Studies have also shown the close associations between emotional stress and psychosocial risk factors with subsequent progression of coronary artery disease, arterial inflammation, and adverse clinical outcomes^28–30. Neuroanatomical and neurophysiologic studies have demonstrated that brain areas including the anterior insula, anterior cingulate cortex, amygdala, hypothalamus, periaqueductal gray (PAG), parabrachial nucleus, and several regions of the medulla may be able to control cardiac function. These areas are critically involved in emotional behavior, stress responses, and homeostatic reflexes and exert their influence on cardiac function via the sympathetic and parasympathetic nervous systems³¹. For instance, the nucleus of the solitary tract (NTS) is activated by blood pressure elevation, which initiates various compensatory changes in cardiac and vasomotor activity to reduce blood pressure^32,33. Important regions for the central integration of sympathetic outflow such as the paraventricular nucleus (PVN) and the rostral ventrolateral medulla (RVLM) are also associated with cardiovascular function^34–37. However, whether or how these regions are involved in regulating MIRI is still unclear.

Here we have combined cell-type-specific viral tracing approaches, in vivo electrophysiological recording, and other methods in a mouse MIRI model to accurately elucidate the brain nuclei and neural circuits that regulate MIRI. In addition, we explored whether the treatment of MIRI could be achieved by manipulating the relevant neural circuits through chemogenetic and pharmacological methods. In conclusion, we demonstrate that the PVN glutamatergic neurons (Glu^PVN)→RVLM tyrosine hydroxylase–expressing neurons (TH^RVLM)→CSNs pathway represents the hierarchical neural architecture underlying MIRI, which provides new neural interventional targets for MIRI.

Unraveling the pivotal supraspinal regions associated with MIRI

We first constructed the MIRI model by ligating the left anterior descending (LAD) coronary artery in the hearts of C57BL/6J mice^38,39. At the end of reperfusion, blood samples from the ventricle were collected for ELISA analysis. Meanwhile, the heart was collected for triphenyl-tetrazolium chloride (TTC) staining (Fig. 1a). We next examined the myocardium infarct size relative to the area at risk (IS/AAR) and serum cardiac troponin I (cTnI) concentration, two gold standards of myocardium injury^40,41. Both the IS/AAR and serum cTnI concentration were significantly increased in MIRI mice (Fig. 1b–d and Extended Data Fig. 1a), indicating that the MIRI model was successfully established. Furthermore, the MIRI mice also exhibited a higher serum norepinephrine (NE) level than did the sham mice, suggesting an enhanced sympathetic excitability⁴²(Fig. 1e).

We then asked whether this MIRI-caused sympathetic hyperexcitability can be centrally regulated. To answer this question, we injected pseudorabies virus (PRV) that encodes enhanced green fluorescent protein (EGFP) under the control of the constitutive CAG promoter (PRV-CAG-EGFP) directly into the apex region of the heart to retrogradely and trans-synaptically label neurons upstream of the heart in the central nervous system (Fig. 1f). Viral-induced fluorescence was detected in the myocardium of the left ventricle (LV) within 24 to 48 h of PRV injection (Fig. 1f). During that time, the EGFP signal was also detected in the stellate ganglion (SG) and in lamina VII of the spinal cord (SC VII) (Fig. 1g, h). More importantly, at 72–96 h after PRV injection, the EGFP signal was also observed in various brain regions such as the NTS; lateral reticular nucleus (LRt); medial vestibular nucleus, parvicellular part (MVePC); posterior hypothalamic nucleus (PH); lateral hypothalamic area (LH); intermediate nucleus of the lateral lemniscus (ILL); and PAG, especially regions associated with the sympathetic nerves, such as the PVN and RVLM (Fig. 1i, j and Extended Data Fig. 2a, b).

To further validate the involvement of these PRV-labeled regions in MIRI, we examined their c-Fos expression levels, a proxy for neuronal activity⁴³. There was a substantial increase in the c-Fos expression level in the SG, SC VII, RVLM, and PVN of the MIRI mice as compared with the sham mice (Fig. 1k, l and Extended Data Fig. 2c). Our PRV trans-synaptic retrograde tracing and c-Fos detection experiments both suggested the importance of the PVN and RVLM, two vital brain regions for the central integration of sympathetic outflow^34–37. Therefore, we speculated that the PVN and RVLM may be the pivotal nuclei for regulating MIRI.

PVN and RVLM neurons are preferentially activated during MIR

We then examined the real-time activity of PVN and RVLM neurons before and after MIR based on in vivo multi-channel recordings (Fig. 2a, f). A total of 25 PVN neurons from six mice and 20 RVLM neurons from eight mice were recorded, and their firing patterns were consistent with previous studies^44,45(Fig. 2b, g). Notably, reperfusion significantly increased the neuronal spike rate in the PVN (Fig. 2c, d) and RVLM (Fig. 2h, i). The average spontaneous spike rate increased from 3.61 ± 0.98 Hz to 11.03 ± 2.46 Hz in PVN neurons (Fig. 2e) and from 3.58 ± 0.73 Hz to 7.91 ± 2.08 Hz in RVLM neurons (Fig. 2j). These results indicate that PVN and RVLM nuclei are activated after MIR.

Ablation or inactivation of PVN and RVLM neurons reduces MIRI

Next, we ablated neurons of the PVN or RVLM by injecting an AAV that conditionally expresses cell death–inducing active caspase 3 (taCasp3) (Fig. 3a, f). The ablation efficiency was 66.0% in the PVN and 76.8% in the RVLM after intracranial injection of AAV-flex-taCasp3-TEVp virus with an AAV that expresses Cre recombinase (AAV-Cre virus) (Fig. 3b, g). Ablation of either PVN or RVLM neurons reduced MIRI, as reflected by the significant reduction in IS/AAR and serum cTnI concentration (Fig. 3c, d and 3h, i). Ablation of PVN or RVLM neurons also decreased sympathetic excitability, as indicated by the significant reduction in serum NE concentration (Fig. 3e, j).

Next, we inhibited the activity of the PVN or RVLM by using DREADDs (designer receptors exclusively activated by designer drugs) approach. Briefly, a recombinant AAV that conditionally expresses the inhibitory receptor hM4Di (AAV-CaMKIIa-hM4Di-EGFP) or a recombinant AAV without hM4Di (AAV-CaMKIIa-EGFP) as the control was bilaterally injected into the PVN or RVLM (Fig. 3k, p). Injection sites in the PVN and RVLM were verified by EGFP expression (Fig. 3l, q). Four weeks after virus injection, the designer drug clozapine N-oxide (CNO) was injected intraperitoneally (i.p.) 60 min before MIR. When activated by CNO, both PVN- and RVLM-expressed hM4Di significantly reduced the resulting MIRI (Fig. 3m–o and 3r–t). These results indicate that PVN and RVLM are functionally regulating MIRI development.

The PVN→RVLM pathway contributes to MIRI

Accumulating evidence suggests that PVN neurons directly project to the RVLM^46,47. As both the PVN and RVLM are activated during MIRI, we wondered whether the PVN→RVLM projection contributes to MIRI. To further confirm the existence of this pathway, we delivered the retrograde transport virus Retro-AAV-EGFP into the RVLM (Fig. 4a). Four weeks later, EGFP was well expressed in the RVLM, indicating the accurate injection of the viral vector (Fig. 4b). Meanwhile, the EGFP signal was observed in the whole PVN and other nuclei (Fig. 4c, d and Extended Data Fig. 5). EGFP signals were more abundant in the PVN ipsilateral to the site of RVLM injection relative to those in the contralateral PVN (Fig. 4c, d). Among all the PVN→RVLM projecting neurons, EGFP⁺ PVN neurons co-labeled for c-Fos expression represented 9.4% of all EGFP⁺ PVN neurons in sham mice (Fig. 4f, g), which was increased to 65.3% in MIR mice (Fig. 4h, i), suggesting an activation of the PVN→RVLM projection in MIRI.

To further evaluate the role of the PVN→RVLM projection in MIRI, we conditionally inhibited this projection through bilateral injection of AAV-CaMKIIa-hM4Di-EGFP into the PVN of C57BL/6J mice or of AAV-CaMKIIa-EGFP into the same region as a control (Fig. 4j, k). Three weeks after virus injection, two cannulas were bilaterally implanted onto the RVLM area of the mice for CNO injection. Four weeks after virus injection, CNO was microinjected into the RVLM via the cannulas (Intra. CNO) 20 minutes before MIR. Chemogenetic inactivation of the PVN→RVLM projection after intra-RVLM injection of CNO dramatically alleviated MIRI (Fig. 4l–o). Thus, these results indicate that activation of the PVN→RVLM projection promotes MIRI.

The RVLM tyrosine hydroxylase–expressing (TH^RVLM) neurons contribute to MIRI

TH^RVLM neurons are essential for sympathetic outflow^48,49. We thus used virus tracing, immunostaining, transgenic mice, and chemogenetic tools to clarify the role of the TH^RVLM neurons in the PVN→RVLM→CSN pathway in MIRI. We first injected PRV-CAG-EGFP into the apex region of the heart to retrogradely label neurons in the RVLM that project to the heart (Fig. 5a). At 72 h after virus injection, abundant EGFP signals were observed in the RVLM, and 65.3% of the EGFP⁺ RVLM neurons were co-labeled for TH expression (Fig. 5b), suggesting that most RVLM neurons that project to the heart were TH⁺ neurons. Next, a mixture of AAV-hSyn-Cre and AAV-DIO-EGFP was unilaterally co-injected into the PVN to test whether PVN neurons can directly project to the TH^RVLM neurons (Fig. 5c). Four weeks later, the correct injection site in the PVN was verified by EGFP (Fig. 5d). Meanwhile, EGFP signals were observed in the RVLM (Fig. 5e) and were closely associated with cell bodies and dendrites of the TH^RVLM(Fig. 5f), suggesting an anatomical connection between the PVN and the TH^RVLM.

Further, to confirm whether the TH^RVLM neurons projecting from the PVN are involved in the regulation of MIRI, a Cre-dependent anterograde monosynaptic tracing virus, AAV2/1-Cre, was injected into the PVN. In addition, AAV2/8-DIO-mCherry was injected into the RVLM of C57BL/6J mice. Four weeks later, we injected PRV-CAG-EGFP into the apex region of the heart to retrogradely label the neurons in the RVLM that project to the heart (Fig. 5g). At 72 h after PRV injection, mCherry signals anterograde monosynaptic projecting from PVN were co-labeled with EGFP expression in RVLM neurons, and these neurons were mainly TH⁺ neurons (Fig. 5h), providing direct anatomic evidence for the existence of a PVN→TH^RVLM→heart pathway.

We next examined the role of TH^RVLM neurons in MIRI using dopamine β-hydroxylase (DBH)-Cre transgenic mice in which Cre is specifically expressed in the TH⁺ neurons⁴⁷. AAV-DIO-hM4Di-mCherry or the control AAV-DIO-mCherry was bilaterally injected into the RVLM of DBH-Cre mice. Four weeks later, CNO was injected i.p. 60 minutes before MIR (Fig. 5i). Appropriate targeting of virus injections in the RVLM was verified post hoc by marking the injection sites with mCherry (Extended Data Fig. 7b). Inhibition of TH^RVLM neurons by CNO reduced the MIRI, as indicated by the significant reduction in the IS/AAR and serum cTnI and serum NE concentrations (Fig. 5j–m). Collectively, these findings suggest that TH^RVLM neurons play a pivotal role in MIRI.

The Glu^PVN→TH^RVLM projection contributes to MIRI

We next identified the subtype of neurons projecting to TH^RVLM neurons in the PVN. Retro-AAV-EGFP was unilaterally injected into the RVLM (Extended Data Fig. 8a). Four weeks later, many retrogradely labeled EGFP⁺ neurons were observed in the PVN, and 78.6% of these were glutamate (Glu) immunopositive (Extended Data Fig. 8b, c). Few EGFP⁺ neurons were GABA or TH immunopositive (Extended Data Fig. 8d–g). Retro-AAV-DIO-EGFP was then injected into the RVLM of Vglut2-Cre mice in which the Cre is specifically expressed in the glutamatergic neurons (Fig. 6a). Four weeks later, mice underwent MIR, and c-Fos expression in PVN EGFP⁺ neurons was examined to determine whether such Glu^PVN neurons are activated during MIRI. Appropriate targeting of virus injections in the RVLM was verified post hoc by marking injection sites with EGFP (Fig. 6b, e). Among the retrogradely traced EGFP⁺ Glu^PVN neurons, 11.1% were co-labeled for c-Fos expression in sham mice (Fig. 6c, d), which increased to 71.6% in MIR mice (Fig. 6f, g), suggesting an activation of Glu^PVN neurons during MIRI. Furthermore, extensive EGFP fluorescence was closely associated with the cell bodies and dendrites of TH^RVLM neurons after unilateral injection of AAV-DIO-EGFP into the PVN of Vglut2-Cre mice (Fig. 6h–k), indicating a glutamatergic projection from the PVN to TH-expressing neurons in the RVLM (Glu^PVN→TH^RVLM).

Next, chemogenetic manipulation was used to test the role of the Glu^PVN→TH^RVLM projection in MIRI. In brief, AAV-DIO-hM4Di-mCherry was bilaterally injected into the PVN of Vglut2-Cre mice (Fig. 6l). Four weeks later, CNO was microinjected into the RVLM 20 minutes before MIR (Fig. 6l). Appropriate targeting of virus injections in the PVN was verified post hoc by marking injection sites with mCherry (Extended Data Fig. 9b). Inhibition of the Glu^PVN→TH^RVLM pathway by injecting CNO into the RVLM relieved MIRI, as reflected by the significantly reduced IS/AAR and serum cTnI and NE concentrations (Fig. 6m–p). Together, these findings suggest that activation of the Glu^PVN→TH^RVLM projection exacerbates MIRI.

Downstream nodes of the Glu^PVN→TH^RVLM pathway modulate MIRI

Our above results indicated that MIRI is associated with cardiac sympathetic hyperexcitation. As CSNs always receive inputs from spinal preganglionic neurons (SPNs) and SG neurons^50–53, we therefore wondered whether the PVN-projecting TH^RVLM neurons can directly innervate the SPNs and SG neurons. To achieve this, we first unilaterally injected an anterograde monosynaptic tracing virus, AAV2/1-DIO-FLP, accompanied by cholera toxin subunit B (CTB) into the PVN and AAV-fDIO-EGFP-2a-Synaptophysin-mRuby into the ipsilateral RVLM of DBH-Cre mice (Fig. 7a). Appropriate targeting of virus injections into the PVN was verified post hoc by CTB, which marked the injection sites (Fig. 7b). Similarly, the EGFP signal was restricted to the RVLM based on the trans-synaptic spreading of DIO and FLP (Fig. 7c). Moreover, mRuby⁺ presynaptic terminals of the TH^RVLM were observed in SC VII (Fig. 7d), an area containing a massive number of SPNs that project to the SG⁵⁴.

To further demonstrate the anatomical connection between TH^RVLM and the SG, we unilaterally injected an anterograde monosynaptic tracing virus, AAV-DIO-WGA-FLP, accompanied by CTB into the RVLM and injected AAV-fDIO-EGFP-2a-Synaptophysin-mRuby into SC VII of DBH-Cre mice (Fig. 7e). Appropriate targeting of virus injections in the RVLM was verified post hoc by CTB (Fig. 7f), and the EGFP signal was restricted to SC VII (Fig. 7g). Notably, mRuby signals from the SPNs in SC VII were observed in the SG (Fig. 7h). These findings suggest that SPNs and the SG are downstream of the Glu^PVN→TH^RVLM pathway.

Next, we examined the function of the SPN→SG projection in MIRI. As SPNs send nerve projections through the cervical sympathetic trunk (CST) to the SG neurons, we transected the left or right CST to evaluate the role of the SPNs→SG pathway in MIRI (Fig. 7i). As compared with the control group of mice, transection of the CST alleviated MIRI (Fig. 7j), as reflected by the significantly reduced IS/AAR (Fig. 7k) and serum cTnI (Fig. 7l) and serum NE (Fig. 7m) concentrations.

Activation of CSNs always causes massive release of NE in the heart^55,56, which has also been evidenced in our MIRI model. Based on this, we examined whether blockade of the NE receptor in the myocardium affects MIRI. Metoprolol, a β-adrenoceptor blocker, was injected intracardially 10 minutes before MIR (Fig. 7n). Metoprolol administration reduced MIRI (Fig. 7o), as indicated by the significant decrease in IS/AAR (Fig. 7p) and serum cTnI concentration (Fig. 7q). However, the metoprolol failed to affect the serum NE concentration (Fig. 7r). Taken together, these findings reveal a Glu^PVN→TH^RVLM→CSNs pathway that modulates MIRI.

Current clinical treatments for MIRI are far from satisfactory, and there is an urgent need to better understand the regulatory mechanisms involved in this process. Recently, therapeutic approaches targeting sympathetic excitement caused by MIRI such as β-receptor blockers, spironolactone, nicorandil, and high thoracic epidural anesthesia have been used in clinical trials. Studies have indicated that these approaches show beneficial effects on myocardial injury and ventricular remodeling in patients with acute MIR^{19–23,57−62}. Although sporadic clinical and animal studies have reported that some brain areas, including the PVN, RVLM, and subthalamic nucleus (STN), may affect the cardiovascular outcome by targeting central sympathetic outflow^63–65, research regarding the central hierarchy that affects MIRI is still very limited. In this study, we identified a novel Glu^PVN→TH^RVLM→CSNs pathway that contributes to the development of MIRI. In vivo multi-channel recordings showed that the activation of PVN and RVLM neurons was increased during MIR. Both pharmacological and chemogenetic inhibition of the Glu^PVN→TH^RVLM→CSNs pathway were able to reduce MIRI.

There is some evidence that PVN neurons send projections directly to the RVLM, the region widely known for regulating sympathoexcitation and blood glucose content^46,47,49. However, the neuronal subtypes involved and the function of this pathway during MIRI have not been investigated. Here we first confirmed that the PVN and RVLM neurons were preferentially activated during MIR. In vivo multi-channel recordings showed that the spike rate of PVN and RVLM neurons immediately increased at the onset of reperfusion and gradually recovered after 20 seconds. This may be associated with the rapid influx of noxious substances produced by ischemia into the ischemic myocardium at the onset of reperfusion⁶⁶. In development, we identified a neural pathway involving glutamatergic projections from the PVN to TH-expressing neurons in the RVLM (Glu^PVN→TH^RVLM) that contributes to the development of MIRI. However, it remains unclear how this Glu^PVN→TH^RVLM pathway is activated during MIR. Some previous studies have reported that PVN neurons are activated under various pathological conditions such as pain, stress, and inflammation^67–70. Based on this, it is likely that the angina and stress response during MIR lead to the activation of PVN neurons, which then activate the downstream RVLM neurons and the CSNs.

NE, the major neurotransmitter released from the CSNs, aggravates myocardial injury during MIR^71,72. Metoprolol, the widely used blocker of NE receptors, has also been shown to reduce myocardial injury in patients with acute MIR^19–23. Here we found that activation of the supraspinal Glu^PVN→TH^RVLM pathway accelerated the release of NE in the heart during MIR. This pathway also showed a strong connection with SC VII, and then with the SG and CSNs. It is likely that activation of this multilevel neural pathway is one main cause for the increase in NE release and myocardial injuries during MIR. However, the neuronal subtype in SC VII and the SG of this circuit is not investigated. In addition, the parasympathetic nervous system has also been reported to regulate MIRI^73–75. Thus, further research is needed to resolve the central and peripheral mechanisms underlying MIRI.

Although various clinical strategies have been attempted to lessen MIRI over the past decades, their therapeutic effects remain disappointing. For instance, ischemia pre-conditioning (IPC) can limit the size of myocardial infarcts that caused by MIRI⁷⁶. However, there are operational and ethical limitations to the implementation of IPC. Several pharmacological interventions have been tested in the research laboratory with some encouraging results, but their translation to the clinical setting has been largely disappointing⁷⁷. In addition, some trials using pharmacological agents have suggested that only patients with ischemia of a short duration can benefit⁷⁸. In the present study, we explored MIRI from a holistic perspective. We provide direct anatomical evidence for the existence of a sympathetic axis between the brain and heart and suggest that a hierarchical neural architecture contributes to MIRI. Furthermore, some other basic and clinical studies have also shown that therapeutic strategies targeting the CSNs are also able to prevent life-threatening arrhythmias and myocardial injury^{19–23,79−81}. Thus, our findings provide new insights indicating that the Glu^PVN→TH^RVLM→CSNs pathway is a promising clinical target for reducing MIRI.

Animals. All animal procedures and protocols used in this study were approved by and performed in accordance with the guidelines of the Laboratory Animal Ethics Committee of Anhui Medical University (No. LLSC20190476). In all experiments, C57BL/6J mice (purchased from Beijing Vital River Laboratory or Jackson Laboratories) and Vglut2-Cre and DBH-Cre mice (obtained from Wei Xiong, Institute of Artificial Intelligence, Hefei Comprehensive National Science Center) at 8–10 weeks of age were used. All animals were housed in an environment with a temperature of 22 ± 1ºC, relative humidity of 50 ± 1%, and a light/dark cycle of 12 h/12 h.

Animal models of myocardial ischemia/reperfusion injury (MIRI). Surgery was performed after the mice were anesthetized with intraperitoneal administration of urethane (800 mg/kg) and α-chloralose (40 mg/kg), and with a maintenance dose (200 mg/kg urethane and 10 mg/kg α-chloralose) repeatedly injected i.p. every hour. The procedure was as described for experimental models of myocardial ischemia and infarction^82,83. Briefly, all mice underwent tracheal intubation. Mechanical ventilation was provided with an Apparatus Rodent Respirator (R407, RWD Life Science, Shenzhen, China), and the mice were ventilated with room air at 110–120 breaths/min. Body temperature was monitored and maintained at 37 ± 1ºC using a heating pad. Subcutaneous stainless steel electrodes were connected to a PowerLab monitoring system to monitor the lead II electrocardiogram and heart rate. A left thoracotomy was performed to expose the heart at the third and fourth intercostal spaces; an 8 − 0 Prolene loop was placed at the origin of the left coronary artery. After surgical preparation, the mice were stabilized for 10 min. Regional ischemia and reperfusion were induced by ligating and releasing the Prolene loop, respectively. MIRI was induced by exposing animals to 60 min of ischemia by occlusion of the left anterior descending (LAD) coronary artery followed by 120 min of reperfusion by release of the occlusion. Ischemia was confirmed by electrocardiographic changes and cardiac cyanosis.

Transection of the cervical sympathetic trunk (CST). In individual mice, either the right or left CST was transected to induce sympathetic blockade. A 2-cm vertical incision was created in the neck region. Then, the right or left sternomastoid muscle and omohyoid muscle were separated (not transected). The right or left common carotid artery was exposed, and the CST was identified along the proximal side of the superior cervical ganglion⁸⁴. The right or left side of the cervical sympathetic trunk was transected, and the incision was closed. Sympathetic blockade was confirmed by blepharoptosis on the operated side.

Myocardial infarction analysis. Infarct size of each heart in this experiment was calculated as described⁸⁵. Briefly, at the end of the reperfusion period, the LAD suture was re-tightened, and 1 ml of 0.25% Evans blue dye was perfused through the heart to delineate the area at risk of infarction. The hearts were then frozen at − 20ºC before being sectioned into four transverse slices and stained for viable tissue by immersion in 1% triphenyl-tetrazolium chloride (Sigma, USA) at 37ºC for 15 min. After fixation in 4% formalin for 24 h, the sections were digitally scanned for analysis. Analysis of infarct size (IS) as a proportion of the area at risk (AAR) was calculated via planimetry using ImageJ software (version 1.45, National Institutes of Health, USA).

Enzyme-linked immunosorbent assay (ELISA). For ELISA of serum cTnI and NE, blood samples from the ventricle were collected at the end of reperfusion. Samples were then centrifuged at 12000 g for 10 min. The serum was collected for evaluation of the cTnI and NE concentration using ELISA kits (Elabscience, Wuhan, China).

Pseudorabies virus (PRV) trans-synaptic retrograde tracer technique. Mice were anesthetized with 3% isoflurane and ventilated. Anesthesia was maintained with 1% isoflurane. The left third and fourth intercostal spaces were opened by incision, and then a pericardial incision was made to expose the apex of the heart. PRV-CAG-EGFP (2.00 × 10⁹ PFU/ml, 0.2 µl/site; BrainVTA, Wuhan, China) was injected directly into four sites of the ventricular myocardium in the apex region. The wound was closed using cutaneous interrupted sutures. The mice were sacrificed every 24 h, and tissues including the heart, SG, spinal cord, and brain were collected and fixed with paraformaldehyde, followed by a frozen sectioning process, which was used to view the autofluorescence of the PRV.

In vivo electrophysiological recordings. Neuronal recordings were performed at a 30-kHz sampling rate using the Cereplex µ headstage, a digital hub, and a neuronal signal processor (Blackrock Microsystems). For the RVLM and the PVN recordings, urethane (800 mg/kg) and α-chloralose (40 mg/kg) were administrated i.p. to induce anesthesia, with a maintenance dose (200 mg/kg urethane and 10 mg/kg α-chloralose) repeatedly injected i.p. every hour. Mice were fixed in a stereotaxic frame (RWD Life Science, Shenzhen, China), the skull and dura were removed at coordinates overlying the PVN nucleus (bregma, − 0.80 mm; lateral, ± 0.13 mm; and dura, − 4.55 mm) or RVLM nucleus (bregma, − 6.65 mm; lateral, ± 1.20 mm; and dura, − 4.40 mm). After an electrode was fixed to the PVN or RVLM with dental cement, the MIR surgery was performed. After surgical preparation, the mice were stabilized for 10 min. Signals were recorded continuously during ischemia/reperfusion after a 15-min period of baseline recording. To verify the recording sites, mouse brains were removed and stained with DAPI.

Spike sorting. Neurons were isolated offline using BOSS software (Blackrock Microsystems). A notch filter was applied to eliminate the line noise, then a high-pass filter of 250 Hz was applied, and a threshold multiplier was used to extract spikes from the signal energy between 1 and 5 kHz. We then used principal component analysis to extract spike features and to manually cluster the spikes in different units. We eliminated spikes that did not have a refractory period of 2–3 ms and spikes that were simultaneously recorded in 10 or more channels (as potential artifacts).

Spike train analysis. Sorted files were then processed in Neuroexplorer to extract unit timestamps and relevant event markers. These data were subsequently analyzed in MATLAB (Natick, MA). We then calculated the spike rate (T_interval = 0.1 ms, bin = 1 s) as follows: Spike rate (T_neuron) = Number_timestamps(n|T_neuron – bin ≤ n < T_neuron)/bin, where T_interval is how often it is calculated, T_neuron is the timing that we calculated the spike rate, Number_timestamps is the spike number during bin, and the bin refers to the time before T_neuron.

Stereotaxic injection. Mice were anesthetized with an intraperitoneal injection of pentobarbital sodium (0.5%, 0.01 ml/g) and mounted on a stereotaxic apparatus (68046, RWD Life Science, Shenzhen, China). Ophthalmic ointment was applied to prevent their eyes from drying out. The skull was exposed by a midline scalp incision, and a craniotomy was drilled unilaterally or bilaterally above each injection site. Viruses were delivered at a rate of 50 nl/min through the glass pipette, which was connected to a pump (KDS LEGATO 130, RWD Life Science, Shenzhen, China). After completion of the injection, the glass pipette was held in place for 10 min before being withdrawn to allow for diffusion of the viruses. The coordinates used for PVN and RVLM injection were as described above.

For the ablation of PVN or RVLM neurons, a 1:1 mixture of rAAV-hSyn-Cre-WPRE-hGH-pA (AAV2/9, 5.20 × 10¹² v.g./ml, BrainVTA) and rAAV-EF1a-flex-taCasp3-TEVp-WPRE-hGH-pA (AAV2/9, 5.13 × 10¹² v.g./ml, BrainVTA) was bilaterally injected into the PVN or RVLM of C57BL/6J mice at a volume of 150 nl for each side. A 1:1 mixture of rAAV-hSyn-Cre-WPRE-hGH-pA (AAV2/9, 5.20 × 10¹² v.g./ml, BrainVTA) and rAAV-EF1a-DIO-mCherry-WPRE-hGH-pA (AAV2/9, 5.26 × 10¹² v.g./ml, BrainVTA) was used as a control.

For chemogenetic inactivation of PVN or RVLM neurons, rAAV-CaMKIIa-hM4Di-EGFP-WPRE-pA (AAV2/9, 5.00 × 10¹² v.g./ml, BrainVTA) was bilaterally injected into the PVN or RVLM of C57BL/6J mice at a volume of 150 nl for each side. rAAV-CaMKIIa-EGFP-WPRE-hGH-pA (AAV2/9, 2.00 × 10¹² v.g./ml, BrainVTA) was used as a control.

For retrograde monosynaptic tracing of RVLM-projecting neurons, C57BL/6J mice were unilaterally injected with 150 nl of rAAV-hSyn-EGFP-WPRE-hGH-pA (AAV2/R, 5.00 × 10¹² v.g./ml, BrainVTA) into the RVLM.

For chemogenetic inactivation of PVN→RVLM projection, rAAV-CaMKIIa-hM4Di-EGFP-WPRE-pA (AAV2/9, 5.00 × 10¹² v.g./ml, BrainVTA) was bilaterally injected into the PVN of C57BL/6J mice at a volume of 150 nl for each side. rAAV-CaMKIIa-EGFP-WPRE-hGH-pA (AAV2/9, 2.00 × 10¹² v.g./ml, BrainVTA) was used as a control.

For anterograde monosynaptic tracing of the PVN→RVLM projection, C57BL/6J mice were unilaterally injected with a 1:1 mixture of rAAV-hSyn-Cre-WPRE-hGH-pA (AAV2/9, 5.20 × 10¹² v.g./ml, BrainVTA) and rAAV-EF1a-DIO-EGFP-WPRE-hGH-pA (AAV2/9, 5.00 × 10¹² v.g./ml, BrainVTA) into the PVN at a volume of 150 nl.

For trans-synaptic tracing of the PVN→TH^RVLM→heart pathway, C57BL/6J mice were unilaterally injected with 150 nl of ScAAV-hSyn-Cre-WPREs (AAV2/1, 1.10 × 10¹³ v.g./ml, BrainVTA) into the PVN and 150 nl of rAAV-hSyn-DIO-mCherry-WPRE-hGH-pA (AAV2/8, 2.58 × 10¹² v.g./ml, BrainVTA) into the ipsilateral RVLM. Four weeks later, PRV-CAG-EGFP (2.00 × 10⁹ PFU/ml, 0.2 µl/site; BrainVTA) was injected directly into four sites of the ventricular myocardium in the apex region.

For chemogenetic inhibition of TH^RVLM neurons, rAAV-EF1α-DIO-hM4Di-mCherry-WPRE (AAV2/9, 5.00 × 10¹² v.g./ml, BrainVTA) was bilaterally injected into the RVLM of DBH-Cre mice at a volume of 150 nl. For chemogenetic excitement of TH^RVLM neurons, rAAV-EF1α-DIO-hM3Dq-mCherry-WPRE (AAV2/9, 5.27 × 10¹² v.g./ml, BrainVTA) was bilaterally injected into the RVLM of DBH-Cre mice at a volume of 150 nl. rAAV-EF1a-DIO-mCherry-WPRE-pA (AAV2/9, 5.26 × 10¹² v.g./ml, BrainVTA) was used as a control for both sets of experiments.

For retrograde monosynaptic tracing of the PVN→RVLM glutamatergic projection, Vglut2-Cre mice were unilaterally injected with 150 nl rAAV-EF1a-DIO-EGFP-WPRE-hGH-pA (AAV2/R, 5.00 × 10¹² v.g./ml, BrainVTA ) into the RVLM.

For anterograde monosynaptic tracing of the PVN→RVLM glutamatergic projection, Vglut2-Cre mice were unilaterally injected with 150 nl rAAV-EF1a-DIO-EGFP-WPRE-hGH-pA (AAV2/9, 5.00 × 10¹² v.g./ml, BrainVTA) into the PVN.

For chemogenetic inhibition of Glu^PVN→TH^RVLM projection, rAAV-EF1α-DIO-hM4Di-mCherry-WPRE (AAV2/9, 5.00 × 10¹² v.g./ml, BrainVTA) was bilaterally injected into the PVN of Vglut2-Cre mice at a volume of 150 nl. For chemogenetic excitement of Glu^PVN→TH^RVLM projection, rAAV-EF1α-DIO-hM3Dq-mCherry-WPRE (AAV2/9, 5.27 × 10¹² v.g./ml, BrainVTA) was bilaterally injected into the PVN of Vglut2-Cre mice at a volume of 150 nl. rAAV-EF1a-DIO-mCherry-WPRE-pA (AAV2/9, 5.26 × 10¹² v.g./ml, BrainVTA) was used as a control for both sets of experiments.

For di-synaptic tracing of the PVN→RVLM→SPN pathway, DBH-Cre mice were unilaterally injected with 150 nl rAAV-EF1α-DIO-FLP-WPRE-hGH-pA (AAV2/1, 1.00 × 10¹³ v.g./ml, BrainVTA) into the PVN and 150 nl rAAV-hSyn-fDIO-H2B-EGFP-2A-Synaptophysin-mRuby-WPRE (AAV2/9, 5.00 × 10¹² v.g./ml, BrainVTA) into the ipsilateral RVLM.

For di-synaptic tracing of the RVLM→SPN→SG pathway, DBH-Cre mice were unilaterally injected with 150 nl rAAV-CAG-DIO-WGA-FLP-WPRE-hGH-pA (AAV2/9, 5.27 × 10¹² v.g./ml, BrainVTA) into the RVLM and 250 nl rAAV-hSyn-fDIO-H2B-EGFP-2A-Synaptophysin-mRuby-WPRE (AAV2/9, 5.00 × 10¹² v.g./ml, BrainVTA) into the ipsilateral spinal cord.

Histological procedures. For experiments involving MIR, at the end of reperfusion, mice were sequentially transcardially with ice-cold PBS (0.1 M, 20 ml) followed by 4% paraformaldehyde in 0.1 M PBS. The brain, spinal cord, DRG, SG, and heart were removed from each mouse and then post-fixed in 4% paraformaldehyde at 4°C overnight and dehydrated using 30% (w/v) sucrose in 0.1 M PBS at 4°C overnight. Samples were flash-frozen on dry ice. Coronal brain sections (40 µm thick) and sections of all other samples (10 µm thick) were cut on a cryostat (Leica CM1860) and used for immunofluorescence (see below).

For virus injection validation, the mice were deeply anesthetized with urethane (800 mg/kg) and α-chloralose (40 mg/kg). Coronal brain sections and sections of all other samples were then prepared as described above and mounted on slides. After three 5-min washes with 0.1 M PBS, the slides were cover slipped with Antifade Mounting Medium with DAPI (P0131, Beyotime, Shanghai, China). Fluorescence images were acquired with a Zeiss LSM900 confocal fluorescence microscope (Zeiss, USA, San Diego, CA).

Immunofluorescence. Tissue sections prepared as described above were incubated in 0.3% (v/v) Triton X-100 for 0.5 h, blocked with 10% fetal bovine serum for 1 h at room temperature, and incubated with primary antibodies at 4°C for 24 h. These antibodies included anti-c-Fos (1:500, rabbit, Abcam), anti-NeuN (1:200, mouse, CST), anti-Glutamate (1:400, rabbit, Sigma-Aldrich), anti-GABA (1:400, rabbit, Sigma-Aldrich), and anti-TH (1:1000, rabbit, Abcam). The sections were washed 3 times for 5 min each in PBS and incubated with the corresponding fluorophore-conjugated secondary antibodies for 2 h at room temperature. These secondary antibodies were goat anti-rabbit Alexa Fluor 647 (1:1000, Thermo Fisher), goat anti-rabbit Alexa Fluor 568 (1:1000, Thermo Fisher), goat anti-rabbit Alexa Fluor 488 (1:1000, Thermo Fisher), and goat anti-mouse Alexa Fluor 488 (1:1000, Thermo Fisher). Finally, sections were stained with Antifade Mounting Medium with DAPI. Fluorescence images were acquired with a Zeiss LSM900 confocal fluorescence microscope (Zeiss, USA, San Diego, CA).

Drug infusion. For chemogenetic inhibition/excitement of PVN or RVLM neurons, C57BL/6J, Vglut2-Cre, or DBH-Cre mice were injected i.p. with CNO (5 mg/kg, Sigma) 60 min before MIR.

For chemogenetic inhibition/excitement of PVN→RVLM projections, C57BL/6J or Vglut2-Cre mice were injected with CNO (0.2 µg/µl, 150 nl/side, Sigma) into the RVLM 20 min before MIR.

For β-adrenoceptor blocker experiments, metoprolol (12.5 mg/kg) was injected intracardially 10 minutes before MIR.

Statistics and reproducibility. Data were expressed as the mean ± standard error of measurement (s.e.m). The comparisons between groups were performed with paired t-tests, unpaired t-tests, or one-way analysis of variance (ANOVA) followed by Dunn’s post-hoc test, as appropriate. All statistical analyses were conducted with GraphPad Prism (Version 8.0, San Diego, CA, USA). Two-sided P values less than 0.05 were considered as statistically significant.

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A Paraventricular Nucleus–Rostral Ventrolateral Medulla Pathway Contributes to Myocardial Ischemia/Reperfusion Injury

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