The authors thank Kathy Gage for her editorial support. This study was supported by funds from the Department of Anesthesiology (Duke University Medical Center), American Heart Association grant (18CSA34080277), and National Institutes of Health (NIH) grants (NS099590, HL157354, NS117973, and NS127163).

All the procedures described here were conducted in accordance with the National Institutes of Health (NIH) guidelines for the care and use of animals in research, and the protocol was approved by the Duke Institute of Animal Care and Use Committee (IACUC). C57BL/6 male and female&#160;mice aged 8-10 weeks old were used for the present study.
1. Surgical preparation
<ol>
	<li>Weigh a mouse on a digital scale, and place it into a 4 in x 4 in x 7 in plexiglass anesthesia induction box.</li>
	<li>Adjust the anesthesia vaporizer to 5% isoflurane, the oxygen flow meter to 30, and the nitrogen flow meter to 70 (see Table of Materials).</li>
	<li>Take the animal out of the induction box, and lay it in a supine position on the surgical bench when its respiratory rate has decreased to 30-40 breaths per minute.</li>
	<li>Pull out the tongue with blunt forceps, and hold it using the non-dominant hand. Use the dominant&#160;hand to insert a laryngoscope (see Table of Materials) into the mouse&#39;s mouth and visualize the vocal cord.</li>
	<li>Use the non-dominant&#160;hand to insert a guide wire and 20 G intravenous catheter into the mouth. Gently insert the guide wire into the trachea.</li>
	<li>Push the catheter into the trachea until the wing part of the catheter is even with the nose tip. 
	NOTE: Do not intubate a mouse that is not fully anesthetized since this may injure the trachea and cause airway bleeding.</li>
	<li>Connect the intubated mouse to a small animal ventilator (see Table of Materials), and reduce the isoflurane to 1.5%.</li>
	<li>Input the mouse&#39;s body weight into the control panel of the ventilator to determine the tidal volume and respiratory rate.</li>
	<li>Keep the mouse in a supine position under a heat lamp, and maintain the rectal temperature at 37 &#176;C with a temperature controller.</li>
	<li>Shave the inguinal areas, disinfect the surgical area at least three times with iodine and alcohol&#160;(see Table of Materials),&#160;and cover the area with a sterile surgical drape.</li>
	<li>Apply eye ointment to both eyes and administer 5 mg/kg carprofen subcutaneously before surgery.</li>
	<li>Open the sterile instrument package for surgery. Make a 1 cm skin incision with surgical scissors to access the femoral arteries on both sides. Dissect and ligate the distal femoral artery with a single strand of 4-0 silk suture (see&#160;Table of Materials), and apply one drop of lidocaine.</li>
	<li>Apply an aneurysm clip at the proximal femoral artery&#160;and make a small cut on the artery distal to the clip. Insert a polyethylene 10 (PE-10, see Table of Materials) catheter into the left and the right femoral arteries. 
	NOTE: The left arterial line is used for blood pressure monitoring, while the right one is used for blood withdrawal and resuscitation mixture infusion.</li>
	<li>Inject 50 &#181;L of 1:10 heparinized saline into each arterial line to prevent clotting in the line.</li>
	<li>Turn the mouse to the prone position, and mount it on a stereotaxic head frame.</li>
	<li>Connect three needle electrodes (red, green, and black) to the left arm, left leg, and right arm for electrocardiogram (ECG, see Table of Materials) monitoring.</li>
	<li>Glue a flexible plastic fiber probe onto the intact temporal skull through a 0.5 cm skin incision for cerebral blood flow monitoring. This step is optional.</li>
	<li>Shave the top of the head,&#160;disinfect the surgical area at least three times with iodine and alcohol, and cover the area with a sterile surgical drapel.</li>
	<li>Cut a 2.5 cm midline skin incision, and use four small retractors to expose the entire skull surface for brain imaging.</li>
	<li>Place a monitoring imager (e.g., a laser speckle contrast imager, see Table of Materials) above the head. 
	NOTE: A few drops of saline can be added to the skull surface to facilitate laser speckle contrast imaging.</li>
</ol>
2. Induction of cardiac arrest
<ol>
	<li>Fill a 1 mL plastic syringe with 26 &#181;L of the resuscitation cocktail stock solution. 
	NOTE: Each milliliter of this solution contains 400 &#181;L of 1 mg/mL epinephrine, 500 &#181;L of 8.4% sodium bicarbonate, 50 &#181;L of 1,000 U/mL heparin, and 50 &#181;L of 0.9% sodium chloride (see Table of Materials).</li>
	<li>Wait until the body temperature reaches 37 &#176;C. Adjust the oxygen meter to 100% to oxygenate the blood for 2 min.</li>
	<li>Withdraw the oxygenated arterial blood up to 200 &#181;L via the right femoral artery into the prepared plastic syringe containing 26 &#181;L of resuscitation cocktail stock solution.</li>
	<li>Switch off the oxygen, and increase the nitrogen to 100% to induce anoxia. 
	NOTE: After approximately 45 s, the heart will fail to function, and the heart rate will rapidly decrease, indicating the onset of CA. After about 2 min of oxygen deprivation, the ECG monitoring will indicate an asystole, and there will be no measurable systemic blood pressure and negligible cerebral blood flow.</li>
	<li>Turn off the ventilator, isoflurane vaporizer, temperature controller, and nitrogen flowmeter. Adjust the oxygen to 100% in preparation for resuscitation.</li>
</ol>
3. Resuscitation procedure
<ol>
	<li>Turn the ventilator on at 8 min after CA onset.</li>
	<li>Immediately start to infuse the withdrawn oxygenated blood mixed with the resuscitation cocktail into the blood circulation via the right femoral artery in 1 min. 
	NOTE: The infusion leads to a gradual increase in the heart rate and the restoration of blood perfusion; eventually, the return of spontaneous circulation (ROSC) is achieved.</li>
</ol>
4. Post-CA recovery
<ol>
	<li>Place the mouse in the supine position after removing it from the stereotaxic frame, and remove&#160;the PE-10 catheters from the femoral arteries.</li>
	<li>Apply 0.25% bupivacaine to the skin incision, and suture the skin incisions using a 6-0 nylon suture (see Table of Materials). Apply antibiotic ointment to the surface of the skin incision.</li>
	<li>Disconnect the mouse ventilator when spontaneous respiration is restored.</li>
	<li>Transfer the mouse to a recovery chamber with a controlled temperature of 32 &#176;C.</li>
	<li>After 2 h of recovery, extubate the mouse, and return to the home cage. Inject 0.5 mL of normal saline subcutaneously to prevent dehydration.</li>
</ol>

A Mouse Model to Study Brain Physiology During Cardiac Arrest

<ol>
	<li>Smith, A., Masters, S., Ball, S., Finn, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+incidence+and+outcomes+of+out-of-hospital+cardiac+arrest+in+metropolitan+versus+rural+locations:+A+systematic+review+and+meta-analysis.">The incidence and outcomes of out-of-hospital cardiac arrest in metropolitan versus rural locations: A systematic review and meta-analysis.</a> Resuscitation. 185, 109655 (2022).</li><li>Amacher, S. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Predicting+neurological+outcome+in+adult+patients+with+cardiac+arrest:+systematic+review+and+meta-analysis+of+prediction+model+performance.">Predicting neurological outcome in adult patients with cardiac arrest: systematic review and meta-analysis of prediction model performance.</a> Critical Care. 26 (1), 382 (2022).</li><li>Matsuyama, T., Ohta, B., Kiyohara, K., Kitamura, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intra-arrest+partial+carbon+dioxide+level+and+favorable+neurological+outcome+after+out-of-hospital+cardiac+arrest:+A+nationwide+multicenter+observational+study+in+Japan+(the+JAAM-OHCA+registry).">Intra-arrest partial carbon dioxide level and favorable neurological outcome after out-of-hospital cardiac arrest: A nationwide multicenter observational study in Japan (the JAAM-OHCA registry).</a> European Heart Journal of Acute Cardiovascular Care. 12 (1), 14-21 (2023).</li><li>Takahagi, M., Sawano, H., Moriyama, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+neurological+outcome+of+extracorporeal+cardiopulmonary+resuscitation+for+out-of-hospital+cardiac+arrest+patients+with+nonshockable+rhythms:+A+single-center,+consecutive,+retrospective+observational+study.">Long-term neurological outcome of extracorporeal cardiopulmonary resuscitation for out-of-hospital cardiac arrest patients with nonshockable rhythms: A single-center, consecutive, retrospective observational study.</a> The Journal of Emergency Medicine. 63 (3), 367-375 (2022).</li><li>Mork, S. R., Botker, M. T., Christensen, S., Tang, M., Terkelsen, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survival+and+neurological+outcome+after+out-of-hospital+cardiac+arrest+treated+with+and+without+mechanical+circulatory+support.">Survival and neurological outcome after out-of-hospital cardiac arrest treated with and without mechanical circulatory support.</a> Resuscition Plus. 10, 100230 (2022).</li><li>Koenig, M. A., Kaplan, P. W., Thakor, N. 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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multimodal+monitoring+including+early+EEG+improves+stratification+of+brain+injury+severity+after+pediatric+cardiac+arrest.">Multimodal monitoring including early EEG improves stratification of brain injury severity after pediatric cardiac arrest.</a> Resuscitation. 167, 282-288 (2021).</li><li>Beekman, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bedside+monitoring+of+hypoxic+ischemic+brain+injury+using+low-field,+portable+brain+magnetic+resonance+imaging+after+cardiac+arrest.">Bedside monitoring of hypoxic ischemic brain injury using low-field, portable brain magnetic resonance imaging after cardiac arrest.</a> Resuscitation. 176, 150-158 (2022).</li><li>Sinha, N., Parnia, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monitoring+the+brain+after+cardiac+arrest:+A+new+era.">Monitoring the brain after cardiac arrest: A new era.</a> Current Neurology Neuroscience Report. 17 (8), 62 (2017).</li><li>Reis, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pathophysiology+and+the+monitoring+methods+for+cardiac+arrest+associated+brain+injury.">Pathophysiology and the monitoring methods for cardiac arrest associated brain injury.</a> International Journal of Molecular Sciences. 18 (1), 129 (2017).</li><li>Zhou, H., Lin, C., Liu, J., Wang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Continuous+monitoring+of+brain+perfusion+by+cerebral+oximetry+after+spontaneous+return+of+circulation+in+cardiac+arrest:+A+case+report.">Continuous monitoring of brain perfusion by cerebral oximetry after spontaneous return of circulation in cardiac arrest: A case report.</a> BMC Neurology. 22 (1), 365 (2022).</li><li>Takegawa, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+brain+monitoring+by+near-infrared+spectroscopy+predicts+neurological+outcome+after+cardiac+arrest+and+resuscitation+in+rats:+A+proof+of+concept+study+of+a+novel+prognostic+measure+after+cardiac+arrest.">Real-time brain monitoring by near-infrared spectroscopy predicts neurological outcome after cardiac arrest and resuscitation in rats: A proof of concept study of a novel prognostic measure after cardiac arrest.</a> Journal Clinical Medicine. 11 (1), 131 (2021).</li><li>Zhang, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Invasion+of+peripheral+immune+cells+into+brain+parenchyma+after+cardiac+arrest+and+resuscitation.">Invasion of peripheral immune cells into brain parenchyma after cardiac arrest and resuscitation.</a> Aging and Disease. 9 (3), 412-425 (2018).</li><li>Duan, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cervical+vagus+nerve+stimulation+improves+neurologic+outcome+after+cardiac+arrest+in+mice+by+attenuating+oxidative+stress+and+excessive+autophagy.">Cervical vagus nerve stimulation improves neurologic outcome after cardiac arrest in mice by attenuating oxidative stress and excessive autophagy.</a> Neuromodulation. 25 (3), 414-423 (2022).</li><li>Liu, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+modification+of+potassium+chloride+induced+cardiac+arrest+model+for+aged+mice.">Novel modification of potassium chloride induced cardiac arrest model for aged mice.</a> Aging and Disease. 9 (1), 31-39 (2018).</li><li>Shen, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aging+is+associated+with+impaired+activation+of+protein+homeostasis-related+pathways+after+cardiac+arrest+in+mice.">Aging is associated with impaired activation of protein homeostasis-related pathways after cardiac arrest in mice.</a> Journal of American Heart Association. 7 (17), e009634 (2018).</li><li>Wang, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Manganese+porphyrin+promotes+post+cardiac+arrest+recovery+in+mice+and+rats.">Manganese porphyrin promotes post cardiac arrest recovery in mice and rats.</a> Biology. 11 (7), 957 (2022).</li><li>Wang, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+evaluation+of+a+novel+mouse+model+of+asphyxial+cardiac+arrest+revealed+severely+impaired+lymphopoiesis+after+resuscitation.">Development and evaluation of a novel mouse model of asphyxial cardiac arrest revealed severely impaired lymphopoiesis after resuscitation.</a> Journal of American Heart Association. 10 (11), e019142 (2021).</li><li>Li, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+the+XBP1s/O-GlcNAcylation+pathway+improves+functional+outcome+after+cardiac+arrest+and+resuscitation+in+young+and+aged+mice.">Activation of the XBP1s/O-GlcNAcylation pathway improves functional outcome after cardiac arrest and resuscitation in young and aged mice.</a> Shock. 56 (5), 755-761 (2021).</li><li>Shen, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+the+ATF6+(activating+transcription+factor+6)+signaling+pathway+in+neurons+improves+outcome+after+cardiac+arrest+in+mice.">Activation of the ATF6 (activating transcription factor 6) signaling pathway in neurons improves outcome after cardiac arrest in mice.</a> Journal American Heart Association. 10 (12), e020216 (2021).</li><li>Jiang, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MCC950,+a+selective+NLPR3+inflammasome+inhibitor,+improves+neurologic+function+and+survival+after+cardiac+arrest+and+resuscitation.">MCC950, a selective NLPR3 inflammasome inhibitor, improves neurologic function and survival after cardiac arrest and resuscitation.</a> Journal of Neuroinflammation. 17 (1), 256 (2020).</li><li>Zhao, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+arrest+and+resuscitation+activates+the+hypothalamic-pituitary-adrenal+axis+and+results+in+severe+immunosuppression.">Cardiac arrest and resuscitation activates the hypothalamic-pituitary-adrenal axis and results in severe immunosuppression.</a> Journal of Cerebral Blood Flow &amp; Metabolism. 41 (5), 1091-1102 (2021).</li></ol>

The authors have no conflicts of interest.

In experimental CA studies, asphyxia, potassium chloride injections, or electrical current-derived ventricular fibrillation have been used to induce CA16,17,18,19,20,21,22,23. Normally, CPR is required for resuscitation in these CA models, especially in mic...

Cardiac arrest (CA) remains a global public health crisis1. More than 356,000 out-of-hospital and 290,000 in-hospital CA cases are reported annually in the US alone, and most CA victims are over 60 years old. Notably, post-CA neurologic impairments are common among survivors, and these represent a major challenge for CA management2,3,4,5. To understand post-CA brain pathologic changes and their effects on neurologic outcomes, various neurophysiologic monitoring and brain tissue monitoring techniques have been applied in patients6,7,8,9,10,11,12. Using near-infrared spectroscopy, real-time brain monitoring has also been performed in CA rats to predict neurologic outcomes13.
However, in murine CA models, such an imaging approach has been complicated by the need for chest compressions to restore spontaneous circulation, which always entails substantial physical motion and, thus, hinders delicate imaging procedures. Moreover, CA models are normally performed with mice in a supine position, whereas the mice must be turned to the prone position for many brain imaging modalities. Thus, a mouse model with minimal body movement during the surgery is required in many cases in order to perform real-time imaging/monitoring of the brain during the whole CA procedure, spanning from pre-CA to post-resuscitation.
Previously, Zhang et al. reported a mouse CA model that could be useful for brain imaging14. In their model, CA was induced by bolus injections of vecuronium and esmolol followed by the cessation of mechanical ventilation. They showed that after 5 min of CA, resuscitation could be achieved by infusing a resuscitation mixture. Notably, however, circulatory arrest in their model occurred only about 10 s after the esmolol injection. Thus, this model does not recapitulate the progression of asphyxia-induced CA in patients, including hypercapnia and tissue hypoxia during the prearrest period.
The overall goal of the current surgical procedure is to model clinical asphyxia CA in mice followed by resuscitation without chest compressions. This CA model, therefore, allows the use of complex imaging techniques to study brain physiology in mice15.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >Adrenalin</td>
			<td >Par Pharmaceutical</td>
			<td >NDC 42023-159-01</td>
			<td ></td>
		</tr>
		<tr >
			<td >Alcohol swabs</td>
			<td >BD</td>
			<td >326895</td>
			<td></td>
		</tr>
		<tr >
			<td >Animal Bio Amp</td>
			<td >ADInstruments</td>
			<td >FE232</td>
			<td></td>
		</tr>
		<tr >
			<td >BP transducer</td>
			<td >ADInstruments</td>
			<td >MLT0699</td>
			<td></td>
		</tr>
		<tr >
			<td >Bridge Amp</td>
			<td >ADInstruments</td>
			<td >FE117</td>
			<td></td>
		</tr>
		<tr >
			<td >Heparin sodium injection, USP</td>
			<td >Fresenius Kabi</td>
			<td >NDC 63323-540-05</td>
			<td></td>
		</tr>
		<tr >
			<td >Isoflurane</td>
			<td >Covetrus</td>
			<td >NDC 11695-6777-2</td>
			<td></td>
		</tr>
		<tr >
			<td >Laser Doppler perfusion monitor</td>
			<td >Moor Instruments</td>
			<td >moorVMS-LDF1</td>
			<td></td>
		</tr>
		<tr >
			<td >Laser speckle imaging system</td>
			<td >RWD</td>
			<td >RFLSI III</td>
			<td></td>
		</tr>
		<tr >
			<td >Lubricant eye ointment</td>
			<td >Bausch + Lomb</td>
			<td >339081</td>
			<td></td>
		</tr>
		<tr >
			<td >Micro clip</td>
			<td >Roboz</td>
			<td >RS-5431</td>
			<td></td>
		</tr>
		<tr >
			<td >Mouse rectal probe</td>
			<td >Physitemp</td>
			<td >RET-3</td>
			<td></td>
		</tr>
		<tr >
			<td >Needle electrode</td>
			<td >ADInstruments</td>
			<td >MLA1213</td>
			<td>29 Ga, 1.5 mm socket</td>
		</tr>
		<tr >
			<td >Nitrogen</td>
			<td >Airgas</td>
			<td >UN1066</td>
			<td></td>
		</tr>
		<tr >
			<td >Optic plastic fibre</td>
			<td >Moor Instruments</td>
			<td >POF500</td>
			<td></td>
		</tr>
		<tr >
			<td >Otoscope</td>
			<td >Welchallyn</td>
			<td >728</td>
			<td>2.5 mm Speculum</td>
		</tr>
		<tr >
			<td >Oxygen</td>
			<td >Airgas</td>
			<td >UN1072</td>
			<td></td>
		</tr>
		<tr >
			<td >PE-10 tubing</td>
			<td >BD</td>
			<td >427401</td>
			<td>Polyethylene tubing</td>
		</tr>
		<tr >
			<td >Povidone-iodine</td>
			<td >CVS</td>
			<td >955338</td>
			<td></td>
		</tr>
		<tr >
			<td >PowerLab 8/35</td>
			<td >ADInstruments</td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Rimadyl (carprofen)</td>
			<td >Zoetis</td>
			<td >6100701</td>
			<td>Injectable 50 mg/ml</td>
		</tr>
		<tr >
			<td >Small animal ventilator</td>
			<td >Kent Scientific</td>
			<td >RoVent Jr.</td>
			<td></td>
		</tr>
		<tr >
			<td >Temperature controller</td>
			<td >Physitemp</td>
			<td >TCAT-2DF</td>
			<td></td>
		</tr>
		<tr >
			<td >Triple antibioric &#38; pain relief</td>
			<td >CVS</td>
			<td >NDC 59770-823-56</td>
			<td></td>
		</tr>
		<tr >
			<td >Vaporizer</td>
			<td >RWD</td>
			<td >R583S</td>
			<td></td>
		</tr>
		<tr >
			<td >0.25% bupivacaine</td>
			<td >Hospira</td>
			<td >NDC 0409-1159-18</td>
			<td></td>
		</tr>
		<tr >
			<td >0.9% sodium chroride</td>
			<td >ICU Medical</td>
			<td >NDC 0990-7983-03</td>
			<td></td>
		</tr>
		<tr >
			<td >1 mL plastic syringe</td>
			<td >BD</td>
			<td >309659</td>
			<td></td>
		</tr>
		<tr >
			<td >4-0 silk suture</td>
			<td >Look</td>
			<td >SP116</td>
			<td>Black braided silk</td>
		</tr>
		<tr >
			<td >6-0 nylon suture</td>
			<td >Ethilon</td>
			<td >1698G</td>
			<td></td>
		</tr>
		<tr >
			<td >8.4% sodium bicarbonate Inj., USP</td>
			<td >Hospira</td>
			<td >NDC 0409-6625-02</td>
			<td></td>
		</tr>
		<tr >
			<td >20 G IV catheter</td>
			<td >BD</td>
			<td >381534</td>
			<td>20GA 1.6 IN</td>
		</tr>
		<tr >
			<td >30 G PrecisionGlide needle</td>
			<td >BD</td>
			<td >305106</td>
			<td>30 G X 1/2</td>
		</tr>
	</tbody>
</table>

mouse cardiac arrest model for brain imaging and brain physiology monitoring during ischemia and resuscitation

Most cardiac arrest (CA) survivors experience varying degrees of neurologic deficits. To understand the mechanisms that underpin CA-induced brain injury and, subsequently, develop effective treatments, experimental CA research is essential. To this end, a few mouse CA models have been established. In most of these models, the mice are placed in the supine position in order to perform chest compression for cardiopulmonary resuscitation (CPR). However, this resuscitation procedure makes the real-time imaging/monitoring of brain physiology during CA and resuscitation challenging. To obtain such critical knowledge, the present protocol presents a mouse asphyxia CA model that does not require the chest compression CPR step. This model allows for the study of dynamic changes in blood flow, vascular structure, electrical potentials, and brain tissue oxygen from the pre-CA baseline to early post-CA reperfusion. Importantly, this model applies to aged mice. Thus, this mouse CA model is expected to be a critical tool for deciphering the impact of CA on brain physiology.

To induce CA, the mouse was anesthetized with 1.5% isoflurane and ventilated with 100% nitrogen. This condition led to severe bradycardia in 45 s (Figure 1). Following 2 min of anoxia, the heart rate dramatically reduced (Figure 2), the blood pressure decreased below 20 mmHg, and the cerebral blood flow ceased completely (Figure 1).&#160;As the isoflurane was turned off, the body temperature was no longer managed and slowly dropped ...

Watch this Scientific Journal Video about A Unique Mouse Model of Asphyxia-Induced Cardiac Arrest at JoVE.com

Author Spotlight: A Unique Mouse Model of Asphyxia-Induced Cardiac Arrest 

This protocol demonstrates a unique mouse model of asphyxia cardiac arrest that does not require chest compression for resuscitation. This model is useful for monitoring and imaging the dynamics of brain physiology during cardiac arrest and resuscitation.

Mouse Cardiac Arrest Model for Brain Imaging and Brain Physiology Monitoring During Ischemia and Resuscitation

Most cardiac arrest (CA) survivors experience varying degrees of neurologic deficits. To understand the mechanisms that underpin CA-induced brain injury ...

mouse-cardiac-arrest-model-for-brain-imaging-brain-physiology

Research

JoVE Journal

Neuroscience

1.6K Views.  Duke University Medical Center. This protocol demonstrates a unique mouse model of asphyxia cardiac arrest that does not require chest compression for resuscitation. This model is useful for monitoring and imaging the dynamics of brain physiology during cardiac arrest and resuscitation.

Video: Author Spotlight: A Unique Mouse Model of Asphyxia-Induced Cardiac Arrest 

State-Dependency Effects on TMS:  A Look at Motive Phosphene Behavior

Transcranial magnetic stimulation (TMS) is a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate cortical excitability (either increasing or decreasing it) via the application of localized magnetic field pulses.1,2 Within the field of TMS, the term state dependency refers to the initial, baseline condition of the particular neural region targeted for stimulation. As can be inferred, the effects of TMS can (and do) vary according to this primary susceptibility and responsiveness of the targeted cortical area.3,4,5

In this experiment, we will examine this concept of state dependency through the elicitation and subjective experience of motive phosphenes. Phosphenes are visually perceived flashes of small lights triggered by electromagnetic pulses to the visual cortex. These small lights can assume varied characteristics depending upon which type of visual cortex is being stimulated. In this particular study, we will be targeting motive phosphenes as elicited through the stimulation of V1/V2 and the V5/MT+ complex visual regions.6

In this article, we examine the effects of visually relevant state dependency on TMS induced motive phosphenic presentations.

Transcranial magnetic stimulation (TMS) is a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate ...

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Whole-mount Imaging of Mouse Embryo Sensory Axon Projections

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we describe a method to label in situ a subset of dorsal root ganglion (DRG) axon projections to assess their phenotypic characteristics using several genetically manipulated mouse lines. The TrkA-positive neurons are nociceptor neurons, dedicated to the transmission of pain signals. We utilize a TrkAtaulacZ mouse line to label the trajectories of all TrkA-positive peripheral axons in the intact mouse embryo. We further breed the TrkAtaulacZ line onto a Bax null background, which essentially abolishes neuronal apoptosis, in order to assess growth-related questions independently of possible effects of genetic manipulations on neuronal survival. Subsequently, genetically modified mice of interest are bred with the TrkAtaulacZ/Bax null line and are then ready for study using the techniques described herein. This presentation includes detailed information on mouse breeding plans, genotyping at the time of dissection, tissue preparation, staining and clearing to allow for visualization of full-length axonal trajectories in whole-mount preparation.

We present here an optimized protocol to genotype, stain and prepare fetal mice for the imaging of peripheral nociceptor axon projections in the whole animal, as an effective method to assess sensory axon growth phenotypes in developing genetically engineered mice.

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we ...

Chronic Implantation of Whole-cortical Electrocorticographic Array in the Common Marmoset

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent development of thin, flexible ECoG electrodes has enabled conduction of stable recordings of large-scale cortical activity. We have developed a whole-cortical ECoG array for the common marmoset. The array continuously covers almost the entire lateral surface of cortical hemisphere, from the occipital pole to the temporal and frontal poles, and it captures whole-cortical neural activity in one shot. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain. Marmosets have two advantages regarding ECoG recordings, one being the homologous organization of anatomical structures in humans and macaques, including frontal, parietal, and temporal complexes. The other advantage is that the marmoset brain is lissencephalic and contains a large number of complexes, which are more difficult to access in macaques with ECoG, that are exposed to the brain surface.These features allow direct access to most cortical areas beneath the surface of the brain. This system provides an opportunity to investigate global cortical information processing with high resolutions at a sub-millisecond order in time and millimeter order in space.

We have developed a whole-cortical electrocorticographic array for the common marmoset that continuously covers almost the entire lateral surface of cortex, from the occipital pole to the temporal and frontal poles. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain.

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...

Continuous Video Electroencephalogram during Hypoxia-Ischemia in Neonatal Mice

Hypoxia ischemia is the most common cause of neonatal seizures. Animal models are crucial for understanding the mechanisms and physiology underlying neonatal seizures and hypoxia ischemia. This manuscript describes a method for continuous video electroencephalogram (EEG) monitoring in neonatal mice to detect seizures and analyze EEG background during hypoxia ischemia. Use of video and EEG in conjunction allows description of seizure semiology and confirmation of seizures. This method also allows analysis of power spectrograms and EEG background pattern trends over the experimental time period. In this hypoxia ischemia model, the method allows EEG recording prior to injury to obtain a normative baseline and during injury and recovery. Total monitoring time is limited by the inability to separate pups from the mother for longer than four hours. Although, we have used a model of hypoxic-ischemic seizures in this manuscript, this method for neonatal video EEG monitoring could be applied to diverse disease and seizure models in rodents.

This manuscript describes a method for continuous video EEG recordings using multiple depth electrodes in neonatal mice undergoing hypoxia-ischemia.

Hypoxia ischemia is the most common cause of neonatal seizures. Animal models are crucial for understanding the mechanisms and physiology underlying ...