We thank Aylin Rodan for critical reading of the manuscript, and members of the lab for helpful discussion. This research is supported by the NIH, R01AA019526.

1. Creating the exposure chamber
<ol>
<li>Cut a vial plug in half using a razor blade. Alternatively, use cotton balls.</li>
<li>Push the vial plug half, or the cotton ball, to the bottom of a food vial to make a smooth surface that flies will not get stuck in.</li>
<li>If making multiple exposure chambers, make sure that the vial plug and food take up the same volume in each chamber. A mark on the side of the vial can facilitate this.</li>
</ol>
2. Exposing the flies
<ol>
<li>Collect 8 flies of one sex, age 1-5 days, the day before exposure into regular food vials without yeast.</li>
<li>Label them in code, such that the experimenter is blinded to the actual genotype/strain used.</li>
<li>Transfer the eight flies into the exposure chambers by tapping. Do not anesthetize.</li>
<li>Add a small amount of food dye, e.g. acid blue 9, to 200 proof ethanol to make it easily visible.</li>
<li>Coat the bottom of a vial plug, or cotton ball, with 0.5 ml ethanol.</li>
<li>Insert the ethanol plug into the exposure vial. Ensure that the alcohol is facing into the vial, and not towards the wall of the vial.</li>
<li>Start a timer counting up.</li>
</ol>
3. Record ethanol-induced intoxication
<ol>
<li>Each minute after the timer is started, tap the exposure chamber on the bench to startle the flies and observe them for ten seconds.</li>
<li>Record the number of stationary flies for each minute. Flies are stationary if they cannot get off their backs, remain in one location, or rapidly vibrate their wings throughout the entire ten-second observation-period. The latter flies are considered stationary because the vast majority of these flies are stationary between periods of rapid wing movement. Flies are NOT stationary if they can get off their backs and walk away and/or up the vial wall.</li>
<li>An exposure trial is finished when 4 or more flies are recorded as stationary.</li>
<li>Discard flies and let the exposure chamber air out for later reuse.</li>
</ol>
4. Determine the time to 50% sedation

The time to 50% sedation (ST50) is the time after the start of the ethanol exposure when four of the eight flies remain stationary. This is determined as follows: 
Either: the first measurement minute when exactly four flies are stationary. 
Or: if less than four flies (Y1) are stationary at one time point (minute X), and more than four (Y2) at the next measuring time point (minute X+1), then linearly interpolate the ST50 using the formula: 
ST50 = (4 - Y1)/(Y2 - Y1) + X


5. Repeat exposure to determine tolerance

Exposing the flies
<ol>
<li>Determine the ST50(wt) for the wild-type control strain used in your experiment.</li>
<li>Expose the experimental groups of flies to ethanol as described above.</li>
<li>Measure their ST50(1), but continue to expose until you reach the time 2x ST50(wt).</li>
<li>Transfer the flies back into an empty food vial, and hold for 4 hours.</li>
<li>Re-expose the flies, and determine the ST50(2) of exposure 2.</li>
</ol>
Determine tolerance
Calculate tolerance in % as (ST50(2)/ ST50(1) - 1)x100.

6. Representative Results:
Shown here, are some representative results we obtained with 6-8 vials of flies per genotype and dose. If your standard error is substantially larger, then a systematic problem likely occurred. Make sure that the ethanol faces into the exposure chambers, and that the chambers are the same volume.
 
<img src="/files/ftp_upload/2541/2541fig1.jpg" alt="Figure 1" /> 
Figure 1. Dose response curve of wild type flies exposed to increasing percentages of ethanol. Eight vials of flies were exposed per dose and sex. Male and female flies show the same ST50 at each dose (p>0.22, 2-way ANOVA), while the ST50 across doses decreases significantly with increasing ethanol dose (p&lt;0.001). Note that flies from over-crowded bottles can be substantially smaller than those from healthy bottles, and this size difference can affect the ST50. Seed each bottle with 15-20 females laying for 1-2 days to get optimal larval density per bottle.
<img src="/files/ftp_upload/2541/2541fig2.jpg" alt="Figure 2" /> 
Figure 2. Exposing flies to either 90%, or 100% ethanol reproduces the previously described ethanol-resistance caused by the white rabbit1 mutation2 (p&lt;0.001, 2-way ANOVA).
<img src="/files/ftp_upload/2541/2541fig3.jpg" alt="Figure 3" /> 
Figure 3. Repeat exposure to 100% ethanol, spaced by a 4 hour recovery interval, causes a significant increase in ST50 after the second exposure (p&lt;0.001, paired t-test). Note that the flies were exposed for 24 minutes during the first exposure (corresponding to twice their na ve ST50). Exposure for a shorter amount of time (12 minutes) did not cause a significant increase in ST50 upon re-exposure (data not shown).

<ol>
	<li>Bainton, R. J., Tsai, L. T. -. Y., Singh, C. S., Moore, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Curr. Biol. 10, 187-194 (2000).</li><li>Rothenfluh, A., Threlkeld, R. J., Bainton, R. J., Tsai, L. T. -. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Cell. 127, 199-211 (2006).</li><li>Devineni, A. V., Heberlein, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Curr. Biol. 19, 2126-2132 (2009).</li><li>Scholz, H., Ramond, J., Singh, C. S., Heberlein, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Neuron. 28, 261-271 (2000).</li><li>Allen, M. J., Godenschwege, T. A., Tanouye, M. A., Phelan, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Semin. Cell Dev. Biol. 17, 31-41 (2006).</li><li>Feany, M. B., Bender, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Nature. 404, 394-398 (2000).</li></ol>

No conflicts of interest declared.

In this presentation, we have outlined a simple way to determine ethanol-induced intoxication based on previous assays described by Bainton et al. 20001, and Rothenfluh et al. 20062. Since alcohol-induced sedation is correlated with alcohol drinking preference3, this assay is relevant for the study of alcohol use disorders. The assay described is based on startle-induced negative geotaxis, and how an alcohol exposure interferes with this innate response. Before starting the exposures, make sure all your flies show a normal response to such a startle, i.e. after tapping down the flies in the food vial, they should all move up from the bottom within a 10 second observation time. Examples of flies having reduced negative geotaxis include bang-sensitive seizure mutants5, and locomotion-deficient flies caused by degeneration of dopamine neurons6.
Even though the described assay is relatively simple, two variables merit the most attention to ensure reproducible results. First, all exposure vials should be the same volume. This includes making sure that the bottom of the vial contains the same volume of food + vial plug/cotton ball, as well as inserting the ethanol-containing plug the same distance for every vial. A completely empty vial can be used for this assay, but it leads to longer ST50 values, decreasing the throughput of the assay.
This assay is also suitable to determine the amount of tolerance flies develop upon repeat ethanol exposure. A number of variables should be considered when doing repeat exposures. First, the inter-exposure interval is here described as 4 hours. This variable can be changed, but we recommend it not be lower than 2 hours, to ensure clearance of ethanol from the first exposure. See Scholz et al. 20004 for the kinetics of ethanol tolerance development and decay. The second variable to consider is the duration of the first exposure. This obviously has to be the same for the control and the experimental flies. However, this may not result in the same amount of ethanol delivered to both these fly strains. Sedated flies absorb more ethanol than active flies2, therefore a strain that sedates earlier may receive a higher dose than a later-sedating strain, and thus develop more tolerance due to a higher initial dose. We strongly caution the reader when comparing tolerance development of strains that show differences in the initial ST50(1).
The assay we describe here is a useful tool in easily and reproducibly determining flies behavioral responses to alcohol.

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a simple way to measure ethanol sensitivity in flies

Low doses of ethanol cause flies to become hyperactive, while high doses are sedating. The sensitivity to ethanol-induced sedation of a given fly strain is correlated with that strain s ethanol preference, and therefore sedation is a highly relevant measure to study the genetics of alcohol responses and drinking. We demonstrate a simple way to expose flies to ethanol and measure its intoxicating effects. The assay we describe can determine acute sensitivity, as well as ethanol tolerance induced by repeat exposure. It does not require a technically involved setup, and can therefore be applied in any laboratory with basic fly culture tools.

Watch this Scientific Journal Video about A Simple Way to Measure Ethanol Sensitivity in Flies at JoVE.com

A Simple Way to Measure Ethanol Sensitivity in Flies

A simple assay to measure the sedating effects of ethanol on Drosophila flies, based on the loss of righting reflex, is described.

Low doses of ethanol cause flies to become hyperactive, while high doses are sedating. The sensitivity to ethanol-induced sedation of a given fly strain ...

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23.2K vistas.  University of Texas Southwestern Medical Center. Una prueba sencilla para medir los efectos sedantes del alcohol en Drosophila Moscas, basado en la pérdida de los reflejos, se describe.

Video: A Simple Way to Measure Ethanol Sensitivity in Flies

Hyponeophagia: A Measure of Anxiety in the Mouse

Before the present day, when fast-acting and potent rodenticides such as alpha-chloralose were not yet in use, the work of pest controllers was often hampered by a phenomenon known as "bait shyness". Mice and rats cannot vomit, due to the tightness of the cardiac sphincter of the stomach, so to overcome the problem of potential food toxicity they have evolved a strategy of first ingesting only very small amounts of novel substances. The amounts ingested then gradually increase until the animal has determined whether the substance is safe and nutritious. So the old rat-catchers would first put a palatable substance such as oatmeal, which was to be the vehicle for the toxin, in the infested area. Only when large amounts were being readily consumed would they then add the poison, in amounts calculated not to affect the taste of the vehicle. The poisoned bait, which the animals were now readily eating in large amounts, would then swiftly perform its function.
Bait shyness is now used in the behavioural laboratory as a way of measuring anxiety. A highly palatable but novel substance, such as sweet corn, nuts or sweetened condensed milk, is offered to the mice (or rats) in a novel situation, such as a new cage. The latency to consume a defined amount of the new food is then measured.
Robert M.J. Deacon can be reach at <a href="mailto:robert.deacon@psy.ox.ac.uk">robert.deacon@psy.ox.ac.uk</a>

Mice and rats, due to their innate cautiousness, are initially slow in consuming a novel food, particularly in a novel place. This hyponeophagia can readily be measured in the laboratory, even though laboratory animals are much less anxious than their wild counterparts

Before the present day, when fast-acting and potent rodenticides such as alpha-chloralose were not yet in use, the work of pest controllers was often ...

Microdialysis of Ethanol During Operant Ethanol Self-administration and Ethanol Determination by Gas Chromatography

Operant self-administration methods are commonly used to study the behavioral and pharmacological effects of many drugs of abuse, including ethanol. However, ethanol is typically self-administered orally, rather than intravenously like many other drugs of abuse. The pharmacokinetics of orally administered drugs are more complex than intravenously administered drugs. Because understanding the relationship between the pharmacological and behavioral effects of ethanol requires knowledge of the time course of ethanol reaching the brain during and after drinking, we use in vivo microdialysis and gas chromatography with flame ionization detection to monitor brain dialysate ethanol concentrations over time. 
Combined microdialysis-behavioral experiments involve the use of several techniques. In this article, stereotaxic surgery, behavioral training and microdialysis, which can be adapted to test a multitude of self-administration and neurochemical centered hypotheses, are included only to illustrate how they relate to the subsequent phases of sample collection and dialysate ethanol analysis. Dialysate ethanol concentration analysis via gas chromatography with flame-ionization detection, which is specific to ethanol studies, is described in detail. Data produced by these methods reveal the pattern of ethanol reaching the brain during the self-administration procedure, and when paired with neurochemical analysis of the same dialysate samples, allows conclusions to be made regarding the pharmacological and behavioral effects of ethanol.

A method to determine the time course of ethanol concentration in the brains of rats during operant ethanol self-administration is described. Gas chromatography with flame ionization detection is used to quantify ethanol in the dialysate samples, because it has the sensitivity required for the small volumes that are generated.

Operant self-administration methods are commonly used to study the behavioral and pharmacological effects of many drugs of abuse, including ethanol.  ...

The FlyBar: Administering Alcohol to Flies

Fruit flies (Drosophila melanogaster) are an established model for both alcohol research and circadian biology. Recently, we showed that the circadian clock modulates alcohol sensitivity, but not the formation of tolerance. Here, we describe our protocol in detail. Alcohol is administered to the flies using the FlyBar. In this setup, saturated alcohol vapor is mixed with humidified air in set proportions, and administered to the flies in four tubes simultaneously. Flies are reared under standardized conditions in order to minimize variation between the replicates. Three-day old flies of different genotypes or treatments are used for the experiments, preferably by matching flies of two different time points (e.g., CT 5 and CT 17) making direct comparisons possible. During the experiment, flies are exposed for 1 hr&#160;to the pre-determined percentage of alcohol vapor and the number of flies that exhibit the Loss of Righting reflex (LoRR) or sedation are counted every 5 min. The data can be analyzed using three different statistical approaches. The first is to determine the time at which 50% of the flies have lost their righting reflex and use an Analysis of the Variance (ANOVA) to determine whether significant differences exist between time points. The second is to determine the percentage flies that show LoRR after a specified number of minutes, followed by an ANOVA analysis. The last method is to analyze the whole times series using multivariate statistics. The protocol can also be used for non-circadian experiments or comparisons between genotypes.

Drosophila has emerged as a significant model system for dissecting the cellular and molecular underpinnings of behavioral responses to alcohol. Here we present a protocol for the collection of alcohol sensitivity data in a circadian context that can be easily applied to other experiments and is well-suited for undergraduate research.

Fruit flies (Drosophila melanogaster) are an established model for both alcohol research and circadian biology. Recently, we showed that the circadian ...

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion channel, and transporter cell surface presentation on a minutes time scale.&#160;There is a broad diversity of mechanisms that control endocytic trafficking of individual proteins. Studies investigating the molecular underpinnings of trafficking have primarily relied upon surface biotinylation to quantitatively measure changes in membrane protein surface expression in response to exogenous stimuli and gene manipulation.&#160;However, this approach has been mainly limited to cultured cells, which may not faithfully reflect the physiologically relevant mechanisms at play in adult neurons.&#160;Moreover, cultured cell approaches may underestimate region-specific differences in trafficking mechanisms.&#160;Here, we describe an approach that extends cell surface biotinylation to the acute brain slice preparation.&#160;We demonstrate that this method provides a high-fidelity approach to measure rapid changes in membrane protein surface levels in adult neurons.&#160;This approach is likely to have broad utility in the field of neuronal endocytic trafficking.

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Neuronal membrane trafficking dynamically controls plasma membrane protein availability and significantly impacts neurotransmission.&#160;To date, it has been challenging to measure neuronal endocytic trafficking in adult neurons.&#160;Here, we describe a highly effective, quantitative method to measure rapid changes in surface protein expression ex vivo in acute brain slices.

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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.

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An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Alcohol use disorder (AUD) is a serious health challenge. Despite a large hereditary component to AUD, few genes have been unambiguously implicated in their etiology. The fruit fly, Drosophila melanogaster, is a powerful model for exploring molecular-genetic mechanisms underlying alcohol-related behaviors and therefore holds great promise for identifying and understanding the function of genes that influence AUD. The use of the Drosophila model for these types of studies depends on the availability of assays that reliably measure behavioral responses to ethanol. This report describes an assay suitable for assessing ethanol sensitivity and rapid tolerance in flies. Ethanol sensitivity measured in this assay is influenced by the volume and concentration of ethanol used, a variety of previously reported genetic manipulations, and also the length of time the flies are housed without food immediately prior to testing. In contrast, ethanol sensitivity measured in this assay is not affected by the vigor of fly handling, sex of the flies, and supplementation of growth medium with antibiotics or live yeast. Three different methods for quantitating ethanol sensitivity are described, all leading to essentially indistinguishable ethanol sensitivity results. The scalable nature of this assay, combined with its overall simplicity to set-up and relatively low expense, make it suitable for small and large scale genetic analysis of ethanol sensitivity and rapid tolerance in Drosophila.

An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Straightforward assays for measuring ethanol sensitivity and rapid tolerance in Drosophila facilitate the use of this model organism for investigating these important ethanol-related behaviors. Here, a relatively simple, scalable assay for measuring ethanol sensitivity and rapid tolerance in flies is described.

Alcohol use disorder (AUD) is a serious health challenge. Despite a large hereditary component to AUD, few genes have been unambiguously implicated in ...

A Simple Alternative to Stereotactic Injection for Brain Specific Knockdown of miRNA

MicroRNAs (miRNAs) are key regulators of gene expression. In the brain, vital processes like neurodevelopment and neuronal functions depend on the correct expression of microRNAs. Perturbation of microRNAs in the brain can be used to model neurodegenerative diseases by modulating neuronal cell death. Currently, stereotactic injection is used to deliver miRNA knockdown agents to specific location in the brain. Here, we discuss strategies to design antagomirs against miRNA with locked nucleotide modifications (LNA). Subsequently describe a method for brain specific delivery of antagomirs, uniformly across different regions of the brain. This method is simple and widely applicable since it overcomes the surgery, associated injury and limitation of local delivery in stereotactic injections. We prepared a complex of neurotropic, cell-penetrating peptide Rabies Virus Glycoprotein (RVG) with antagomir against miRNA-29 and injected through tail vein, to specifically deliver in the brain. The antagomir design incorporated features that allow specific targeting of the miRNA and formation of non-covalent complexes with the peptide. The knock-down of the miRNA in neuronal cells, resulted in apoptotic cell death and associated behavioural defects. Thus, the method can be used for acute models of neuro-degeneration through the perturbation of miRNAs.

MicroRNAs play crucial roles in the brain and are potential targets for modeling neuro-degeneration. However, perturbing miRNA levels is challenging due to the short length of miRNA and inaccessibility of the brain tissue. This video presents a method for antagomir design and brain specific delivery using a neuropeptide in mice.

MicroRNAs (miRNAs) are key regulators of gene expression.  In the brain, vital processes like neurodevelopment and neuronal functions depend on the ...

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We demonstrate a simple way to tap into the fly reward system, modify experiences related to natural reward, and use voluntary ethanol consumption as a measure for changes in reward states. The approach serves as a relevant tool to study the neurons and genes that play a role in experience-mediated changes of internal state. The protocol is composed of two discrete parts: exposing the flies to rewarding and nonrewarding experiences, and assaying voluntary ethanol consumption as a measure of the motivation to obtain a drug reward. The two parts can be used independently to induce the modulation of experience as an initial step for further downstream assays or as an independent two-choice feeding assay, respectively. The protocol does not require a complicated setup and can therefore be applied in any laboratory with basic fly culture tools.

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for inducing rewarding and nonrewarding experiences in fruit flies (Drosophila melanogaster) using voluntary ethanol consumption as a measure for changes in reward states.

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A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Using Enzyme-based Biosensors to Measure Tonic and Phasic Glutamate in Alzheimer's Mouse Models

Neurotransmitter disruption is often a key component of diseases of the central nervous system (CNS), playing a role in the pathology underlying Alzheimer's disease, Parkinson's disease, depression, and anxiety. Traditionally, microdialysis has been the most common (lauded) technique to examine neurotransmitter changes that occur in these disorders. But because microdialysis has the ability to measure slow 1-20 minute changes across large areas of tissue, it has the disadvantage of invasiveness, potentially destroying intrinsic connections within the brain and a slow sampling capability. A relatively newer technique, the microelectrode array (MEA), has numerous advantages for measuring specific neurotransmitter changes within discrete brain regions as they occur, making for a spatially and temporally precise approach. In addition, using MEAs is minimally invasive, allowing for measurement of neurotransmitter alterations in vivo. In our laboratory, we have been specifically interested in changes in the neurotransmitter, glutamate, related to Alzheimer's disease pathology. As such, the method described here has been used to assess potential hippocampal disruptions in glutamate in a transgenic mouse model of Alzheimer's disease. Briefly, the method used involves coating a multi-site microelectrode with an enzyme very selective for the neurotransmitter of interest and using self-referencing sites to subtract out background noise and interferents. After plating and calibration, the MEA can be constructed with a micropipette and lowered into the brain region of interest using a stereotaxic device. Here, the method described involves anesthetizing rTg(TauP301L)4510 mice and using a stereotaxic device to precisely target sub-regions (DG, CA1, and CA3) of the hippocampus.

Using Enzyme-based Biosensors to Measure Tonic and Phasic Glutamate in Alzheimer&#039;s Mouse Models

Here, we describe the setup, software navigation, and data analysis for a spatially and temporally precise method of measuring tonic and phasic extracellular glutamate changes in vivo using enzyme-linked microelectrode arrays (MEA).