The authors would like to thank undergraduate Daniel Korus for his help running the Matlab code and undergraduate Alex Boettcher for his assistance in running the behavioral experiments. This project is supported by NSF IOS 1256799 to R.L.M., and by the National Institute on Drug Abuse of the National Institutes of Health under Award Number T32DA007234.

All procedures described here were approved by the Institutional Animal Care and Use Committee (IACUC) of The University of Minnesota, and are in accordance with The Guide for the Care and Use of Laboratory Animals13.
1. Animals and Cannulation Surgery
<ol>
	<li>Obtain Syrian hamsters from a common animal supplier at approximately 55 days of age. 
	NOTE: Although the age of animals will vary due to constraints of various experimental paradigms, for sexual behavior it is important to obtain sexually mature animals that are all approximately the same age.</li>
	<li>House animals in a controlled temperature (22 &#176;C) and lighting (14 h light followed by 10 h dark with lights out at 13:00 h) environment, with food and water available ad libitum except during periods of experimental testing. 
	NOTE: Since hamsters seasonally reproduce, this 14:10 light/dark cycle mimics their natural breeding conditions.</li>
	<li>Following a 1-week acclimation to the laboratory, perform aseptic bilateral ovariectomies and intracranial stereotaxic cannulations under general anesthesia (e.g., pentobarbital), and administer appropriate institutionally-approved post-operative analgesic and antibiotic treatment as previously described14,15. 
	NOTE: For the study of sexual behavior, ovariectomies are necessary to remove the major endogenous source of sexual hormones so that the animals can be uniformly induced to sexual receptivity with exogenous hormone in accordance with the experimental timeline.
	<ol>
		<li>For both dopaminergic and glutamatergic probe implantation, stereotaxically implant guide cannulae (0.7 mm diameter; Figure 3) into the area of interest (e.g., this lab targeted the right nucleus accumbens (NAc)) as described in stepwise detail elsewhere15. 
		NOTE: Although the data presented are from single sensor recordings of one glutamate or dopamine probe per animal, this system allows for simultaneous recording of up to 4 sensors in a single animal; thus, 1 - 4 cannulae could be implanted depending on desired experimental design and outcomes.
		<ol>
			<li>Determine the implant coordinates of the region of interest using a stereotaxic atlas (e.g., The Golden Hamster Brain by Morin &#38; Wood), keeping in mind that the sensor extends 1 mm below the cannula into brain tissue.</li>
		</ol>
		</li>
		<li>After a cannula is stereotaxically lowered to the desired dorsal-ventral location, affix it to the skull using a cap constructed from dental acrylic, secured with stainless steel bone screws and a plastic head mount (Figure 3; see15 for more details).</li>
		<li>For carbon fiber testing, insert a reference electrode (350 &#956;m diameter, 7.5 mm total length) into the contralateral hemisphere. 
		NOTE: A specific location or distance from the cannula is not required; reference electrodes may be placed anywhere in the contralateral hemisphere that is convenient.</li>
	</ol>
	</li>
</ol>
2. Biosensor and Carbon Fiber Testing
<ol>
	<li>Following 1-week of recovery from surgery, administer the hormone priming regimen to induce sexual receptivity.
	<ol>
		<li>Inject 10 &#181;g of estradiol benzoate in 0.1 mL of cottonseed oil subcutaneously (s.c.), approximately 48 h and 24 h prior to the sexual behavior testing, to induce sexual receptivity towards the male11.</li>
		<li>Inject 500 &#181;g of progesterone in 0.1 mL of cottonseed oil, s.c., 4 h prior to the introduction of the male to induce sexual receptivity11.</li>
	</ol>
	</li>
	<li>Test all animals at the start of the dark phase of their daily cycle, as social behaviors of rodents are subject to alteration under white light and red light testing conditions16,17,18.</li>
	<li>For enzymatic biosensor glutamate testing, calibrate the sensors in vitro (Figure 4) before use as described previously15,19.
	<ol>
		<li>Make the calibration solutions fresh on the day of testing in 20 mL glass centrifuge tubes. For the analyte solution, dissolve 7.4 mg of L-glutamic acid into 10 mL of ultra-pure H2O by gently inverting the tube. Make the interferent solution by dissolving 176.1 mg AA in 10 mL of ultra-pure H2O.</li>
		<li>Set up for the calibration by placing a magnetic stirrer on the base of a basic lab ring stand. Place a 20-mL jacketed beaker on top of the magnetic stirrer, and clamp in place using a medium 2-prong clamp.
		<ol>
			<li>Add a magnetic stir bar and 20 mL of 100 mM phosphate buffered saline (PBS) into the jacketed beaker. Connect the jacketed beaker to a circulating water bath to heat the buffer solution to 37 &#176;C. 
			NOTE: The enzymatic biosensor calibrations must be performed at body temperature since enzymatic activity is influenced by temperature.</li>
		</ol>
		</li>
		<li>Place the sensor calibration holder on top of the jacketed beaker, and set a 4-channel calibration preamplifier on top, securing in place using a right-angle clamp. Connect the preamplifier to a data conditioning and acquisition device to record calibration data.</li>
		<li>Test the sensitivity of each enzymatic biosensor to glutamate (or other analytes of interest for other commercially available biosensor probes e.g., glucose) before the experimental recording by placing the probe into the calibration holder atop a jacketed beaker, submerging the sensing cavity and some portion of the silver chloride (AgCl) reference wire in the buffer solution.
		<ol>
			<li>Before connecting each sensor (up to 4) being tested to a port on a 4-channel calibration preamplifier, initiate a new recording on the computer interface utilizing free downloadable acquisition software. Allow the sensors to reach a stable baseline.</li>
		</ol>
		</li>
		<li>Begin adding the analyte injections when the sensors have reached a stable baseline. Use the hole in the calibration holder to introduce a pipette tip to the buffer solution. Use the quick key annotation tool in the recording software to annotate when an injection is made.
		<ol>
			<li>Make 10 &#181;M additions of glutamate by pipetting 40 &#956;L of the analyte solution into the buffer solution. Wait for the sensor to stabilize between additions. Add 3 injections of analyte solution before adding a single injection from interferent AA solution (50 &#956;L injection volume for 250 &#956;M change in concentration). 
			NOTE: The calibration allows for conversion of the changes in the amperometric signal (seen during the enzymatic experimental recording in step 2.12) to changes in glutamate concentration, if desired. See reference15 for stepwise instruction. The dopaminergic probes used by this lab are calibrated during the manufacturing process by the commercial provider. Glutamate probes must be calibrated at the time of use as their sensitivity can decrease over time as the enzymatic components degrade19. Because carbon fiber electrodes do not undergo degradation, the calibration performed by the company prior to shipment is sufficient along as there is an increased likelihood of damaging the thin carbon fiber (350 &#956;m diameter) through additional handling.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Line the test chamber with bedding taken from the animal&#39;s home cage. 
	NOTE: This increases the female&#39;s familiarity with the testing chamber, as well as exposes the male to sexually stimulating olfactory mating cues that were distributed by the female in response to the estradiol treatment.</li>
	<li>Lightly anesthetize the animals prior to insertion of the probe by either isoflurane or another volatile anesthetic in an induction chamber.</li>
	<li>Remove the occlusive obdurator from the guide cannula, and insert either the carbon fiber electrode or enzymatic probe through the guide shaft.</li>
	<li>After the probe insertion, place the animal in the testing chamber. 
	NOTE: This lab uses a 10-gal glass aquarium (24.5 x 48.9 x 29.4 cm), but round chambers of equal area may also be employed.</li>
	<li>Begin recording the video and time-locked amperometric signal in a commercially available software program, described in detail elsewhere15.</li>
	<li>Connect the sensor to the recording system: the potentiostat via an electrically shielded cable and an electrical swivel. Stabilize the sensor connection by screwing a pin attachment onto the head mount. Reinforce the connection using lab film if necessary. 
	NOTE: A customized connector is needed to attach both the carbon fiber electrode and reference electrode to the potentiostat, while the enzymatic glutamate sensors can be directly connected to the potentiostat.</li>
	<li>Allow the sensor to equilibrate in the brain before experimental testing. 
	NOTE: Equilibration times differ depending on the recording sensor (e.g., enzymatic sensors require 2 - 4 h equilibration, while carbon-fiber electrodes only require about 30 min). If both types of sensors are implanted, equilibrate for the longer enzymatic sensor time.</li>
	<li>After sensor equilibration, introduce a stimulus male into the testing chamber.</li>
	<li>Following the first mount with penile insertion (termed intromission) by the stimulus male, continue recording for an additional 10 - 30 min, depending on the experimental goals.</li>
	<li>Following behavioral testing, disconnect the female&#39;s sensor.</li>
	<li>Remove both hamsters from the testing chamber. Remove the bedding and clean the chamber using 70% ethanol. 
	NOTE: Due to the brief equilibration required for carbon fiber recordings, steps 2.3 - 2.14 can be repeated to test multiple animals in the same day. In contrast, because the enzymatic sensors require about 4 h of equilibration, test only one animal per day to limit possible inter-subject variation due to differences in the animals&#39; circadian time at testing.</li>
</ol>
3. Sacrifice and Perfusion
<ol>
	<li>Following the conclusion of the testing period, deeply anesthetize the female test subject with a euthanizing agent (e.g., this lab uses an intraperitoneal injection of 0.2 mL phenytoin and pentobarbital solution).</li>
	<li>Transcardially perfuse the animal with 25 mM PBS, pH 7.6, to remove all circulating blood, followed by a 20 min fixation using 4% paraformaldehyde-PBS.</li>
	<li>Remove the brain and postfix in ~ 30 mL of the 4% paraformaldehyde solution in a 50-mL conical tube overnight.
	<ol>
		<li>Store the brain in a 10% sucrose-PBS solution until ready to serial section. 
		NOTE: Brains may be stored up to 2-weeks, with weekly sucrose solution changes to avoid potential growth of contaminants.</li>
	</ol>
	</li>
	<li>Serial section the brain in 40 &#181;m slices through the implanted region with either a freezing microtome or cryostat as previously described19.</li>
	<li>Mount slices serially on commercially available adhesive-coated slides (or see20 to make). Allow to dry, at least overnight.</li>
	<li>Stain the slides with cresyl violet dye, clear, and coverslip as previously described20.</li>
	<li>Image the slides using bright field microscopy to confirm the anatomical placement of the sensor.</li>
</ol>
4. Behavioral Coding
<ol>
	<li>View and annotate the videos using free commercially downloadable software (see Table of Materials) in slow motion to precisely code behaviors time-locked to the amperometric signal. 
	NOTE: To use the MATLAB code in the analysis (see below), annotate the start and end of each of the female&#39;s and male&#39;s behaviors.
	<ol>
		<li>Annotate startLordosis in the frame when the female initiates a dorsoflexion of her back and defects her tail upwards. Annotate endLordosis when the female terminates this posture, which often occurs when the female readjusts to another location in the testing chamber.</li>
		<li>Annotate startAI when the female is in the lordosis posture and the male moves his snout towards the female&#39;s anogenital region, where sniffing, licking, or nuzzling of her perineal region may occur. Annotate endAI when the male removes his snout from close proximity to the female&#39;s anogenital region.</li>
		<li>Annotate startMount when the male approaches and places his forepaws on the female in a mounting posture, regardless of the orientation of the mount attempt (e.g., side, rump).</li>
		<li>Annotate startIntromission when successful thrusting gains the mounted male penile access to the female&#39;s vagina. 
		NOTE: An intromission is behaviorally distinguishable from the mounted thrusting that occurs prior to intromission, as the male visibly pulls his upper limbs towards his body, thrusts his pelvis forward, and curls his tail upwards, indicating penetration (see Figure 4C).
		<ol>
			<li>When applicable, annotate ejaculation. 
			NOTE: At the end of a mating bout and in conjunction with a successful intromission, the male will ejaculate. Although one is unable to visualize the actual emission, male hamsters have a characteristic treading motion with their hind foot during an intromission21, thus annotate the ejaculation at the occurrence of the foot movement.</li>
		</ol>
		</li>
		<li>Annotate endIntromission and endMount simultaneously in the frame when the male has removed his front two paws and thus, has no contact with the female. Due to the frequent inability to visualize his withdrawal, simultaneous code these two behaviors to allow for consistency and reliability across mating bouts. 
		Note: A mating bout either follows a successful ejaculation or is when the female breaks the lordosis posture following at least one mount.</li>
	</ol>
	</li>
</ol>
5. Example Data Analysis
Note: The use of fixed-potential carbon fiber and enzymatic recordings for dopamine and glutamate, respectively, allows for the ability to measure both tonic and phasic patterns of electrochemical release. This lab uses a free, commercial acquisition system and software to record and convert recorded signals from analog to digital (bias 0.600 V, sampling rate = 1 Hz).
<ol>
	<li>Description of example data analysis protocol
	<ol>
		<li>Although other programs can be used, quantify the tonic and phasic (transient) changes in the recorded electrochemical signal using MATLAB.
		<ol>
			<li>For the presented data, calculate the tonic signal using a 50-point moving average filter. A transient-only normalized representation of the signal was calculated by subtracting the tonic signal from the raw data. When identifying phasic peaks, there are various algorithmic options (e.g., labeling peaks as changes in the signal more than 1 SD above the mean).</li>
			<li>Identify the locations of these peaks, i.e., local maxima, from the normalized signal using the MATLAB function peakfinder with the minimum peak amplitude set to the root mean square of the signal. The location of peaks detected from the normalized signal was used to extract the amplitude from each peak. The ability to detect peaks allows for the identification of any time-locked behaviors that may be coinciding with, or driving the relative peak in the neurochemical under investigation.</li>
		</ol>
		</li>
		<li>Segment the processed data in accordance with the behavioral annotations (see step 3); a Student&#39;s t-test was used to evaluate differences in the tonic neurotransmitter level, the occurrence of transients, and the amplitude of transients during the pre-mating bout lordosis versus lordosis during mating bouts. To measure changes to tonic patterns of electrochemical release, the data points occurring during the Lordosis position directly before a mating bout (i.e., pre-mating bout lordosis) were compared to the data points during a mating bout session using a t-test.</li>
	</ol>
	</li>
	<li>Preparation of raw data for described data analysis protocol 
	Note: As stated, although other programs or analyses may be equally viable options, in order to use the MATLAB code described above, the raw data must be prepared in the following way:
	<ol>
		<li>Export the annotation and voltage files. To do this, under the &#39;File&#39; tab, select the &#39;Export&#39; function and select the &#39;Annotations&#39; tab to save the annotations. Under the &#39;File&#39; tab, select the &#39;Export&#39; function and choose the &#34;TSV&#39; tab to save the voltage measurements as a TSV extension file.</li>
		<li>Open each file with a spreadsheet program and save as &#39;xls/xlsx&#39; to make the files compatible with MATLAB. 
		NOTE: For the &#39;Annotation&#39; file, there are additional annotations that must manually added post-export in a spreadsheet program to dictate the start and end of both the pre-mating bout lordosis (annotated as PreMBlordorsis) and the mating bout so that these time points can be directly compared as mentioned in step 5.1.2.</li>
		<li>Annotate the start of the pre-mating bout lordosis (startPreMBlordosis) at the same time as a lordosis behavior begins (startLordosis) that also includes a mount behavior, and also at the start of a new mating bout if the female remains in the lordosis posture.</li>
		<li>Annotate the end of the pre-mating bout lordosis (endPreMBlordosis) when the first mount begins.</li>
		<li>Annotate the start of a mating bout (startMB) at the same time of the endPreMBlordosis, as this signifies the start of a mating bout.</li>
		<li>Annotate the end of a mating bout (endMB) at either the end of a mount that includes an ejaculation (see step 4.1.4.1), or following the female&#39;s termination of the lordosis posture (endLordosis).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Although relatively straightforward, some issues can arise when employing this technique. First, the stereotaxic placement of the probes must be precise: unlike microdialysis that samples a wider radius of the extracellular milieu surrounding the probe, this technique only allows the measurement of a neurotransmitter that comes into direct contact with the probe. Second, in the case of the carbon fiber recording, due to the small width of fiber, breakage can occur, and the probe must be inserted with deliberate care. In ...

The ability to measure neurotransmitter release in awake behaving animals allows researchers to link specific behaviors with spatial and temporal patterns of neurotransmission-a powerful tool to investigate mechanisms and circuitry underlying both natural and operant behaviors in real-time. Historically, microdialysis has been employed to measure both electrically reactive and nonreactive substances in the extracellular milieu of the brain1. This technique uses a continuous flow of an aqueous solution of similar ionic composition to the extracellular fluid, through a microdialysis probe composed of a small shaft with a tip made of a semipermeable hollow fiber membrane2. After insertion of the probe, neurotransmitters or other analytes of interest can cross the semipermeable membrane by passive diffusion before being collected at intervals for subsequent analysis by high-performance liquid chromatography (HPLC), an analytical chemistry technique commonly utilized to separate, identify, and quantify components in a heterogeneous mixture3.
Although microdialysis is a sensitive technique that can be used to measure virtually any analyte of interest, the temporal resolution is low, with maximum sampling rates on the order of minutes to tens of minutes1,2. The invention of fast scan cyclic voltammetry (FSCV), a technique that relies on the redox potential of electroactive species, can elucidate near instantaneous concentrations of the analyte of interest in the extracellular fluid. In brief (see Robinson et al.4 for an extensive review), an electrode is applied to raise and lower the voltage in a triangular wave fashion on a fast time scale4. When the voltage is in the correct range, the compound of interest is repeatedly oxidized and reduced. This oxidization and reduction results in a movement of electrons that creates a small alternating current. Scan rates take place on the sub-second scale with oxidization and reduction of compounds occurring in microseconds. By subtracting the background current created by the probe from the resulting current, one can generate a voltage vs. current plot unique to each compound. Since the time scale of the voltage oscillations is known, these data can be used to calculate a plot of the current as a function of time. Thus, the relative concentrations of the compound may be determined as long as the number of electrons transferred in each oxidation and reduction reaction is known4.
This chemical specificity and high temporal resolution make FSCV a powerful technique for detecting changing chemical concentrations in vivo. However, despite these manifold advantages, this technique requires extensive technical expertise and expensive equipment and setup. Further, nonelectroactive neurotransmitters (e.g., glutamate) cannot be measured using this technique. Fortunately, technological advancements in the field of electrochemistry5, as well as commercialization of these inventions, has introduced a relatively simple approach to measure non-electroactive neurotransmitters in awake behaving animals without compromising temporal precision-a technique known as enzymatic biosensor technology. This technique uses enzymatic conversion of the nonelectroactive neurotransmitter of interest into two substrates, one of which is electroactive hydrogen peroxide that is detected as an amperometric oxidation current generated by an applied potential5. Commercially available biosensor probes (see Figure 1) selectively measure analytes of interest by competitively reducing the contribution of endogenous interferents. In the case of glutamate, the contribution of the common interferent ascorbic acid (AA) is competitively reduced to the measured current by co-localizing AA oxidase onto the active enzymatic surface of the sensor, converting AA to non-electroactive dihydroascorbate and water. In addition, a negatively charged Nafion polymer layer present under the enzyme layer excludes endogenous anionic compounds.
This same biosensor experimental setup can measure electroactive neurotransmitters as in FSCV, but instead it employs a fixed-potential recording6. In contrast to the oscillating voltage applied in FSCV, in a fixed-potential recording the voltage is kept at the redox potential for the analyte of interest. Although it is less chemically selective than FSCV as multiple neurotransmitters may have the same redox potential, in brain areas that overwhelmingly skew towards one neurotransmitter, the turn-key nature of this approach outweighs the lack of chemical specificity.
The ability to measure both electroactive and nonelectroactive neurotransmitter release in near real-time and link it to specific behavioral events provides an opportunity to examine converging neurotransmitter release. This manuscript details the use of this system to interrogate both dopamine and glutamate neurotransmission in response to natural reward in awake behaving hamsters. The aim of this paper is to detail the process of measuring this neurotransmitter release during sexual behavior in female hamsters, with the goal of demonstrating its feasibility for examining other behaviors and experimental paradigms.
Hamsters are an ideal model for use in electrochemical recordings 
Historically, rat and mice models have been employed in the study of sexual behavior. These rodent species engage in a dynamic copulatory sequence, involving numerous female solicitation behaviors that include hopping, darting, and ear wiggling to entice the male to chase and ultimately mount the female7. The mounting by the male (with or without vaginal penetration) lasts only a few seconds, during which the female engages in her sexual behavior posture (termed lordosis) also only for a few seconds before resuming active solicitation behaviors. This pattern of behavior, composed of high levels of activity interspersed with brief periods of immobility, is problematic for measuring neurotransmission in behaving animals. First, there can be movement artifacts in the amperometric recordings that are unrelated to neural activity. Second, the locomotion is associated with the release of particular neurotransmitters in certain brain regions. For example, dopamine release has been coupled to locomotor activity in the dorsal and ventral striatum8,9, a finding that formed the basis for microdialysis measurements of dopamine following psychostimulant administration10. Because the female-typical solicitation behaviors in most rodents involve high levels of locomotor activity, and are represented by the bulk of a 10 minute sexual behavior test, this makes it difficult to ascribe changes in neurotransmission to the explicit components of sexual behavior that collectively last only minutes.
To analyze the neurochemical profile of female sexual behavior, this lab sought out a species in which there is minimal locomotor activity accompanying sexual behavior. The copulatory sequence in Syrian hamsters (Mesocricetus auratus) is ideal for neurochemical recordings due to the lack of solicitation behaviors typically seen in rats and mice11. As a consequence, female hamsters will enter and maintain the lordosis posture for upwards of 9 minutes out of a 10 minute testing session12. With the lack of extraneous locomotor movements by the female, in vivo electrochemical recordings that can be associated with components of sexual interactions with the male can be obtained.
Copulatory bouts in hamsters 
After the introduction of a male stimulus animal into the testing chamber, the male will initially engage in anogenital investigation (AI) of the female before mounting her (Figure 2A). In order for the male to mount, the female must assume a receptive sexual posture known as lordosis, in which she arches her back and deflects her tail so that the mounting male can gain penile access to her vagina. The male will mount the female, clasping her hindquarters with both paws (Figure 2B), and begin thrusting in an attempt to gain penile intromission (Figure 2C). The male will mount the female (without insertion) as well as intromit a number of times before eventually achieving ejaculation. This sequence of mounts and intromissions leading to ejaculation is termed a &#34;copulatory bout&#34;. Males will have several copulatory bouts within a single session.

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			<td style="width:183px;">Oak Pharmaceuticals Inc.</td>
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			<td style="width:504px;">Pentobarbital sodium injection, USP. This lab uses 8.5mg/100g body weight, injected intraperotineally.&#160;</td>
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			<td height="42" style="height:42px;width:329px;">Loxicom analgesic&#160;</td>
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			<td style="width:504px;">NSAID antinflammatory and analgesic used for post-operative pain control. Generic: meloxicam.&#160;</td>
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			<td style="width:183px;">Norbrook Laboratories&#160;</td>
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			<td style="width:504px;">Fluoroquinolone antibiotic for post-operative infection prevention. Generic: Enrofloxacin.</td>
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			<td style="width:504px;">Pentobarbital Sodium, Phenytoin Sodium euthanasia agent.</td>
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			<td>Pinnacle Technologies, Inc.</td>
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			<td>Pinnacle Technologies, Inc.</td>
			<td>8400-K2</td>
			<td style="width:504px;">Pinnacle offers complete hardware kits for new users of our tethered biosensor system for rats. Kits include a commutator, preamplifier, and data conditioning and acquisition system</td>
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			<td>Pinnacle Technologies, Inc.</td>
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			<td style="width:504px;">The base computer package includes a preconfigured computer with ample hard disk storage, a high-definition monitor, a keyboard and mouse, an uninterruptible power supply, and all necessary cables.&#160;</td>
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			<td height="60" style="height:60px;">Video EQ700 EverFocus camera&#160; package</td>
			<td>Pinnacle Technologies, Inc.</td>
			<td>&#160;9000-K10&#160;</td>
			<td style="width:504px;">EQ700 night vision capable box camera with independent IR source was obtained as part of Pinnacle video computer package. Dome camera (9000-K9) and HD camera (9000-K11) options are also available.&#160;</td>
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			<td height="126" style="height:126px;">Sirenia Acquisition software</td>
			<td>Pinnacle Technologies, Inc.</td>
			<td>Free--available to download from pinnaclet.com</td>
			<td style="width:504px;">Sirenia&#160;Acquisition provides a single platform for recording data from any Pinnacle hardware system. The software features synchronization of all data streams, user-configurable settings, data consolidation, and multiple export options. In addition, the software includes basic review and analysis modules for biosensor recordings. Sirenia&#160;delivers free ll-in-one software that is ideal for data acquisition and review.</td>
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			<td height="147" style="height:147px;">Tethered rat&#160;in vitro&#160;calibration kit</td>
			<td>Pinnacle Technologies, Inc.</td>
			<td>7000-K2-T-BAS&#160;</td>
			<td style="width:504px;">In order to relate the current changes measured by a biosensor to actual changes in analyte concentration, it is necessary to calibrate the biosensor prior to implantation into the animal. The process also confirms the integrity and selectivity of the sensors.&#160;Calibration kit includes 20 mL jacketed beaker (#7058), 1/2&#34; by 1/8&#34; magnetic stir bar (#7059), right angle clamp (#7056), 2 prong single-adjustment clamp (#7055), 4-channel calibration preamplifer (#7053), and calibration holder (#7051).&#160;</td>
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			<td>Corning</td>
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			<td style="width:504px;">Corning digital Stirrer, 5&#34; x 7&#34;, 120 VAC used to spin magnetic stirrer in jacketed beaker during in vitro calibration of glutamate biosensors.</td>
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			<td height="63" style="height:63px;">Water bath capable of closed loop circulation</td>
			<td>PolyScience</td>
			<td>8006A11B</td>
			<td style="width:504px;">PolyScience 8006A11B 6L Standard Digital Heated Circulating Bath, 120VAC water bath was used with plastic tubing to heat jacketed beaker to physiological temperature.&#160;</td>
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			<td height="21" style="height:21px;">Carbon fiber sensor with BASi rat cannulae</td>
			<td>Pinnacle Technology, Inc.</td>
			<td>7002-CFS</td>
			<td style="width:504px;">Carbon fiber electrode used for recording dopamine neurotransmission.</td>
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			<td height="21" style="height:21px;">Ag/AgCl reference electrode</td>
			<td>Pinnacle Technology, Inc.</td>
			<td>7065</td>
			<td style="width:504px;">Necessary for carbon fiber recordings.</td>
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			<td height="42" style="height:42px;">Glutamate biosensors&#160;</td>
			<td>Pinnacle Technology, Inc.</td>
			<td>7001&#160;</td>
			<td style="width:504px;">Enzymatic biosensor probe used for recording glutamatergic neurotransmission.</td>
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			<td height="21" style="height:21px;">BASi guide cannulae</td>
			<td>Pinnacle Technologies, Inc.</td>
			<td>7030&#160;</td>
			<td style="width:504px;">Guide cannulae implanted into brain region of interest to guide probe.</td>
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			<td height="21" style="height:21px;">BASi cannula plastic headpiece for rats&#160;</td>
			<td>Pinnacle Technologies, Inc.</td>
			<td>7011&#160;</td>
			<td style="width:504px;">Headmount stabilizes probe and attaches to potentiostat.</td>
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measuring in vivo changes in extracellular neurotransmitters during naturally rewarding behaviors in female syrian hamsters

The ability to measure neurotransmitter release on a rapid time scale allows patterns of neurotransmission to be linked to specific behaviors or manipulations; a powerful tool in elucidating underlying mechanisms and circuitry. While the technique of microdialysis has been used for decades to measure nearly any analyte of interest in the brain, this technique is limited in temporal resolution. Alternatively, fast scan cyclic voltammetry is both temporally precise and extremely sensitive; however, because this technically difficult method relies on the electroactivity of the analyte of interest, the possibility to detect nonelectroactive substances (e.g., the neurotransmitter glutamate) is eliminated. This paper details the use of a turn-key system that combines fixed-potential amperometry and enzymatic biosensing to measure both electroactive and nonelectroactive neurotransmitters with temporal precision. The pairing of these two powerful techniques allows for the measurement of both tonic and phasic neurotransmission with relative ease, and permits recording of multiple neurotransmitters simultaneously. The aim of this manuscript is to demonstrate the process of measuring dopamine and glutamate neurotransmission in vivo using a naturally rewarding behavior (i.e., sexual behavior) in female hamsters, with the ultimate goal of displaying the technical feasibility of this assay for examining other behaviors and experimental paradigms.

Using the electrochemical and behavioral coding methodology described above, this lab has begun to characterize both tonic and phasic fluctuations in both dopamine and glutamate during in vivo recordings of sexual behavior. Due to the temporally-precise manner of this methodology, we can more accurately characterize neurotransmission during sexual behavior; as well as ascribe specific changes in release patterns to corresponding tonic changes during mating bouts, and correspondin...

Watch this Scientific Journal Video about Measuring In Vivo Changes in Extracellular Neurotransmitters During Naturally Rewarding Behaviors in Female Syrian Hamsters at JoVE.com

Measuring In Vivo Changes in Extracellular Neurotransmitters During Naturally Rewarding Behaviors in Female Syrian Hamsters

This paper details the use of fixed-potential amperometric recordings using carbon fiber electrodes and enzymatic biosensor technology to measure the release of dopamine and glutamate with high temporal resolution during natural rewarding behavior in the female hamster.

Measuring In Vivo Changes in Extracellular Neurotransmitters During Naturally Rewarding Behaviors in Female Syrian Hamsters

The ability to measure neurotransmitter release on a rapid time scale allows patterns of neurotransmission to be linked to specific behaviors or ...

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10.1K 조회수.  University of Minnesota. 이 문서 내용을 사용 하 여 탄소 섬유 전극 효소 바이오 센서 기술에 자연적인 보상 행동 동안 도파민과 높은 시간 해상도와 조미료의 릴리스를 측정 하는 고정 잠재력 amperometric 녹음의 사용은 여자는 햄스터

비디오: Measuring In Vivo Changes in Extracellular Neurotransmitters During Naturally Rewarding Behaviors in Female Syrian Hamsters

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

In vivo Laser Axotomy in C. elegans

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron is damaged by injury or disease, it may regenerate. Cell-intrinsic and extrinsic factors influence the ability of a neuron to regenerate and restore function. Recently, the nematode C. elegans has emerged as an excellent model organism to identify genes and signaling pathways that influence the regeneration of neurons1-6. The main way to initiate neuronal regeneration in C. elegans is laser-mediated cutting, or axotomy. During axotomy, a fluorescently-labeled neuronal process is severed using high-energy pulses. Initially, neuronal regeneration in C. elegans was examined using an amplified femtosecond laser5. However, subsequent regeneration studies have shown that a conventional pulsed laser can be used to accurately sever neurons in vivo and elicit a similar regenerative response1,3,7.
We present a protocol for performing in vivo laser axotomy in the worm using a MicroPoint pulsed laser, a turnkey system that is readily available and that has been widely used for targeted cell ablation. We describe aligning the laser, mounting the worms, cutting specific neurons, and assessing subsequent regeneration. The system provides the ability to cut large numbers of neurons in multiple worms during one experiment. Thus, laser axotomy as described herein is an efficient system for initiating and analyzing the process of regeneration.

In vivo Laser Axotomy in C. elegans

A protocol to cut neurons in C. elegans with a MicroPoint pulsed laser is presented. We describe setting up the system, immobilizing worms, and severing labeled neurons. Advantages include a relatively low-cost system and the ability to sever neuronal processes or ablate cells in vivo.

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Construction of Cell-based Neurotransmitter Fluorescent Engineered Reporters (CNiFERs) for Optical Detection of Neurotransmitters In Vivo

Cell-based neurotransmitter fluorescent engineered reporters (CNiFERs) provide a new tool for neuroscientists to optically detect the release of neurotransmitters in the brain in vivo. A specific CNiFER is created from a human embryonic kidney cell that stably expresses a specific G protein-coupled receptor, which couples to Gq/11 G proteins, and a FRET-based Ca2+-detector, TN-XXL. Activation of the receptor leads to an increase in the FRET signal. CNiFERs have nM sensitivity and a temporal response of seconds because a CNiFER clone utilizes the native receptor for a particular neurotransmitter, e.g., D2R for dopamine. CNiFERs are directly implanted into the brain, enabling them to sense neurotransmitter release with a spatial resolution of less than one hundred &#181;m, making them ideal to measure volume transmission in vivo. CNiFERs can also be used to screen other drugs for potential cross-reactivity in vivo. We recently expanded the family of CNiFERs to include GPCRs that couple to Gi/o G proteins. CNiFERs are available for detecting acetylcholine (ACh), dopamine (DA) and norepinephrine (NE). Given that any GPCR can be used to create a novel CNiFER and that there are approximately 800 GPCRs in the human genome, we describe here the general procedure to design, realize, and test any type of CNiFER.

Construction of Cell-based Neurotransmitter Fluorescent Engineered Reporters CNiFERs for Optical Detection of Neurotransmitters In Vivo

We present a protocol to create cell-based neurotransmitter fluorescent engineered reporters (CNiFERs) for the optical detection of volumetric neurotransmitter release.