Name: experimental approach to examine leptin signaling in the carotid bodies and its effects on control of breathing
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All experimental protocols have been approved by the Institutional Animal Care and Use Committee (MO18M211).
1. Leptin Infusion
NOTE: In order to examine the effect of leptin on breathing, we infused leptin subcutaneously in lean C57BL/6J mice by an osmotic pump to raise circulating leptin levels to those observed in obese mice.
<ol>
	<li>Osmotic pump preparation
	<ol>
		<li>Weigh the empty pump to check the net weight of the solution loaded.</li>
		<li>Add Leptin (5 mg/mL) to the osmotic pump (1 &#956;L/h for 3 days). Fill the pump with a small syringe (1 mL). After filling, close the pump with the provided blunt tipped 27 gauge filling tube. 
		NOTE: The syringe and the attached tube should be free of air bubbles.</li>
		<li>After filling, weigh the pump again to check the net weight of the solution.</li>
		<li>Insert the leptin pump subcutaneously in the interscapular area. 
		NOTE: If you want to start the infusion immediately, incubate the pre-filled pump in the sterile saline at 37 &#176;C for at least 4 to 6 h (preferably overnight).</li>
	</ol>
	</li>
</ol>
2. Hypoxic Ventilatory Response (HVR)
NOTE: Thermoneutral conditions eliminate stressful factors imposed by cool ambient temperature and significantly modify metabolism in mice. Therefore, all respiratory measurements should be performed at the thermoneutral conditions (t = 30 &#176;C) using a neonatal incubator12 (Figure 1A). The whole body plethysmography chamber (WBP) has been used for all measurements. All animals should be acclimated to the barometric plethysmography chamber and to a sham neck collar for subsequent pulse oximetry recording for 3-5 days prior to the HVR measurements. HVR is recorded between 10 AM and 5 PM. HVR is suppressed during sleep, therefore it can be measured separately during sleep and quiet wakefulness13. To ensure the specific sleep-wake stage of the animal during the HVR measurement, EEG/EMG electrodes should be implanted as an EEG/EMG headmount as previously described14. Animals should be allowed to recover for at least 72 h after the headmount. Sleep staging should be performed in 5 s epochs. NREM sleep is recognized by the slow wave EEG activity occupying greater than 50% of the epoch. Quiet wakefulness is manifested by the alpha EEG activity in the absence of movements. REM sleep is identified by the predominant alpha and theta EEG activity in the presence of reduced muscle tone on EMG. Typically, REM sleep is not observed during the HVR gas challenges.
<ol>
	<li>The HVR recording protocol
	<ol>
		<li>Use the WBP chamber of the following size: internal diameter of 80 mm, height of 50.5 mm, and volume of 0.4 L. The WBP chamber consists of two chambers, sealed and reference chambers, and a circular platform to place the mouse14 (Figure 1B). Inflow and outflow inside the chamber are controlled by positive and negative pressure sources maintaining the atmospheric pressure. This control creates a steady bias flow in the chamber and prevents CO2 retention. For more details about the WBP set up, see Hernandez and colleagues15.</li>
		<li>Measure the temperature inside the chamber and the relative humidity in the room prior to placing the mouse in the WBP.</li>
		<li>Measure the body weight and rectal temperature.</li>
		<li>Put the oximeter collar around the neck of the mouse (the area should be previously shaved).</li>
		<li>Place the mouse inside the WBP and ensure that the chamber is completely sealed in order to avoid air leakages.</li>
		<li>Wait for approximately 30 min until the mouse is quiet and the chamber is at a constant volume.</li>
		<li>Normoxia: after 30 min, if the mouse is quiet, start recording the respiratory signals and peripheral oxygen saturation (SpO2) at normoxia phase (21% FiO2 at 1 atm of pressure) for 20 min, using LabChart 7 Pro (version 7.2).</li>
		<li>Hypoxia: after 20 min of quiet normoxia, start recording the first cycle of acute hypoxia and SpO2. For hypoxia phase, expose the animals to a constant mixed gas flow composed by 10% O2 and 3% CO2 for 5 min.
		<ol>
			<li>During the first 30 sec of hypoxia, the mixed gas flows through the chamber via 2 small side ports at the base allowing the FiO2 to drop from 21% (at 1 atm of pressure) to 10%.</li>
			<li>After the initial 30 s, close one of the small side ports of the WBP chamber and keep recording at the constant hypoxia at 10% FiO2 for 5 min. 
			NOTE: The mixture of 10% O2 and 3% CO2 at hypoxia is used in order to maintain eucapnea11.</li>
		</ol>
		</li>
		<li>After the 5 min of hypoxia, expose the mouse to room air again (21% FiO2 at 1 atm of pressure) by switching the inflow source.</li>
		<li>Wait for at least 30 min until the next normoxia recording in order to recover from the previous hypoxic exposure to avoid the ventilatory roll-off phenomenon (i.e., central suppression of breathing during hypoxia16). 
		NOTE: Repeat normoxia/hypoxia cycles three times in each mouse to assure reproducibility of the measurements. In our experience, additional (greater than 3 times on a given day) hypoxic exposures should be avoided, because of the ventilatory adaptation (the roll-off phenomenon).</li>
		<li>At the end of the experiment, calibrate the WBP chamber (with the mouse is still inside) by injecting 1 mL of room air 3 times with a syringe plugged in one of the small side ports at the base of the chamber.</li>
		<li>Measure the temperature in the chamber again with the animal inside it.</li>
		<li>Open the chamber and measure the rectal temperature of the mouse before placing it back to its home cage.</li>
	</ol>
	</li>
	<li>HVR calculation
	<ol>
		<li>Digitize all signals for HVR calculation at 1,000 Hz (sampling frequency per channel). 
		NOTE: WBP tidal volume analysis is performed in Lab Chart 7 based on Drorbaugh and Fenn equation17. Required variables comprise mouse rectal temperature and the chamber temperature (before and after HVR measurement), relative humidity, and the chamber gas constant. This constant is resultant from WBP pressure deflection after the 1 mL injections of air in the calibration phase. For more details, see Hernandez and colleagues15.</li>
		<li>After calculating the Drorbaugh and Fenn equation, multiply the tidal volume (Vt) chamber channel by the constant.</li>
		<li>Selection of recordings for the analysis
		<ol>
			<li>Normoxia: select only sections of steady breathing with constant tidal volume. Avoid sections in close proximity to mouse&#8217;s movements.</li>
			<li>Hypoxia: discard first 30 s when O2 levels are declining from 21% to 10%, select sections from 30 s to 2 min of 10% O2 exposure (a 90 s interval). Within this interval, select only sections with steady breathing with constant tidal volume. Avoid sections in close proximity to mouse&#8217;s movements. The analysis is reliable if at least 10 s of hypoxia is selected in each cycle. 
			NOTE: The peripheral component of chemoreflex governed by CB predominates during the first 1-2 min of exposure16,18,19. During the second phase, between 2 min and 5 min of hypoxic exposure, both peripheral and central component play a role. Finally, the third phase, &#62; 5 min, is characterized by hypoventilation (the roll-off phenomenon) governed predominantly by central chemoreceptors. Our experience shows that mice are often awake during hypoxic exposition because of the manual switches in the airflow source.</li>
		</ol>
		</li>
		<li>After the selection, calculate the mean minute ventilation (VE) at normoxic and hypoxic conditions using the formula VE = respiratory rate X tidal volume.</li>
		<li>Calculate the mean SpO2 at normoxia and hypoxia conditions.</li>
		<li>Calculate the HVR manually using the formula HVR = (VE (10% O2) - VE (21%O2)) / (SpO2 (10% O2) &#8211; SpO2 (21% O2)). 
		NOTE: In C57BL/6J mice, the osmotic pump for leptin infusion may impair the accurate SpO2 signal detection by the neck collar. In this case, HVR is calculated based on the FiO2 values at normoxic and hypoxic conditions as HVR = &#916;VE / &#916;FiO211.</li>
	</ol>
	</li>
</ol>
3. Carotid Body Denervation or Carotid Sinus Nerve Dissection (CSND)
NOTE: We performed combined surgical and chemical denervation one week apart, because surgical denervation alone does not abolish the hypoxic chemoreflex.
<ol>
	<li>Surgical preparation
	<ol>
		<li>Perform all the procedures on adult male C57BL/6J mice. Sterilize all surgical instruments. Use sterile surgical gloves, syringes and cotton tipped applicators.</li>
		<li>Anesthetize male C57BL/6J mice with 1-2% isoflurane using a nose cone and place a warm blanket to prevent hypothermia. Isoflurane is carefully titrated to maintain respiratory rate at 1 Hz (60 breaths/min). The adequacy of anesthesia prior to starting surgeries will be assessed by the breathing frequency, the absence of movements and audible noises, the absence of response to tactile stimuli and the forelimb or hindlimb pedal withdrawal reflex.</li>
		<li>Administer Buprenorphine (0.05 mg/kg) intraperitoneally to prevent pain discomfort. Remove the hair at ventral region of the neck and disinfect the area with betadine and alcohol.</li>
		<li>To prevent corneal desiccation, lubricate the mice&#8217;s eyes with sterile ophthalmic ointment during anesthesia.</li>
	</ol>
	</li>
	<li>CSND procedures 
	Stage 1: Surgical denervation
	<ol>
		<li>After midline incision, expose the bifurcation of the common carotid arteries by removing connective tissues and adipose tissue.</li>
		<li>Identify the hypoglossal nerve, which is usually very prominent, and then lift it up to expose the glossopharyngeal nerve immediately underneath. Dissect carotid sinus nerves bilaterally using micro spring scissors.</li>
		<li>Close the incision with 6-0 silk suture.</li>
		<li>Administer 1 mL of normal saline subcutaneously to prevent dehydration.</li>
		<li>House the mice in a recovery chamber and monitor their behavior every 15 min for initial 1 h until the mice regain consciousness to maintain sternal recumbency. Return the mice to their home cages after they are fully recovered. Keep monitoring the mice twice per day for next 3 days. Give additional buprenorphine if mice show any signs of pain (e.g., reduced appetite, restlessness). 
		Stage 2: Chemical denervation</li>
		<li>One week after the surgery, paint the carotid artery with a solution of 2% phenol diluted in ethanol at the points of branching from the glossopharyngeal nerve to the cranial pole of the CB. The same post-operative care should be provided after the chemical denervation as described above in the surgical denervation section. 
		NOTE: For a sham surgery group, the same procedures are performed except the carotid sinus nerve dissection.</li>
		<li>Let the animals recover for 5-7 days before HVR measurements.</li>
	</ol>
	</li>
</ol>
4. Expression of LepRb in the CB using an adenoviral vector (Ad-LepRb) vs control (Ad-LacZ)
<ol>
	<li>Ad-LepRb and Ad-LacZ suspension
	<ol>
		<li>Transfect the mice with adenovirus (Ad-LepRb-GFP, 2-5x1010 pfu/mL for overexpression, Ad-Lacz, 1 x 1010 pfu/mL for control) to the CB area.</li>
		<li>Thaw the Matrigel matrix by submerging the vial in ice in a 4 &#176;C refrigerator overnight.</li>
		<li>Gently suspend adenovirus in liquid Matrigel matrix at 1:5 (1 &#956;L of Matrigel matrix and 4 &#956;L of adenovirus applied bilaterally).</li>
		<li>Always keep viral suspension on ice until it is applied in the CB.</li>
	</ol>
	</li>
	<li>Surgical preparation
	<ol>
		<li>Use the same surgical instruments as the CSND protocol.</li>
		<li>Anesthetize LepRb-deficient db/db mice with 2-2.5% isoflurane using a nose cone and place the animals on a warm blanket to prevent hypothermia. Isoflurane is carefully titrated to maintain respiratory rate at 1 Hz (60 breaths/min). The adequacy of anesthesia will be assessed by the breathing frequency, the absence of movements and audible noises, the absence of response to tactile stimuli and the forelimb or hindlimb pedal withdrawal reflex.</li>
		<li>Administer buprenorphine (0.05 mg/kg) intraperitoneally.</li>
		<li>Remove the hair at ventral region of the neck and disinfect the area with betadine and alcohol as in the CSND protocol.</li>
		<li>To prevent corneal desiccation, lubricate the mice&#8217;s eyes with sterile ophthalmic ointment during anesthesia.</li>
	</ol>
	</li>
	<li>Adenovirus treatment of the CB
	<ol>
		<li>Expose CBs as described in the CSND protocol and apply 5 &#956;L of the viral suspension to the CB area bilaterally.</li>
		<li>Wait 2-3 min until the liquid Matrigel matrix becomes a gel. After confirming that the viral suspension was congealed, close the incision with 6.0 silk suture.</li>
	</ol>
	</li>
	<li>After surgery
	<ol>
		<li>House the mice in a recovery chamber and monitor their behavior every 15 min for initial 1 h until the mice regain consciousness to maintain sternal recumbency. Return the mice to their home cages after they are fully recovered. Keep monitoring the mice twice per day for next 3 days. Give additional buprenorphine if mice show any signs of pain (e.g., reduced appetite, restlessness).</li>
		<li>Administer Buprenorphine (0.05 mg/kg) intraperitoneally as needed to prevent pain discomfort during the post-operative period.</li>
		<li>Measure HVR 9 days after the adenovirus transfection.</li>
	</ol>
	</li>
</ol>

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The authors have no conflicting interests or disclosures.

The main focus of our study was to examine respiratory effects of leptin signaling in the CB. Several protocols have been developed to assess the role of leptin in a mechanistic fashion. First, specific contribution of CB to the HVR was analyzed by careful quantification of the HVR during the first 2 min of hypoxic exposure. Second, the relevance of CB in leptin-mediated up-regulation of control of breathing was examined by two complementary approaches. In lean wild-type mice with low leptin levels, the HVR was measured at baseline and after continuous infusion of leptin; the experiment was repeated after CB denervation. In LepRb-deficient db/db mice, the HVR was measured at baseline and after LepRb expression in the CB.
Several protocols have been used to measure HVR in rodents, exposing the animals to hypoxic gases. We have developed an HVR protocol to examine the role of CB in the peripheral chemoreflex and the effects of leptin in CB on the control of breathing. CB chemoreflex has a relatively short time domain. The first 1-2 min of hypoxia are characterized by an acute augmentation of ventilatory response followed by a slow return to baseline ventilation after 2 min of the carotid sinus nerve stimulation16,18,19. Thus, in our HVR protocol, the analyses are conducted within a 90 s interval between the first 30 s of hypoxia and 2 min of 10% O2 exposure, corresponding to the domination of peripheral chemoreflex governed by CB. This short-time analysis avoids the hypoxic ventilatory depression in the animals. In our HVR protocol, we also used a fixed 3% CO2 tension in hypoxia in order to circumvent the hypocapnia induced by hyperventilation during acute hypoxic exposure11. Finally, we have developed the HVR protocol under constant thermoneutral conditions, keeping the mice at a temperature of around 30 &#176;C. Our experience shows that short exposures to hypoxia can decrease rectal temperature in mice if exposure occurs at room temperature11, while hypoxia at thermoneutrality does not induce significant metabolic changes12.
CSND in mice is technically challenging, because of the small size of the animals and their CBs. We have developed a consistently successful approach with nearly 100% survival rate by strict adherence to our protocol. Controlled conditions in our protocol includes thermoneutral environment, carefully controlled anesthesia and standard sterile microsurgical techniques with visualization of the glossopharyngeal nerve as vigilant post-operative management with pain control. Our experience shows that surgical denervation alone does not abolish the hypoxic chemoreflex. The second step, chemical denervation, is also followed by careful post-operative management to improve survival.
Our most innovative technique is selective gene over-expression in the CB area. This approach has not been implemented previously, because of the small size of the CBs and the lack of specific drivers allowing to express a gene of interest in a particular cell type. In fact, type I CB cells are very similar to sympathetic neurons or cells of the adrenal medulla, whereas type II cells are similar to astrocytes20,21. We took advantage of db/db mice, which lack the LepRb gene, our ability to apply adenoviral suspension almost exclusively to the CB area, and properties of the Matrigel matrix, which rapidly solidifies at 37 &#176;C. Our novel approach can be used in the future to study a role of any gene expressed in CB using mice with the whole body tyrosine hydroxylase specific (type I cells) or GFAP specific (type II cells) knockouts.
Our protocols have several limitations. First, we used 3% CO2 to determine the HVR and the question remains open if the fraction of the HVR can actually be attributed to the hypercapnic response. In order to address this limitation, investigators can simultaneously measure responses to 3% CO2 balanced in hyperoxic gas, which would turn the CB off. Second, HVR may not be completely eliminated by CSND22. This phenomenon can be attributed to neuroplasticity, which is particularly prominent in mice. Therefore, it is important to study HVR as soon as possible after CSND and always use sham surgery control. Third, our CB gene expression approach lacked cell type and organ specificity. Molecular techniques with the future use of more selective promoters may help to counter this limitation.
In conclusion, despite described above limitations, our protocols in combination allow to study the role of specific CB genes in physiological responses to hypoxia.

An adipocyte produced hormone leptin acts in the hypothalamus to suppress food intake and increase metabolic rate. Studies performed in our laboratory1,2 and by other investigators3,4 showed that leptin increases the hypercapnic ventilatory response (HVR) preventing obesity hypoventilation in leptin deficient obesity. However, a majority of obese individuals have high plasma leptin levels and demonstrate resistance to the metabolic and respiratory effects of the hormone5,6,7,8. Resistance to leptin is multifactorial, but limited permeability of the blood-brain barrier (BBB) to leptin plays a major role. We propose that leptin acts below BBB in a key organ of peripheral hypoxic sensitivity, the carotid bodies (CB), to defend breathing in obese individuals. CBs express the long functional isoform of leptin receptor, LepRb, but the role of CB in respiratory effects of leptin has not been sufficiently elucidated9,10.
The goal of our method was to examine the effect of leptin signaling in the CB on HVR. Our rationale was to perform (a) loss of function experiments infusing leptin in mice with intact carotid bodies and denervated carotid bodies followed by HVR measurements; (b) gain of function experiments in db/db mice lacking LepRb, in which we measured the HVR at baseline and after expression of LepRb exclusively in CB. The advantage of our techniques was that we performed all our experiments in unrestrained unanesthetized mice during sleep and wakefulness. Previous investigators either performed their experiments under anesthesia9 or did not measure effects of leptin during sleep10. In addition, our study is the first to utilize a unique gain of function approach with selective LepRb expression in CB described above.
In the broad context, our approach can be generalized to other receptors expressed in CB and their role in hypoxic sensitivity. Investigators can infuse a ligand to a receptor of interest and measure the HVR at baseline and after CB denervation. As a complementary approach, a receptor of interest can be overexpressed in CB and HVR measurements can be performed before and after overexpression using our technology described in this manuscript.

<table><tbody><tr><td>1 mL Insulin Syringes</td><td>BD Biosciences</td><td>309311</td><td></td></tr><tr><td>1x PBS (pH 7.4)</td><td>Gibco</td><td>10010-023</td><td>500 ml</td></tr><tr><td>&lt;em&gt;Ad-Lacz&lt;/em&gt;</td><td>Dr. Christopher Rhodes (University of Chicago)</td><td></td><td>1 x 10&lt;sup&gt;10&lt;/sup&gt; pfu/mL</td></tr><tr><td>&lt;em&gt;Ad-LepRb-GFP&lt;/em&gt;</td><td>Vector Biolabs</td><td>ADV-263380</td><td>2-5 x 10&lt;sup&gt;10&lt;/sup&gt; pfu/mL</td></tr><tr><td>Anesthetic cart</td><td>Atlantic Biomedical</td><td></td><td></td></tr><tr><td>Betadine</td><td>Purdue Products Ltd.</td><td>12496-0757-5</td><td></td></tr><tr><td>Buprenorphine (Buprenex)</td><td>Reckitt Benckiser Healthcare Ltd.</td><td>12496-0757-5</td><td>0.3 mg/mL</td></tr><tr><td>C57Bl/6J</td><td>Jackson laboratory</td><td>000664</td><td>Mice Strain</td></tr><tr><td>Cotton Gauze Sponges</td><td>Fisherbrand</td><td>22-362-178</td><td></td></tr><tr><td>db/db</td><td>Jackson laboratory</td><td>000697</td><td>Mice Strain</td></tr><tr><td>Ethanol</td><td>Pharmco-AAPER</td><td>111000200</td><td></td></tr><tr><td>Isoflurane</td><td>Vetone</td><td>502017</td><td></td></tr><tr><td>Lab Chart</td><td>Data Science International (DSI)</td><td></td><td>Software</td></tr><tr><td>Matrigel Matrix</td><td>BD Biosciences</td><td>356234</td><td></td></tr><tr><td>Micro Spring Scissors</td><td>World Precision Instruments (WPI)</td><td>14124</td><td></td></tr><tr><td>Mouse Ox Plus</td><td>STARR Life Sciences Corp.</td><td></td><td>Software</td></tr><tr><td>Mouse Ox Plus Collar Sensor</td><td>STARR Life Sciences Corp.</td><td>015022-2</td><td>Medium Collar Clip Special 7&amp;rdquo;</td></tr><tr><td>Mouse Whole Body Plethysmography Chamber</td><td>Data Science International (DSI)</td><td>PLY3211</td><td></td></tr><tr><td>Ohio Care Plus Incubator</td><td>Ohmeda</td><td>HCHD000173</td><td></td></tr><tr><td>Operating Scissors</td><td>World Precision Instruments (WPI)</td><td>501753-G</td><td>Straight</td></tr><tr><td>Osmotic Pump</td><td>Alzet</td><td>1003D</td><td>1 &amp;mu;L/h, 3 days</td></tr><tr><td>Phenol</td><td>Sigma-Aldrich</td><td>P4557</td><td></td></tr><tr><td>Recombinant Mouse Leptin protein</td><td>R&amp;amp;D systems</td><td>498-OB-05M</td><td>5 mg</td></tr><tr><td>Saline</td><td>RICCA Chemical</td><td>7210-16</td><td>0.9% Sodium Chloride</td></tr><tr><td>Sterile Surgical Suture</td><td>DemeTech</td><td>DT-639-1</td><td>Silk, size 6-0</td></tr><tr><td>Thermometer</td><td>Innovative Calibration Solutions (INNOCAL)</td><td>EW 20250-91</td><td></td></tr></tbody></table>

experimental approach to examine leptin signaling in the carotid bodies and its effects on control of breathing

An adipocyte-produced hormone leptin is a potent respiratory stimulant, which may play an important role in defending respiratory function in obesity. The carotid bodies (CB), a key organ of peripheral hypoxic sensitivity, express the long functional isoform of leptin receptor (LepRb) but the role of leptin signaling in control of breathing has not been fully elucidated. We examined the hypoxic ventilatory response (HVR) (1) in C57BL/6J mice before and after leptin infusion at baseline and after CB denervation; (2) in LepRb-deficient obese db/db mice at baseline and after LepRb overexpression in CBs. In C57BL/6J mice, leptin increased HVR and effects of leptin on HVR were abolished by CB denervation. In db/db mice, LepRb expression in CB augmented the HVR. Therefore, we conclude that leptin acts in CB to augment responses to hypoxia.

Continuous infusion of leptin significantly increased HVR in lean C57BL/6J mice from 0.23 to 0.31 mL/min/g/&#916;FiO2 (P &#60; 0.001, Figure 2)11. CSND abolished the leptin-induced increase in HVR (Figure 2), while no attenuating effects of CSND on HVR were observed in sham surgery group after leptin infusion.
LepRb expression in the CB of LepRb-deficient obese db/db mice induced a significant increase in HVR from 0.05 to 0.06 mL/min/g/&#916;SpO2 (Figure 3). In animals transfected with control Ad-LacZ in CB, HVR did not change.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60298/60298fig01.jpg" /> 
Figure 1: HVR measurements.&#160;Experiments should be performed at thermoneutral conditions using (A) a neonatal incubator at 30 &#176;C and are recorded in a (B) whole body plethysmography chamber. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60298/60298fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60298/60298fig02.jpg" /> 
Figure 2: Leptin augmented the hypoxic ventilatory response (HVR) and the effects were abolished by carotid sinus nerve dissection (CSND) in C57BL/6J mice.&#160;This figure has been modified from Caballero-Eraso et al.11. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60298/60298fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60298/60298fig03.jpg" /> 
Figure 3: LepRb expression in the carotid bodies (CB) of LepRb-deficient db/db mice increased the hypoxic ventilatory response (HVR).&#160;This figure has been modified from Caballero-Eraso et al.11. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60298/60298fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Experimental Approach to Examine Leptin Signaling in the Carotid Bodies and its Effects on Control of Breathing

Our study focuses on the effects of leptin signaling in carotid body (CB) on the hypoxic ventilatory response (HVR). We performed &#39;loss of function&#39; experiments measuring the effect of leptin on HVR after CB denervation and &#39;gain of function&#39; experiments measuring HVR after overexpression of the leptin receptor in CB.

An adipocyte-produced hormone leptin is a potent respiratory stimulant, which may play an important role in defending respiratory function in obesity. The ...

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5.9K Views.  Johns Hopkins University. Ons protocol kan worden gebruikt om de ademhalingseffecten van leptine op het halsslagaderlichaam in de perifere chemoreflex ademhalingscontrole en fysiologische reacties op hypoxie te onderzoeken. We kunnen ook de effecten van verlies en winst van halsslagader lichaam functie met behulp van chirurgische en chemische denervation en adenovirus transfectie respectievelijk. Voor het begin van de procedure, meet de temperatuur in een op maat gemaakte hele lichaam plasthemagrophy kamer, en meet de...

Video: Experimental Approach to Examine Leptin Signaling in the Carotid Bodies and its Effects on Control of Breathing

Breathing-controlled Electrical Stimulation (BreEStim) for Management of Neuropathic Pain and Spasticity

Electrical stimulation (EStim) refers to the application of electrical current to muscles or nerves in order to achieve functional and therapeutic goals. It has been extensively used in various clinical settings. Based upon recent discoveries related to the systemic effects of voluntary breathing and intrinsic physiological interactions among systems during voluntary breathing, a new EStim protocol, Breathing-controlled Electrical Stimulation (BreEStim), has been developed to augment the effects of electrical stimulation. In BreEStim, a single-pulse electrical stimulus is triggered and delivered to the target area when the airflow rate of an isolated voluntary inspiration reaches the threshold. BreEStim integrates intrinsic physiological interactions that are activated during voluntary breathing and has demonstrated excellent clinical efficacy. Two representative applications of BreEStim are reported with detailed protocols: management of post-stroke finger flexor spasticity and neuropathic pain in spinal cord injury.

Breathing-controlled Electrical Stimulation BreEStim for Management of Neuropathic Pain and Spasticity

The purpose is to present a new method, breathing-control electrical stimulation (BreEStim) for management of neuropathic pain and spasticity.

Electrical stimulation (EStim) refers to the application of electrical current to muscles or nerves in order to achieve functional and therapeutic goals. ...

Medicine

Method to Obtain Pattern of Breathing in Senescent Mice through Unrestrained Barometric Plethysmography

Unrestrained barometric plethysmography (UBP) is a method for quantifying the pattern of breathing in mice, where breathing frequency, tidal volume, and minute ventilation are routinely reported. Moreover, information can be collected regarding the neural output of breathing, including the existence of central apneas and augmented breaths. An important consideration for UBP is obtaining a breathing segment with a minimal impact of anxious or active behaviors, to elucidate the response to breathing challenges. Here, we present a protocol that allows for short, quiet baselines to be obtained in aged mice, comparable to waiting for longer bouts of quiet breathing. The use of shorter time segments is valuable, as some strains of mice may be increasingly excitable or anxious, and longer periods of quiet breathing may not be achieved within a reasonable timeframe. We placed 22 month-old mice in a UBP chamber and compared four 15 s quiet breathing segments between minutes 60&#8211;120 to a longer 10 min quiet breathing period that took&#160;2&#8211;3 h to acquire. We also obtained counts of central apneas and augmented breaths prior to the quiet breathing segments, following a 30 min familiarization period. We show that 10 min of quiet breathing is comparable to using a much shorter 15 s duration. Additionally, the time leading up to these 15 s quiet breathing segments can be used to gather data regarding apneas of central origin. This protocol allows investigators to collect pattern-of-breathing data in a set amount of time and makes quiet baseline measures feasible for mice that may exhibit increased amounts of excitable behavior. The UBP methodology itself provides a useful and noninvasive way to collect pattern-of-breathing data and allows for mice to be tested over several time points.

Unrestrained barometric plethysmography is used to quantify the pattern of breathing in awake mice. We show that 15 s segments under a standardized protocol display similar values to an extended duration of quiet breathing. This methodology also allows for the quantification of apnea and augmented breaths during the first hour in the chamber.

Unrestrained barometric plethysmography (UBP) is a method for quantifying the pattern of breathing in mice, where breathing frequency, tidal volume, and ...

Behavior

A Method of Nodose Ganglia Injection in Sprague-Dawley Rat

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located in the organs of the abdomen and thorax. The vagus nerve communicates information from stimuli such as heart rate, blood pressure, bronchopulmonary irritation, and gastrointestinal distension to the nucleus of solitary tract of the medulla. The cell bodies of the vagus nerve are located in the nodose and petrosal ganglia, of which the majority are located in the former. The nodose ganglia contain a wealth of receptors for amino acids, monoamines, neuropeptides, and other neurochemicals that can modify afferent vagus nerve activity. Modifying vagal afferents through systemic peripheral drug treatments targeted at the receptors on nodose ganglia has the potential of treating diseases such as sleep apnea, gastroesophageal reflux disease, or chronic cough. The protocol here describes a method of injection neurochemicals directly into the nodose ganglion. Injecting neurochemicals directly into the nodose ganglia allows study of effects solely on cell bodies that modulate afferent nerve activity, and prevents the complication of involving the central nervous system as seen in systemic neurochemical treatment. Using readily available and inexpensive equipment, intranodose ganglia injections are easily done in anesthetized Sprague-Dawley rats.

Afferent vagal signaling transmits important information to central nervous system from receptors located in organs of the abdomen and thorax. The nodose ganglia of vagus nerves contain many types of receptors that modulate vagal activity. This protocol describes a method of local injections of neurochemicals into the nodose ganglia.

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located ...

Neuroscience

Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to stimuli remain to be fully understood. To overcome the difficulty of accessing the brainstem in the whole animal, isolation of the brainstem and part of the spinal cord is performed. This preparation is maintained in artificial cerebro-spinal fluid where gases, concentrations, and temperature are controlled and monitored. The output signal from the respiratory network is recorded by a suction electrode placed on the fourth ventral root. In this manner, stimuli can be directly applied onto the brainstem, and the effect can be recorded directly. The signal recorded is linked to the inspiratory signal sent to the diaphragm via the phrenic nerve, and can be described as bursts (around 8 bursts per minute). Analysis of these bursts (frequency, amplitude, length, and area under the curve) allows precise characterization of the stimulus effect on the respiratory network. The main limitation of this method is the viability of the preparation beyond the early post-natal stages. Thus, this method greatly focuses on the study of the whole network without the peripheral inputs in the newborn rat.

The central respiratory drive is located in the brainstem. Spontaneous respiratory motor output from an isolated brainstem-spinal cord is recorded by placing an electrode on the fourth ventral root. This experimental approach is valuable for pharmacological investigations or the assessment of respiratory challenges and genetic manipulations on rhythmic motor behavior.

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to ...

Intraperitoneal Glucose Tolerance Test, Measurement of Lung Function, and Fixation of the Lung to Study the Impact of Obesity and Impaired Metabolism on Pulmonary Outcomes

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese individuals worldwide by 2030, thus representing a growing socioeconomic burden. Simultaneously, obesity-related comorbidities, including diabetes as well as heart and chronic lung diseases, are continuously on the rise. Although obesity has been associated with increased risk for asthma exacerbations, worsening of respiratory symptoms, and poor control, the functional role of obesity and perturbed metabolism in the pathogenesis of chronic lung disease is often underestimated, and underlying molecular mechanisms remain elusive. This article aims to present methods to assess the effect of obesity on metabolism, as well as lung structure and function. Here, we describe three techniques for mice studies: (1) assessment of intraperitoneal glucose tolerance (ipGTT) to analyze the effect of obesity on glucose metabolism; (2) measurement of airway resistance (Res) and respiratory system compliance (Cdyn) to analyze the effect of obesity on lung function; and (3) preparation and fixation of the lung for subsequent quantitative histological assessment. Obesity-related lung diseases are probably multifactorial, stemming from systemic inflammatory and metabolic dysregulation that potentially adversely influence lung function and the response to therapy. Therefore, a standardized methodology to study molecular mechanisms and the effect of novel treatments is essential.

The incidence of obesity is rising and increases the risk of chronic lung diseases. To establish the underlying mechanisms and preventive strategies, well-defined animal models are needed. Here, we provide three methods (glucose-tolerance-test, body plethysmography, and lung fixation) to study the effect of obesity on pulmonary outcomes in mice.

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese ...

Immunology and Infection

Evaluation of Vascular Control Mechanisms Utilizing Video Microscopy of Isolated Resistance Arteries of Rats

This protocol describes the use of in vitro television microscopy to evaluate vascular function in isolated cerebral resistance arteries (and other vessels), and describes techniques for evaluating tissue perfusion using Laser Doppler Flowmetry (LDF) and microvessel density utilizing fluorescently labeled Griffonia simplicifolia (GS1) lectin. Current methods for studying isolated resistance arteries at transmural pressures encountered in vivo and in the absence of parenchymal cell influences provide a critical link between in vivo studies and information gained from molecular reductionist approaches that provide limited insight into integrative responses at the whole animal level. LDF and techniques to selectively identify arterioles and capillaries with fluorescently-labeled GS1 lectin provide practical solutions to enable investigators to extend the knowledge gained from studies of isolated resistance arteries. This paper describes the application of these techniques to gain fundamental knowledge of vascular physiology and pathology in the rat as a general experimental model, and in a variety of specialized genetically engineered &#34;designer&#34; rat strains that can provide important insight into the influence of specific genes on important vascular phenotypes. Utilizing these valuable experimental approaches in rat strains developed by selective breeding strategies and new technologies for producing gene knockout models in the rat, will expand the rigor of scientific premises developed in knockout mouse models and extend that knowledge to a more relevant animal model, with a well understood physiological background and suitability for physiological studies because of its larger size.

This manuscript describes in vitro video microscopy protocols for evaluating vascular function in rat cerebral resistance arteries. The manuscript also describes techniques for evaluating microvessel density with fluorescently labeled lectin and tissue perfusion using Laser Doppler Flowmetry.

This protocol describes the use of in vitro television microscopy to evaluate vascular function in isolated cerebral resistance arteries (and other ...

Studying the Hypothalamic Insulin Signal to Peripheral Glucose Intolerance with a Continuous Drug Infusion System into the Mouse Brain

Insulin regulates systematic metabolism in the hypothalamus and the peripheral insulin response. An inflammatory reaction in peripheral adipose tissues contributes to type 2 diabetes mellitus (T2DM) development and appetite regulation in the hypothalamus. Chemokine CCL5 and C-C chemokine receptor type 5 (CCR5) levels have been suggested to mediate arteriosclerosis and glucose intolerance in type 2 diabetes mellitus (T2DM). In addition, CCL5 plays a neuroendocrine role in the hypothalamus by regulating food intake and body temperature, thus, prompting us to investigate its function in hypothalamic insulin signaling and the regulation of peripheral glucose metabolism.
The micro-osmotic pump brain infusion system is a quick and precise way to manipulate CCL5 function and study its effect in the brain. It also provides a convenient alternative approach to generating a transgenic knockout animal. In this system, CCL5 signaling was blocked by intracerebroventricular (ICV) infusion of its antagonist, MetCCL5, using a micro-osmotic pump. The peripheral glucose metabolism and insulin responsiveness was detected by the Oral Glucose Tolerance Test (OGTT) and Insulin Tolerance Test (ITT). Insulin signaling activity was then analyzed by protein blot from tissue samples derived from the animals.
After 7-14 days of MetCCL5 infusion, the glucose metabolism and insulin responsiveness was impaired in mice, as seen in the results of the OGTT and ITT. The IRS-1 serine302 phosphorylation was increased and the Akt activity was reduced in mice hypothalamic neurons following CCL5 inhibition. Altogether, our data suggest that blocking CCL5 in the mouse brain increases the phosphorylation of IRS-1 S302 and interrupts hypothalamic insulin signaling, leading to a decrease in insulin function in peripheral tissues as well as the impairment of glucose metabolism.

This protocol studies the role of chemokine (C-C motif) ligand 5 (CCL5) in the hypothalamus by delivering an antagonist, MetCCL5, into the mouse brain using a micro-osmotic pump brain infusion system. This transient inhibition of CCL5 activity interrupted hypothalamic insulin signaling, leading to glucose intolerance and peripheral systemic insulin sensitivity.

Insulin regulates systematic metabolism in the hypothalamus and the peripheral insulin response. An inflammatory reaction in peripheral adipose tissues ...

Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to non-invasively assess some features of the respiratory function in humans as well as in laboratory animals. The present article focuses on the application of the double-chamber plethysmography (DCP) in small animals. It provides background information on the methodology as well as a detailed step-by-step procedure to successfully assess respiratory function in conscious, spontaneously breathing animals in a non-invasive manner. The DCP can be used to monitor the respiratory function of multiple animals in parallel, as well as to identify changes induced by aerosolized substances over a chosen time period and in a repeated manner. Experiments on control and allergic mice are used herein to demonstrate the utility of the technique, explain the associated outcome parameters, as well as to discuss the related advantages and shortcomings. Overall, the DCP provides valid and theoretically sound readouts that can be trusted to evaluate the respiratory function of conscious small animals both at baseline and after challenges with aerosolized substances.

The objective of the present article is to provide a detailed description of the recommended procedures to evaluate respiratory function in conscious mice by double-chamber plethysmography.

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to ...

Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative assessment of lung function in rodent models is an important tool in the development of future therapies. Commonly used techniques for assessing respiratory function including invasive plethysmography and forced oscillation. While these techniques provide valuable information, data collection can be fraught with artefacts and experimental variability due to the need for anesthesia and/or invasive instrumentation of the animal. In contrast, unrestrained whole-body plethysmography (UWBP) offers a precise, non-invasive, quantitative way by which to analyze respiratory parameters. This technique avoids the use of anesthesia and restraints, which is common to traditional plethysmography techniques. This video will demonstrate the UWBP procedure including the equipment set up, calibration and lung function recording. It will explain how to analyze the collected data, as well as identify experimental outliers and artefacts that results from animal movement. The respiratory parameters obtained using this technique include tidal volume, minute volume, inspiratory duty cycle, inspiratory flow rate and the ratio of inspiration time to expiration time. UWBP does not rely on specialized skills and is inexpensive to perform. A key feature of UWBP, and most appealing to potential users, is the ability to perform repeated measures of lung function on the same animal.

The assessment of respiratory physiology has traditionally relied upon techniques, which require restraint or sedation of the animal. Unrestrained whole-body plethysmography, however, provides precise, non-invasive, quantitative analysis of respiratory physiology in animal models. In addition, the technique allows repeated respiratory assessment of mice allowing for longitudinal studies.

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative ...

Implantation of Combined Telemetric ECG and Blood Pressure Transmitters to Determine Spontaneous Baroreflex Sensitivity in Conscious Mice

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. The baroreflex is a key homeostatic mechanism to counteract acute, short-term changes in arterial BP and to maintain BP in a relatively narrow physiological range. BP is sensed by baroreceptors located in the aortic arch and carotid sinus. When BP changes, signals are transmitted to the central nervous system and are then communicated to the parasympathetic and sympathetic branches of the autonomic nervous system to adjust HR. A rise in BP causes a reflex decrease in HR, a drop in BP causes a reflex increase in HR.
Baroreflex sensitivity (BRS) is the quantitative relationship between changes in arterial BP and corresponding changes in HR. Cardiovascular diseases are often associated with impaired baroreflex function. In various studies reduced BRS has been reported in e.g., heart failure, myocardial infarction, or coronary artery disease. 
Determination of BRS requires information from both BP and HR, which can be recorded simultaneously using telemetric devices. The surgical procedure is described beginning with the insertion of the pressure sensor into the left carotid artery and positioning of its tip in the aortic arch to monitor arterial pressure followed by the subcutaneous placement of the transmitter and ECG electrodes. We also describe postoperative intensive care and analgesic management. After a two-week period of post-surgery recovery long-term ECG and BP recordings are performed in conscious and unrestrained mice. Finally, we include examples of high-quality recordings and the analysis of spontaneous baroreceptor sensitivity using the sequence method.

The baroreflex is a heart-rate regulation mechanism by the autonomic nervous system in response to blood-pressure changes. We describe a surgical technique to implant telemetry transmitters for continuous and simultaneous measurement of electrocardiogram and blood pressure in mice. This can determine spontaneous baroreflex sensitivity, an important prognostic marker for cardiovascular disease.

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. ...

Experimental Approach to Examine Leptin Signaling in the Carotid Bodies and its Effects on Control of Breathing

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