This work was supported by Ochanomizu University, KAKENHI grants (18K14859, 20K15895), JST FOREST Program, Grant Number JPMJFR204G, Research Foundation for Opto-Science and Technology, Kao Research Council for the Study of Healthcare Science, The Japan Association for Chemical Innovation, and TERUMO life science foundation. The author thanks Dr. Takashi Tsuboi, Kazuki Harada, and Akiyo Natsubori for&#160;their supervision of the experiments. The author also thanks members of the laboratory for their support.

All experimental protocols were approved by the Institutional Animal Care and Use Committee of Ochanomizu University, Japan (animal study protocols 22017). All animal experiments were performed according to the guidelines for animal experimentation of Ochanomizu University that conform with the Fundamental Guidelines for Proper Conduct of Animal Experiment and Related Activities in Academic Research Institutions (Ministry of Education, Culture, Sports, Science and Technology, Japan). Efforts were taken to minimize the number of animals used. This study was carried out in compliance with the ARRIVE guidelines.
1. Preparation of transgenic mice
<ol>
	<li>To follow this experiment, use adult male and female BAC GLT-1-G-CaMP7 #817 transgenic mouse line, namely G7NG817 mice24,25 (older than 8 weeks). The background strain of G7NG817 mice25 is C57BL/6J.</li>
	<li>House the mice under a 12 h /12 h light/dark cycle and raise them in groups of up to five mice.</li>
</ol>
2. Catheter creation
<ol>
	<li>Use scissors to adjust the silicon tube (inner diameter 0.5 mm, outer diameter 1.0 mm) to a precise length of 7 cm (Figure 1A-a). 
	NOTE: Prior to the procedures, scissors and silicon tube were sterilized.</li>
	<li>Adhere diminutive plastic beads (inner diameter 3 mm, outer diameter 5 mm) to the tailored tube, ensuring they are situated 3 mm from its terminus (Figure 1A-b). Affix the beads securely using cyanoacrylate glue (medical or standard variants are acceptable).</li>
	<li>Excise a 23 G needle&#39;s apex and subsequently cut 1.5 cm from the needle tip (Figure 1A-c).</li>
	<li>Introduce the sectioned 23 G needle from step 2.3 into the silicon tube&#39;s extremity, opposite to where the beads were affixed (Figure 1A-d).</li>
</ol>
3. Injector creation
<ol>
	<li>Use sterile pliers to excise 1 cm from the needle tip (Figure 1B-a). 
	NOTE: Ensure the orifice at the severed end retains its integrity and does not get deformed, as any distortion could obstruct the injection solution&#39;s flow.</li>
	<li>Couple the adapted 23 G injection needle from step 3.1.1 to a 2.5 mL syringe (Figure 1B-b).</li>
	<li>Segment a silicon tube to the requisite length (a 15 cm silicon tube in this experiment) and sheath it over the 23 G injection needle (Figure 1B-c). 
	NOTE: The volume of solution dispensed will fluctuate based on the tubing&#39;s length. For instance, with a 15 cm tube, approximately 50 &#181;L of solution will linger post experiment.</li>
	<li>Sever the 23 G injection needle 1.5 cm from the needle tip.</li>
	<li>Join the sectioned portion from step 3.4 with the silicon catheter fashioned in step 3.3 (Figure 1B-d). 
	&#8203;NOTE: Ensure the orifices created in steps 3.1 and 3.4 remain undistorted post incision. If rectification is needed, employ pliers to restore its circular configuration.</li>
</ol>
4. Gastric localization
<ol>
	<li>Administer anesthesia to the mouse via 3.0% isoflurane inhalation in a specified chamber. Apply ophthalmic ointment to prevent eye dryness.</li>
	<li>Delicately transfer the anesthetized mouse from the inhalation chamber to the surgical table upon verifying the mouse&#39;s anesthetized state. Provide thermal support throughout the procedure.</li>
	<li>Lay the mouse supine, aligning its mouth proximate to the inhalation apparatus, and modulate the isoflurane concentration from 3.0% to 2.0%. 
	NOTE: Disengage the inferior segment of the isoflurane mask from its superior counterpart and anchor the separated mask (upper segment) to the surgical table using adhesive tape.</li>
	<li>Supplement the mouse&#39;s oral cavity, forelimbs, and hindlimbs to the surgical table using adhesive tape. 
	NOTE: By fixing the mouse to the table, surgical errors due to movement can be prevented even if the mouse is anesthetized.</li>
	<li>Remove the mouse&#39;s hair from the upper left abdomen (150% of the area around the surgical site) by using depilatory cream. Apply a local analgesic topically on the surgery site and wait for 5 min before proceeding.</li>
	<li>Apply a local anesthetic gel to the abdominal skin and wait for 5 to 10 min. Then, disinfect the surgical area 3 times with alternating rounds of iodophor and 70% ethanol. Confirm the depth of anesthesia via a toe-pinch and cover the animal with sterile surgical drapes. Make a skin incision roughly 1.5 cm in length (Figure 1C-b), situated 1 cm to the right of the abdominal median and 5 mm below the xiphoid process (Figure 1C-c). Then, create a 1.5 cm incision in the abdominal wall at the same location as the initial skin incision. 
	NOTE: Exercise caution to avoid inflicting harm to the underlying liver.</li>
	<li>Delicately reposition the left hepatic lobe laterally using blunt-ended forceps, revealing the stomach beneath.</li>
</ol>
5. Catheter insertion
<ol>
	<li>Elevate the stomach and cautiously extricate it through the incision. 
	NOTE: Exercise utmost care to preserve the integrity of adjacent organs, notably the liver, and refrain from exerting undue force on the stomach.</li>
	<li>Use scissors to create a diminutive perforation (approximately 1.5mm in diameter) in the pyloric antrum.</li>
	<li>Introduce the catheter&#39;s terminus into the perforation, ensuring the bead is in direct contact with the pyloric antrum. 
	NOTE: The catheter is disinfected with a 0.1 to 0.5% chlorhexidine gluconate solution prior to insertion.</li>
	<li>Adhere the catheter&#39;s bead to the stomach employing medical-grade cyanoacrylate glue. 
	NOTE: Ensure the stomach remains unattached to surrounding organs.</li>
	<li>Verify the firm attachment of the catheter to the stomach, then delicately reposition the stomach to its innate locale beneath the left hepatic lobe.</li>
	<li>Stitch the abdominal wall, permitting the catheter to extend externally, using a polydioxanone (PDS) suture. After that, close the skin incision in a manner analogous to the abdominal closure.</li>
	<li>Gently reposition the mouse in a sanitized cage. 
	NOTE: Postoperative mice should be singularly housed. Apply a local analgesic topically on the wound closure and follow analgesic plans as recommeneded by the local institutional guidelines.</li>
	<li>Allow a minimum recovery period of 48 h in a standard environment. 
	NOTE: One end of the catheter inserted into the mouse&#39;s stomach is closed with a 23G needle with a sealed tip to prevent the entry of foreign substances and to avoid biting. Since the catheter protruding from the abdomen is short, it prevents the mouse from dragging it through the bedding, which could cause it to open or become entangled.</li>
</ol>
6. Preparation for in vivo transcranial Ca2+ imaging
<ol>
	<li>Administer anesthesia to the mouse using isoflurane (induction at 2%; maintenance between 0.8% and 1.0%).</li>
	<li>Secure the mouse onto a stereotaxic platform using auxiliary ear bars to mitigate the effects of pulsation and respiration. Subsequently, position the mouse under a fluorescence stereo microscope.</li>
	<li>Employ a wideband blue fluorescence filter set (with excitation at 460-490 nm and emission at 520 nm) in conjunction with a mercury light source.</li>
	<li>Capture the images using a camera and process them using the designated imaging software.</li>
	<li>Carefully remove the hair from the scalp using an electric shaver or hair removal cream. 
	NOTE: If using a razor, ensure that it is thoroughly disinfected and be careful not to cut the skin.</li>
	<li>Disinfect the surface of the scalp using a cotton swab or similar tool with a solution of chlorhexidine gluconate diluted to 0.1-0.5%. Next, disinfect with alcohol and repeat this alternating rounds 3x.</li>
	<li>Apply a local anesthetic gel and wait for 5-10 min.</li>
	<li>Prepare to apply dental acrylic cement.
	<ol>
		<li>Use a sterile scalpel blade to cut the scalp. Either completely remove the scalp or make a straight cut from the back of the head to the forehead.</li>
		<li>Use clips to expose the skull by extending any excess skin.</li>
		<li>Use a cotton swab to remove the connective tissue of the periosteum.</li>
		<li>Immediately apply the acrylic cement after removing the periosteum. Act quickly to prevent the skull surface from becoming opaque due to evaporation from the exposed bone.</li>
		<li>Wait for about 5 min for the cement to dry.</li>
	</ol>
	</li>
</ol>
7. Transcranial Ca2+ imaging with intraduodenal glucose administration
<ol>
	<li>Equilibrate both the experimental solution and saline to ambient temperature. Subject the mice to a dietary restriction for a specified duration (e.g., 2 h).</li>
	<li>Purge any residual content within the mouse-side catheter using ~0.03 mL of saline before initiating the injection.</li>
	<li>Aspirate the experimental solution (for instance, a 10% glucose solution prepared in 300 &#181;L of water, complemented with an additional 300 &#181;L of water) into the syringe, then interface it to the catheter. Ascertain the optimal dosage of the solution tailored to the specific experimental objective. 
	NOTE: During this phase, exercise caution to prevent undue strain on the catheter, which might jeopardize the mouse&#39;s internal anatomy. Endeavor to manipulate the mouse-side tubing with utmost gentleness. Additionally, ensure the syringe remains devoid of air post solution aspiration, as entrapped air and bubbles could induce gastrointestinal distension.</li>
	<li>Assess the mouse&#39;s physiological state and activate the imaging software&#39;s recording function by clicking on the recording process button of the software application.</li>
	<li>Capture images at a resolution of 512 x 512 pixels, 16 bit depth, and a frame rate of 10 Hz. 
	NOTE: The recording parameters can be adjusted to align with specific experimental requirements.</li>
	<li>Acquire spontaneous data (e.g., spanning 50 s), followed by the gradual infusion of the experimental solution at a rate tailored to the experiment (here, 0.035 mL/s). 
	NOTE: At the end of the experiment, the mouse will be euthanized promptly by cervical dislocation under isoflurane anesthesia.</li>
</ol>
8. Image data processing and analysis
<ol>
	<li>Bin image data into 64 x 64 pixels.</li>
	<li>Determine hand-drawn regions of interest (ROIs) using ImageJ. Refer to the mouse brain atlas to designate ROIs as the frontal area (M2), the somatosensory area including the Barrel (Somato), the occipital area (Visual), and the retrosplenial region (RSC).</li>
	<li>Use the MATLAB function ReadImageJROI to extract the coordinates corresponding to the ROIs. Access the MATLAB function through the provided link: https://jp.mathworks.com/matlabcentral/fileexchange/32479-readimagejroi</li>
	<li>Compute the average rate of fluorescence intensity change within the ROI using MATLAB.
	<ol>
		<li>Define the fluorescence intensity change rate (&#916;F/F) with equation (1): 
		<img alt="Equation 1" src="/files/ftp_upload/65902/65902eq01.jpg" style="margin: auto;" />&#160; &#160; &#160;(1) 
		Where Ft represents the fluorescence intensity value at a specific time and the average intensity value from the start of recording until 50 s into the injection is denoted as F0.</li>
	</ol>
	</li>
	<li>Account for the delay in the injection solution reaching the organ. Use the average intensity value spanning the three seconds post injection as a baseline. Target waveforms surpassing the baseline intensity value + 1 SD for analysis if they occur 3 s or later post injection.</li>
</ol>

Studying Gut-Brain Interaction via Transcranial Calcium Imaging in Mice

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The authors have no conflicts of interest to disclose.

This article presents how to attach a catheter to the gut and observe transcranial cortex wide Ca2+ signals with transgenic mice that express a genetically encoded Ca2+ indicator. In this experiment, we did not get the imaging data that use a secondary wavelength of the light source. This is because the transgenic mice we employed in this study exhibit a very high fluorescence intensity change rate (~several tens of percent) in response to sensory stimuli due to the properties of the Ca2+ indicator protein (G-CaMP7) employed. Hence, we thought that contamination from hemodynamics was negligible. Furthermore, the neural responses we focused on occurred rapidly within a few seconds, so we considered it unnecessary to account for subsequently generated local functional hyperemia. This catheter implantation method is able to combine with other genetically modified mice to observe neuronal activity in other brain regions and on different timescales. However, if other genetically modified mice are used, the control experiment described above should be performed.
This method of attaching the catheter to the gut is suitable for single-dose injection at low cost and is easy and less invasive than the method proposed earlier29. Traditionally, absorbent cotton is applied to the digestive organ and sutured. However, suturing can impair post-operative recovery in mice. Therefore, we adopted the method of attaching. The reason that we chose plastic beads is they adhered better to the silicone with the cyanoacrylate glue we use.
It has traditionally been postulated that the brain perceives glucose based on the escalation in plasma blood glucose levels and the concomitant hormone release. Consequently, the brain requires several minutes to hours to detect glucose following its absorption from the gut4. Some research has suggested that gut sensing controls mice&#39;s taste preference in addition to oral sweet taste reception1,3,17,22. As an illustrative example, researchers have provided evidence that intragastric (IG) glucose-paired flavored non-nutritive solution and IG nonmetabolizable &#945;-methyl-d-glucopyranoside (MDG)-paired flavored non-nutritive solution can elicit preference for each flavored non-nutritive solution compared to the IG water-paired group. As intraperitoneal glucose injection did not stimulate non-nutritive solution intake30, it was anticipated that gut nutrient sensing would play a crucial role in the food choices of the mice1,19,22.
A recent study has revealed the presence of an electrically excitable cell in the gut, referred to as a &#34;Neuropod cell,&#34; which employs molecular receptors to detect and transduce sensory stimuli onto a vagus nerve via a glutamatergic synapse. While previous studies have primarily focused on the role of GLP-1 in controlling mice&#39;s food feeding, recent investigations have demonstrated that vagus nerve activation by neuropod cells modulates mouse appetite. The vagus nerve subsequently activates the Nucleus Tractus Solitarius, leading to the subsequent activation of dopaminergic neurons19. However, it is unknown if dopaminergic neuron activation also activates the cortex. To categorize cerebral responses following intragastric glucose infusion, we refined the catheter attachment technique for imaging purposes. When employing this technology, it is imperative to exercise caution to prevent organ damage, with particular regard to the liver, which is susceptible to injury during the manipulation required to ascertain the stomach&#39;s location.
Our findings indicate that intragastric glucose injection activates the prefrontal cortex within seconds. However, intragastric glucose administration initiates multiple signaling pathways simultaneously, such as hormonal routes, the vagus nerve, and the spinal afferent nerve. These signals are processed in the brain stem on different timescales. Therefore, future studies should explore how these intricate signals are processed and integrated within the brain.
Moreover, the BAC GLT-1-G-CaMP7 transgenic mouse line we employed in the present study expresses the Ca2+ sensor, G-CaMP7, in both neurons and astrocytes. In principle, the prefrontal Ca2+ elevation obtained by transcranial cortex wide Ca2+ imaging is derived from neurons and/or astrocytes. However, since astrocytes express dopamine receptors if dopamine signaling is involved, it suggests the potential activation of astrocytic IP3/Ca2+ signaling. Since astrocytic IP3/Ca2+ signaling regulates synaptic plasticity, it may also play an important role in changes in sucrose preference and alterations in feeding behavior after glucose reception. Consequently, imaging at cellular resolution using two-photon microscopy would be required to identify whether the response observed in the prefrontal cortex following glucose administration originated from neurons or astrocytes in future studies. Another effective method involves observing cell-specific Ca2+ indicator expression in mice using adeno-associated virus (AAV) through fiber photometry. Furthermore, it is necessary to verify, using electrophysiological techniques, whether the glucose stimulus to the intestine genuinely stimulated the vagus nerve. If neuropod cells are transmitting signals to the vagus nerve by receiving glucose, we anticipate no response from the vagus nerve when an SGLT-1 inhibitor is administered. Finally, validation using various dopamine receptor antagonists would be necessary to confirm whether the vagus nerve projections to the Nucleus Tractus Solitarii (NTS) activated dopaminergic neurons, increasing prefrontal cortex Ca2+ signals. Overall, this system enables a comprehensive investigation of the rapid neural transmission effects from the gut to the brain. It presents a highly promising methodology that could potentially elucidate mechanisms, such as how stress alters sucrose preference regarding gut-brain interactions.

Primarily, gut sensing of glucose occurs via the sweet taste receptors (Tas1r2, Tas1r3) and the sodium-glucose cotransporter 1 (SGLT-1) located in enteroendocrine cells within the intestinal lumen1,2,3,4,5,6. The sensation of glucose in the brain is a process that typically spans from minutes to hours after its absorption from the gut, an occurrence principally based on increased levels of plasma blood sugar and the release of hormones7,8,9,10,11 (e.g., glucagon-like peptide-1 [GLP-1], Peptide YY (PYY), and glucose-dependent insulinotropic polypeptide [GIP]). They reach the arcuate nucleus of the hypothalamus (ARC), where they bind pro-opiomelanocortin (POMC) neurons and agouti-related protein (AgRP) neurons by blood flow and vagus nerve (parasympathetic nerve)12,13,14. The change in the activity of these neurons leads to control of metabolism and feeding behavior15,16.
Recent literature increasingly highlights the significance of gut-brain neural signaling following glucose absorption. This complex pathway can be broadly categorized into two primary mechanisms. The first involves the activation of spinal afferents, ultimately influencing AgRP neuronal activity in the ARC. The second mechanism entails stimulation of the vagus nerve through connections with epithelial cells. Given the multifaceted ways gut epithelial cells can activate the vagus nerve13, its role is considered versatile.
A significant proportion of gut vagus and spinal afferent nerve activities are associated with feeding behaviors17,18,19. However, emerging evidence suggests that certain vagus nerve fibers, projecting to &#34;neuropod cells,&#34; are implicated in glucose preference. Neuropod cells, a subset of enteroendocrine cells, express SGLT-1 on their luminal side and convey sensory stimuli to a vagus nerve via glutamatergic synapses2,20,21,22. Activation of the vagus nerve triggers dopaminergic neurons in the ventral tegmental area (VTA) within seconds19. Notably, inhibiting vagal activation, particularly in the duodenum, using a glutamate antagonist reduces sucrose preference in mice1. This underscores the critical role of neuropod cell activation in modulating food choice behavior post-glucose intake2.
Despite these advancements, a thorough understanding of the rapid intestinal glucose sensing mechanisms and their effect on cortical activities remains a complex enigma. Though brain activation controlled by AgRP neurons does not include the cerebral cortex23, it is known that dopaminergic neurons in the VTA project to the cerebral cortex. However, whether rapid VTA dopaminergic neuronal activation after glucose sensing truly activates the cerebral cortex is unknown. To elucidate this mechanism, we investigated the potential for intragastric glucose administration to rapidly influence cortical Ca2+ dynamics in transgenic mice expressing a genetically encoded calcium indicator.
This paper presents a low-cost and less invasive method to comprehensively understand the changes in cortical activity following intragastric glucose administration using cortex wide Ca2+ imaging in mice Recently, the transcranial cortex wide Ca2+ imaging technique through an intact skull, using transgenic mice expressing genetically encoded calcium indicators, has gained popularity24. Notably, the BAC GLT-1-G-CaMP7 #817 transgenic mouse line (also known as G7NG817 mouse) employed in this study expresses the Ca2+ sensor, G-CaMP7, in neurons and astrocytes25. Due to its high expression density in the cerebral cortex, this transgenic mouse line is especially suitable for transcranial cortex wide Ca2+ imaging using a standard epifluorescence microscope.

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real-time analysis of gut-brain neural communication: cortex wide calcium dynamics in response to intestinal glucose stimulation

Communication between the gastrointestinal tract and the brain after nutrient absorption plays an essential role in food preference, metabolism, and feeding behaviors. Particularly concerning specific nutrients, many studies have elucidated that the assimilation of glucose within gut epithelial cells instigates the activation of many signaling molecules. Hormones such as glucagon-like peptide-1 are renowned as quintessential signaling mediators. Since hormones predominantly influence the brain through circulatory pathways, they slowly modulate brain activity.
However, recent studies have shown two expeditious gut-brain pathways facilitated by the autonomic nervous system. One operates via the spinal afferent neural pathway, while the vagus nerve mediates the other. Consequently, brain responses following glucose assimilation in the gastrointestinal tract are complicated. Moreover, as intestinal stimulation finally induces diverse cortical activities, including sensory, nociceptive, reward, and motor responses, it is necessary to employ methodologies that facilitate the visualization of localized brain circuits and pan-cortical activities to comprehend gut-brain neural transmission fully. Some studies have indicated precipitous alterations in calcium ion (Ca2+) concentrations within the hypothalamus and ventral tegmental area independently through different pathways after intestinal stimulation. However, whether there are changes in cerebral cortex activity has not been known.
To observe cerebral cortex activity after intragastric glucose injection, we developed an imaging technique for real-time visualization of cortex wide Ca2+ dynamics through a fully intact skull, using transgenic mice expressing genetically encoded Ca2+ indicators. This study presents a comprehensive protocol for a technique designed to monitor intestinal stimulation-induced transcranial cortex wide Ca2+ imaging following intragastric glucose injection via an implanted catheter. The preliminary data suggest that administering glucose solution into the gut activates the frontal cortex, which remains unresponsive to water administration.

Transcranial cortex wide Ca2+ imaging with an isoflurane-anesthetized G7NG817 mouse 
We created the catheter following the procedure illustrated in Figure 1. After carefully identifying the position of the stomach, we attached the catheter to that location (Figure 1). After allowing sufficient recovery time following catheter insertion (approximately 48 h), we measured spontaneous neural activity under mild isoflurane anesthesia (0.8-1.0%) to establish the baseline fluorescence intensity changes for subsequent intestinal stimulation observations. As a result, we confirmed the random Ca2+ oscillation patterns that spanned the entire cortex (Figure 2A). Figure 2 provides representative examples of spontaneous brain activity, with the catheters inserted beforehand. We defined the time of the first observable Ca2+ transient peak in the M2 region following the injection as 0.0 s. The pseudo color representation displayed the peak of this Ca2+ transient as the maximum value, with the mean value plus one standard deviation (1 SD) as the minimum value. It was clear that the left and right hemispheres displayed synchronous activity in all regions.
Next, we established ROIs to characterize the waves within each functional area. As shown in Figure 2B, we presented the temporal changes in fluorescence intensity rate for each ROI (Figure 2B). These observed surface Ca2+ oscillations followed a burst suppression pattern, alternating between periods of low activity and high-amplitude fluorescence changes. This activity pattern aligns with previous findings that simultaneously measured cortical surface EEG and Ca2+ dynamics26.
Despite global or regional surface Ca2+ fluctuations between suppression and burst states, the specific cortical region initiating the calcium surge can differ. As previously reported, when a signal stimulates a local cortical region, the area is expected to activate, subsequently driving a global shift in the cortical state27,28.
Changes in cortex Ca2+ dynamics after intragastric glucose injection 
Next, to elucidate how glucose or water administration influences cortical activity, we monitored cortex wide Ca2+ dynamics during and after the injection. As a result, we found alterations in cortical Ca2+ dynamics following intragastric glucose injection. In contrast to the spontaneous activity illustrated in diverse regions of cortical activation, we observed immediate activation in the prefrontal area or the secondary motor cortex (M2) upon direct gut injection of glucose via the catheter, an effect not induced by intragastric water injection (Figure 3A-C). Moreover, we observed that these transient, prominent Ca2+ events tended to occur continuously over approximately 10 s following intragastric glucose injection. Such persistent phenomena were not apparent when water was administered (Figure 3D-F).
The Ca2+ events in each ROI were characterized by calculating the ratio of fluorescence intensity change ratio, using the auditory cortex (Reference) as a reference, at the first peak occurring within 4-8 s after the completion of glucose administration. As a result, a significant change was observed when glucose was injected after comparing the response in the M2 region using the auditory cortex as a reference before and after injection.
In contrast, no such change was observed when water was injected (Figure 3G, water N = 7: before vs. injection, 2.38 &#177; 2.53 vs. 2.05 &#177; 1.84, p &#62; 0.05; glucose N = 7: before vs. injection, 2.48 &#177; 0.97 vs. 3.76 &#177; 2.76, p &#60; 0.01, Wilcoxon signed-rank test). Next, defining the activation levels in each cortical region by the ratio of the value of after injection to before injection, a significant difference was observed only in the M2 region (Figure 3H, M2: 0.92 &#177; 0.89 vs. 1.30 &#177; 0.32, p = 0.006; Somato: 0.95 &#177; 0.076 vs. 1.04 &#177; 0.13, p = 0.27; Visual: 0.92 &#177; 0.3 vs. 1.00 &#177; 0.06, p = 0.89; RSC: 0.97 &#177; 0.27 vs. 1.10 &#177; 0.27, p = 0.60, t-test).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65902/65902fig01.jpg" /> 
Figure 1: Overview of the method. (A) Catheter creation: (a) Cut the silicon tube. (b) Affix a plastic bead to the silicon tube using medical cyanoacrylate glue. (c) Cut the 23 G needle 1.5 cm from the tip. (d) Attach the cut end of the needle to the silicon tube. (B) Injector assembly: (a) Trim a 23 G injection needle by 1 cm. (b) Connect the trimmed injection needle to a 2.5 mL syringe. (c) Sheath the cut section of the injection needle with a silicon tube. (d) Cut the 23 G needle 1.5 cm from the tip. (e) Attach the cut end of the needle to the silicon tube. (C) Surgical procedure: (a) Depilate the surgical area. (b) Incise the skin and the abdominal wall to a length of approximately 1.5 cm. (c) Create a small perforation (around 1.5 mm in diameter) in the pyloric antrum using scissors and insert the catheter (silicon tube). Secure the catheter to the pyloric antrum using medical cyanoacrylate glue. (d) Stitch the abdominal wall and skin using 5/0 silk suture material and needle. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/65902/65902fig01large.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/65902/65902fig02.jpg" /> 
Figure 2: Spontaneous cortex wide Ca2+ oscillations in G7NG817 mice with an attached catheter. (A) A representative example of cortex wide Ca2+ oscillations displayed at 0.2 s intervals. Changes in fluorescence intensity are represented with a pseudo color overlay. The time window encircled by the dashed line in B shows the first appearing Ca2+ transient, with its peak set as time 0. The pseudo color representation uses the peak of that Ca2+ transient as the maximum value and the mean +1 SD as the minimum value. (B) Time series changes in fluorescence intensity for each ROI in the example. Line colors correspond to different cortical areas: red represents M2; magenta represents the somatosensory area (Somato); blue represents the visual area (Visual); and green represents the retro splenial region. Salmon colored strip shows the injection time. (C) A single peak within the dashed outline is isolated and displayed in an enlarged view. Abbreviations: SD = standard deviation; ROI = region of interest; RSC = retro splenial region. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/65902/65902fig02large.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/65902/65902fig03.jpg" /> 
Figure 3: Cortex wide Ca2+ response during/after glucose or water injection. (A) A representative example of glucose administration displayed at 0.2 s intervals. Changes in fluorescence intensity are represented with a pseudo color overlay. The time window encircled by the dashed line in B shows the first appearing Ca2+ transient after the injection, with its peak set as time 0. The pseudo color representation is based on the peak of the first Ca2+ transient after injection as the maximum value and the mean +1 SD as the minimum value. (B) Time series changes of fluorescence intensity for each ROI in the example. Line colors correspond to different cortical areas: red represents M2, magenta represents the somatosensory area (Somato), blue represents the visual area (Visual), and green represents the retro splenial region. (C) A single peak within the dashed outline is isolated and displayed in an enlarged view. Analyze target calcium wave is the earliest wave that appeared within 4-8 s after injection. (D) A representative example of water injection as glucose control data. (E) Time series changes in fluorescence intensity for each ROI in the example. Line colors are as in B. (F) A single peak within the dashed outline is isolated and displayed in an enlarged view. Analyze target calcium wave is the earliest wave that appeared within 4-8 s after injection. (G) Comparison of M2 spontaneous activity (before) and response after injection (injected). The average fluorescence intensity changes over 50 s before injection, and the peak response in the M2 region between 4 s and 8 s after injection was calculated, using the auditory cortex as a reference. (H) Comparison of activation levels in each cortical region when water or glucose was administered. The ratio of the value of after injection to before injection, as determined in G, was calculated for each region. N.S. represents non-significant (p &#62; 0.05). ** &#60;0.01. Error bars are defined as the standard error of the mean. Abbreviations: SD = standard deviation; ROI = region of interest; RSC = retro splenial region. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/65902/65902fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Real-time Analysis of Gut-brain Neural Communication: Cortex wide Calcium Dynamics in Response to Intestinal Glucose Stimulation at JoVE.com

Real-time Analysis of Gut-brain Neural Communication: Cortex wide Calcium Dynamics in Response to Intestinal Glucose Stimulation

Gut-brain communication, facilitated by the vagus nerve, is crucial for communication between the gastrointestinal endocrine system and the brain. However, whether intragastric glucose injection can change cortical activity is still not understood. Here, we offer a comprehensive protocol to observe changes in cortical activity after glucose injection into the duodenum.

Communication between the gastrointestinal tract and the brain after nutrient absorption plays an essential role in food preference, metabolism, and ...

real-time-analysis-gut-brain-neural-communication-cortex-wide-calcium

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

554 Views.  Ochanomizu University. Gut-brain communication, facilitated by the vagus nerve, is crucial for communication between the gastrointestinal endocrine system and the brain. However, whether intragastric glucose injection can change cortical activity is still not understood. Here, we offer a comprehensive protocol to observe changes in cortical activity after glucose injection into the duodenum.

Video: Real-time Analysis of Gut-brain Neural Communication: Cortex wide Calcium Dynamics in Response to Intestinal Glucose Stimulation

Detection of Microregional Hypoxia in Mouse Cerebral Cortex by Two-photon Imaging of Endogenous NADH Fluorescence

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. Here we present a visualized experimental and methodological protocol to directly visualize microregional tissue hypoxia and to infer perivascular oxygen gradients in the mouse cortex. It is based on the non-linear relationship between nicotinamide adenine dinucleotide (NADH) endogenous fluorescence intensity and oxygen partial pressure in the tissue, where observed tissue NADH fluorescence abruptly increases at tissue oxygen levels below 10 mmHg1. We use two-photon excitation at 740 nm which allows for concurrent excitation of intrinsic NADH tissue fluorescence and blood plasma contrasted with Texas-Red dextran. The advantages of this method over existing approaches include the following: it takes advantage of an intrinsic tissue signal and can be performed using standard two-photon in vivo imaging equipment; it permits continuous monitoring in the whole field of view with a depth resolution of ~50 &mu;m. We demonstrate that brain tissue areas furthest from cerebral blood vessels correspond to vulnerable watershed areas which are the first to become functionally hypoxic following a decline in vascular oxygen supply. This method allows one to image microregional cortical oxygenation and is therefore useful for examining the role of inadequate or restricted tissue oxygen supply in neurovascular diseases and stroke.

Here we describe a method to directly visualize microregional tissue hypoxia in the mouse cortex in vivo. It is based on concurrent two-photon imaging of nicotinamide adenine dinucleotide (NADH) and the cortical microcirculation. This method is useful for high resolution analysis of tissue oxygen supply.

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. ...

Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex

Although the brain represents less than 5% of the body by mass, it utilizes approximately one quarter of the glucose used by the body at rest1. The function of non rapid eye movement sleep (NREMS), the largest portion of sleep by time, is uncertain. However, one salient feature of NREMS is a significant reduction in the rate of cerebral glucose utilization relative to wakefulness2-4. This and other findings have led to the widely held belief that sleep serves a function related to cerebral metabolism. Yet, the mechanisms underlying the reduction in cerebral glucose metabolism during NREMS remain to be elucidated.
	One phenomenon associated with NREMS that might impact cerebral metabolic rate is the occurrence of slow waves, oscillations at frequencies less than 4 Hz, in the electroencephalogram5,6. These slow waves detected at the level of the skull or cerebral cortical surface reflect the oscillations of underlying neurons between a depolarized/up state and a hyperpolarized/down state7. During the down state, cells do not undergo action potentials for intervals of up to several hundred milliseconds. Restoration of ionic concentration gradients subsequent to action potentials represents a significant metabolic load on the cell8; absence of action potentials during down states associated with NREMS may contribute to reduced metabolism relative to wake.
	Two technical challenges had to be addressed in order for this hypothetical relationship to be tested. First, it was necessary to measure cerebral glycolytic metabolism with a temporal resolution reflective of the dynamics of the cerebral EEG (that is, over seconds rather than minutes). To do so, we measured the concentration of lactate, the product of aerobic glycolysis, and therefore a readout of the rate of glucose metabolism in the brains of mice. Lactate was measured using a lactate oxidase based real time sensor embedded in the frontal cortex. The sensing mechanism consists of a platinum-iridium electrode surrounded by a layer of lactate oxidase molecules. Metabolism of lactate by lactate oxidase produces hydrogen peroxide, which produces a current in the platinum-iridium electrode. So a ramping up of cerebral glycolysis provides an increase in the concentration of substrate for lactate oxidase, which then is reflected in increased current at the sensing electrode. It was additionally necessary to measure these variables while manipulating the excitability of the cerebral cortex, in order to isolate this variable from other facets of NREMS.
	We devised an experimental system for simultaneous measurement of neuronal activity via the elecetroencephalogram, measurement of glycolytic flux via a lactate biosensor, and manipulation of cerebral cortical neuronal activity via optogenetic activation of pyramidal neurons. We have utilized this system to document the relationship between sleep-related electroencephalographic waveforms and the moment-to-moment dynamics of lactate concentration in the cerebral cortex. The protocol may be useful for any individual interested in studying, in freely behaving rodents, the relationship between neuronal activity measured at the electroencephalographic level and cellular energetics within the brain.

A procedure is described for manipulating the activity of cerebral cortical pyramidal neurons optogenetically while the electroencephalogram, electromyogram, and cerebral lactate concentration are monitored. Experimental recordings are performed on cable-tethered mice while they undergo spontaneous sleep/wake cycles. Optogenetic equipment is assembled in our laboratory; recording equipment is commercially available.

Although the brain represents less than 5% of the body  by mass, it utilizes approximately one quarter of the glucose used by the body  at rest1. The ...

Long-term Behavioral Tracking of Freely Swimming Weakly Electric Fish

Long-term behavioral tracking can capture and quantify natural animal behaviors, including those occurring infrequently. Behaviors such as exploration and social interactions can be best studied by observing unrestrained, freely behaving animals. Weakly electric fish (WEF) display readily observable exploratory and social behaviors by emitting electric organ discharge (EOD). Here, we describe three effective techniques to synchronously measure the EOD, body position, and posture of a free-swimming WEF for an extended period of time. First, we describe the construction of an experimental tank inside of an isolation chamber designed to block external sources of sensory stimuli such as light, sound, and vibration. The aquarium was partitioned to accommodate four test specimens, and automated gates remotely control the animals&#39; access to the central arena. Second, we describe a precise and reliable real-time EOD timing measurement method from freely swimming WEF. Signal distortions caused by the animal&#39;s body movements are corrected by spatial averaging and temporal processing stages. Third, we describe an underwater near-infrared imaging setup to observe unperturbed nocturnal animal behaviors. Infrared light pulses were used to synchronize the timing between the video and the physiological signal over a long recording duration. Our automated tracking software measures the animal&#39;s body position and posture reliably in an aquatic scene. In combination, these techniques enable long term observation of spontaneous behavior of freely swimming weakly electric fish in a reliable and precise manner. We believe our method can be similarly applied to the study of other aquatic animals by relating their physiological signals with exploratory or social behaviors.

We describe a set of techniques for studying spontaneous behavior of freely swimming weakly electric fish over an extended period of time, by synchronously measuring the animal&#39;s electric organ discharge timing, body position and posture both accurately and reliably in a specially designed aquarium tank inside a sensory isolation chamber.

Long-term behavioral tracking can capture and quantify natural animal behaviors, including those occurring infrequently. Behaviors such as exploration and ...

Real-time Electrophysiology: Using Closed-loop Protocols to Probe Neuronal Dynamics and Beyond

Experimental neuroscience is witnessing an increased interest in the development and application of novel and often complex, closed-loop protocols, where the stimulus applied depends in real-time on the response of the system. Recent applications range from the implementation of virtual reality systems for studying motor responses both in mice1 and in zebrafish2, to control of seizures following cortical stroke using optogenetics3. A key advantage of closed-loop techniques resides in the capability of probing higher dimensional properties that are not directly accessible or that depend on multiple variables, such as neuronal excitability4 and reliability, while at the same time maximizing the experimental throughput. In this contribution and in the context of cellular electrophysiology, we describe how to apply a variety of closed-loop protocols to the study of the response properties of pyramidal cortical neurons, recorded intracellularly with the patch clamp technique in acute brain slices from the somatosensory cortex of juvenile rats. As no commercially available or open source software provides all the features required for efficiently performing the experiments described here, a new software toolbox called LCG5 was developed, whose modular structure maximizes reuse of computer code and facilitates the implementation of novel experimental paradigms. Stimulation waveforms are specified using a compact meta-description and full experimental protocols are described in text-based configuration files. Additionally, LCG has a command-line interface that is suited for repetition of trials and automation of experimental protocols.

Closed-loop protocols are becoming increasingly widespread in modern day electrophysiology. We present a simple, versatile and inexpensive way to perform complex electrophysiological protocols in cortical pyramidal neurons in vitro, using a desktop computer and a digital acquisition board.

Experimental neuroscience is witnessing an increased interest in the development and application of novel and often complex, closed-loop protocols, where ...

Assessment of Neuromuscular Function Using Percutaneous Electrical Nerve Stimulation

Percutaneous electrical nerve stimulation is a non-invasive method commonly used to evaluate neuromuscular function from brain to muscle (supra-spinal, spinal and peripheral levels). The present protocol describes how this method can be used to stimulate the posterior tibial nerve that activates plantar flexor muscles. Percutaneous electrical nerve stimulation consists of inducing an electrical stimulus to a motor nerve to evoke a muscular response. Direct (M-wave) and/or indirect (H-reflex) electrophysiological responses can be recorded at rest using surface electromyography. Mechanical (twitch torque) responses can be quantified with a force/torque ergometer. M-wave and twitch torque reflect neuromuscular transmission and excitation-contraction coupling, whereas H-reflex provides an index of spinal excitability. EMG activity and mechanical (superimposed twitch) responses can also be recorded during maximal voluntary contractions to evaluate voluntary activation level. Percutaneous nerve stimulation provides an assessment of neuromuscular function in humans, and is highly beneficial especially for studies evaluating neuromuscular plasticity following acute (fatigue) or chronic (training/detraining) exercise.

We present a protocol to assess changes in neuromuscular function. Percutaneous electrical nerve stimulation is a non-invasive method that evokes muscular responses. Electrophysiological and mechanical properties of these responses permit the evaluation of neuromuscular function from brain to muscle (supra-spinal, spinal and peripheral levels).

Percutaneous electrical nerve stimulation is a non-invasive method commonly used to evaluate neuromuscular function from brain to muscle (supra-spinal, ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...

Real-time Iontophoresis with Tetramethylammonium to Quantify Volume Fraction and Tortuosity of Brain Extracellular Space

This review describes the basic concepts and protocol to perform the real-time iontophoresis (RTI) method, the gold-standard to explore and quantify the extracellular space (ECS) of the living brain. The ECS surrounds all brain cells and contains both interstitial fluid and extracellular matrix. The transport of many substances required for brain activity, including neurotransmitters, hormones, and nutrients, occurs by diffusion through the ECS. Changes in the volume and geometry of this space occur during normal brain processes, like sleep, and pathological conditions, like ischemia. However, the structure and regulation of brain ECS, particularly in diseased states, remains largely unexplored. The RTI method measures two physical parameters of living brain: volume fraction and tortuosity. Volume fraction is the proportion of tissue volume occupied by ECS. Tortuosity is a measure of the relative hindrance a substance encounters when diffusing through a brain region as compared to a medium with no obstructions. In RTI, an inert molecule is pulsed from a source microelectrode into the brain ECS. As molecules diffuse away from this source, the changing concentration of the ion is measured over time using an ion-selective microelectrode positioned roughly 100 &#181;m away. From the resulting diffusion curve, both volume fraction and tortuosity can be calculated. This technique has been used in brain slices from multiple species (including humans) and in vivo to study acute and chronic changes to ECS. Unlike other methods, RTI can be used to examine both reversible and irreversible changes to the brain ECS in real time.

This protocol describes real-time iontophoresis, a method that measures physical parameters of the extracellular space (ECS) of living brains. The diffusion of an inert molecule released into the ECS is used to calculate the ECS volume fraction and tortuosity. It is ideal for studying acute reversible changes to brain ECS.

This review describes the basic concepts and protocol to perform the real-time iontophoresis (RTI) method, the gold-standard to explore and quantify the ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a ...

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 ...

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural circuits. Due to the fractional change in the light signal, it has been difficult to use this method as a quantitative assay. This article describes special optics and slice handling systems, which render this technique stable and reliable. The present article demonstrates the slice handling, staining, and recording of the VSD-stained hippocampal slices in detail. The system maintains physiological conditions for a long time, with good staining, and prevents mechanical movements of the slice during the recordings. Moreover, it enables staining of slices with a small amount of the dye. The optics achieve high numerical aperture at low magnification, which allows recording of the VSD signal at the maximum frame rate of 10 kHz, with 100 pixel x 100-pixel spatial resolution. Due to the high frame rate and spatial resolution, this technique allows application of the post-recording filters that provide sufficient signal-to-noise ratio to assess the changes in neural circuits.

We introduce a reproducible and stable optical recording method for brain slices using voltage-sensitive dye. The article describes voltage-sensitive dye staining and recording of optical signals using conventional hippocampal slice preparations.

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural ...

Real-time Analysis of Gut-brain Neural Communication: Cortex wide Calcium Dynamics in Response to Intestinal Glucose Stimulation

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Detection of Microregional Hypoxia in Mouse Cerebral Cortex by Two-photon Imaging of Endogenous NADH Fluorescence

Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex

Long-term Behavioral Tracking of Freely Swimming Weakly Electric Fish

Real-time Electrophysiology: Using Closed-loop Protocols to Probe Neuronal Dynamics and Beyond

Assessment of Neuromuscular Function Using Percutaneous Electrical Nerve Stimulation

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Real-time Iontophoresis with Tetramethylammonium to Quantify Volume Fraction and Tortuosity of Brain Extracellular Space

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

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

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye