This work was supported by the National Institute of Neurological Disorders and Stroke Intramural Research Program (NS002986). AS wrote the manuscript and implemented the deconvolution analysis. JSD developed the initial version of the deconvolution analysis and commented on the text.

1. Slice Preparation
<ol>
	<li>Prepare 500 ml slicing/storage solution containing (in mM): 119 NaCl, 2.5 KCl, 0.5 CaCl2, 1.3 MgSO4&middot;7H2O, 4 MgCl2, 26.2 NaHCO3, 1 NaH2PO4, and 22 glucose, 320 mOsm, pH 7.4</li>
	<li>Use a 250 ml beaker to prepare a submersion chamber for the slices, fill it with 200 ml slicing/storage solution, warm it in a water bath at 34 &deg;C and bubble it with 95% O2, 5% CO2.</li>
	<li>Keep the remaining slicing/storage solution in a glass bottle at 4 &deg;C.</li>
	<li>Use a cyanoacrylate adhesive to attach a small agar block (6%, prepared in ACSF) on the vibratome sample holder and store it at 4 &deg;C.</li>
	<li>30 min before starting slicing, place the glass bottle containing the slicing/storage solution in a bucket filled with ice and bubble it with 95% O2, 5% CO2.</li>
</ol>
Note: speed and precision are paramount for the dissection steps described below.
<ol start="6">
	<li>Anesthetize the mouse (P14-21, C57BL/6) with halothane/isoflurane (isoflurane has been reported to enhance astrocytic glutamate uptake15), decapitate it, and dip the head in a 50 ml beaker containing oxygenated, cold slicing/storage solution.</li>
	<li>Perform the dissection of the brain on an ice pack wrapped with paper towels and stored at -20 &deg;C.</li>
	<li>Use a scalpel to make a mid-sagittal skin incision on the dorsal side of the head, from the frontal to the caudal end, and expose the skull.</li>
	<li>Place the lower shear blade of a small pair of surgical scissors in the occipital hole and make two cuts, towards the left and on the right hand side (45&deg; angle).</li>
	<li>Cut the skull along the mid-sagittal line, from the caudal to the frontal end.</li>
	<li>Remove the brain with a spatula and dip it in oxygenated, cold slicing/storage solution.</li>
	<li>With a scalpel, make five cuts to: (1) remove the olfactory bulbs and the frontal cortex; (2) remove the cerebellum; (3) remove the left and (4) right temporal lobes; (5) separate the two brain hemispheres with a mid-sagittal cut.</li>
	<li>Dab the two parts of the brain with paper towels to remove any excess solution.</li>
	<li>Glue the lateral surface of each brain section to the cold vibratome base plate: the dorsal side of the brain should be facing the vibratome blade; the ventral side of the brain should be in contact with the agar block, away from the vibratome blade. 
	Note: to obtain the highest-quality slices, it is important that the two parts of the brain are firmly glued to the vibratome base plate. To do this, use a cyanoacrylate adhesive that is not too liquid and that does not dry out too quickly.</li>
	<li>Secure the vibratome base plate to the dissecting chamber, set the slice thickness to 250 &mu;m and adjust the width of the blade run. Proceed with slicing. 
	Note: if everything is oriented correctly, the vibratome should be cutting parasagittal slices from the dorsal towards the ventral and from the medial towards the lateral side of the brain.</li>
	<li>Once the blade has passed through the cortex and hippocampus, use a scalpel to cut the cortex/hippocampus from the midbrain and place each slice in the submersion chamber at 34 &deg;C.</li>
	<li>Discard the first couple of slices. Typically, 12 slices (250 &mu;m thick) can be obtained from a P14-21 mouse brain.</li>
	<li>Keep the slices at 34 &deg;C for 30 min and let them cool down to room temperature for 30 min before using them for the electrophysiology recordings.</li>
</ol>
2. Astrocyte Identification and Recordings
<ol>
	<li>Prepare an internal solution containing (in mM): 120 KCH3SO3, 10 EGTA, 20 HEPES, 2 MgATP, 0.2 NaGTP, 5 QX-314Br, and 5 NaCl, 290 mOsm, pH 7.2.</li>
	<li>Prepare an extracellular recording solution containing (in mM): 119 NaCl, 2.5 KCl, 2.5 CaCl2, 1.3 MgSO4&middot;7H2O, 1 MgCl2, 26.2 NaHCO3, 1 NaHPO4, and 22 glucose, 300 mOsm, pH 7.4, saturated with 95% O2, 5% CO2.</li>
	<li>Add the following drugs to the extracellular recording solution, to block activation of GluA, GluN, mGluRII, mGluRIII, GABAA, GABAB, and A1 adenosine receptors (in &mu;M): 10 2,3-Dioxo-6-nitro-1,2,3,4-tetrahydrobenzo[&fnof;]quinoxaline-7-sulfonamide disodium salt (NBQX), 10 (RS)-3-(2-carboxypiperazin-4-yl)-propyl-1-phosphonic acid (CPP), (2S)-2-Amino-2-[(1S,2S)-2-carboxycycloprop-1-yl]-3-(xanth-9-yl) propanoic acid disodium salt (LY341495), 100 (R,S)-&alpha;-methylserine-O-phosphate (MSOP), 100 picrotoxin, 5 3-[[(3,4-Dichlorophenyl)methyl]amino]propyl] diethoxymethyl)phosphinic acid (CGP52432), 1 8-cyclopentyl-1,3-dipropylxanthine (DPCPX).</li>
	<li>Set the temperature of the extracellular solution in the recording chamber; typical temperatures are between 34 - 36 &deg;C.</li>
	<li>Prepare patch-clamp electrodes from borosilicate glass capillaries (R &asymp; 2.5 M&Omega;) using a dual-stage, glass micro-pipette puller.</li>
	<li>Take one of the slices and place it in the recording chamber. Hold it down with a metal harp made with platinum wire and nylon strings.</li>
	<li>Visually inspect the slices under Dodt illumination or IR-DIC. Astrocytes can be identified by their small cell body (&Oslash; = 10 &mu;m) and prominent nucleus (Figure 1).</li>
	<li>For synaptic stimulations, place a bipolar stainless steel electrode, ~100 &mu;m away from the astrocyte that you plan to patch.</li>
	<li>Patch the astrocyte and break in the whole-cell configuration by applying a very gentle suction. 
	Note: Astrocytes typically have low input resistance (~10 M&Omega;), hyperpolarized resting membrane potential (~-90 mV), and no firing activity. The astrocytes are kept at its resting membrane potential throughout the experiments in voltage-clamp mode (i.e. the holding current should read 0 pA). Before each stimulation, a 10 ms hyperpolarizing voltage step (-3 mV) is used to monitor the series and input resistance of the astrocyte.</li>
	<li>Discard the recordings if the series resistance changes &gt;20% or if the membrane potential of the astrocyte becomes depolarized. The membrane potential of the astrocyte can be directly measured by switching from voltage-clamp to current-clamp mode. Alternatively, while in voltage-clamp mode, the membrane potential can be monitored by reading the value of the holding potential that results in 0 pA holding current.</li>
	<li>If synaptic stimulations are employed, alternate single and paired stimuli (e.g. 100 msec apart), every 10 - 20 sec.</li>
	<li>For UV photolysis experiments, connect an uncaging Xe lamp to the epifluorescence port of the microscope and add the caged compound to the extracellular solution. Transporter currents of ~100 pA amplitude can be obtained when flash-uncaging 100 &mu;M MNI-L-glutamate in the whole field of view (&Oslash; = 662.5 &mu;m when using a 40X objective 14). Alternate stimulations with the light path open and blocked, every 10 - 20 sec.</li>
	<li>Evoke glutamate transients and record transporter currents in control conditions and in the presence of a sub-saturating concentration of the broad-spectrum glutamate transporter antagonist D,L-threo-&beta;-Benzyloxyaspartic acid (TBOA; 10 &mu;M) to reduce the transporter current amplitude to at least 30% of its control value (see sections 4, 5) or in the presence of a high concentration of TBOA (50 - 100 &mu;M) to block transporter currents completely and isolate the sustained K+-current (see section 3).</li>
</ol>
3. Pharmacological Isolation of the Sustained K+-current
<ol>
	<li>Record astrocytic currents in control conditions and in the presence of a high, saturating concentration of TBOA (50 - 100 &mu;M).</li>
	<li>Average at least 20 sweeps in TBOA (50 - 100 &mu;M).</li>
	<li>Fit the averaged traces obtained in 3.2 with the function: 
	<img alt="Equation 1" fo:src="/files/ftp_upload/50708/50708eq1highres.jpg" src="/files/ftp_upload/50708/50708eq1.jpg" /> 
	Note: the TBOA-insensitive component (i.e. the one that is recorded in the presence of TBOA (50 - 100 &mu;M)) represents the sustained K+-current. Its time course is approximated by the mono-exponential function described above.</li>
	<li>Repeat this fit across different astrocytes and derive an average value of &Tau;rise (i.e. the average rise time of the sustained K+-current (see section 5)).</li>
</ol>
4. Isolation of the Facilitated Portion of Synaptically-Activated Transporter Currents (fSTCs)
<ol>
	<li>Average at least 20 sweeps obtained with paired stimulations, in control conditions and in TBOA (10 &mu;M) (Figure 2a left).</li>
	<li>Average an identical number of sweeps obtained with single stimulations, in control conditions and in TBOA (10 &mu;M) (Figure 2a middle).</li>
	<li>Compare the amplitude of the average current response to the -3 mV voltage step in the four averaged traces (control single pulse, control paired-pulses, TBOA single pulse, TBOA paired-pulses).</li>
	<li>If the amplitude of the current response is the same in all four traces, proceed to step 4.7.</li>
	<li>If the amplitude of the average current response is different in any of the four traces, check that all individual traces were appropriate to be included in the average.</li>
	<li>If the amplitude of the average current is different in any of the four traces, but all individual traces are appropriate to be included in the average, verify that in each trace the size of the fSTC scales linearly with the size of the current response to the test voltage step. If this is the case, scale all traces with respect to each other so that the current responses to the test voltage step are all equal in amplitude. 
	Note: In the recording conditions described here (i.e. intracellular CH3SO3- or C6H11O7-), synaptic stimulations of excitatory axons generate astrocytic currents with complex waveforms consisting of a fast-rising transient inward stoichiometric current and a slow-rising, sustained inward K+-current reflecting K+ re-equilibration in the extracellular space following action potential propagation along neighboring axons. It is critical to remove this sustained K+-current, as any residual current would lead to an overestimate of glutamate lifetime.</li>
	<li>Subtract the average trace obtained with single stimulations from the average trace obtained with paired stimulations (Figure 2a right). This step allows isolating the stoichiometric and the sustained K+-current evoked by the second stimulus.</li>
	<li>Shift the average trace obtained with single stimulations by a time interval that matches the inter-pulse interval used to deliver paired-pulses (i.e. 100 msec) (Figure 2b) .</li>
	<li>Subtract the time-shifted average response obtained with single stimulations from the average response to the second stimulus obtained in 4.7 (Figure 2c). This step isolates the facilitated portion of the transporter current evoked by the second stimulus (the fSTC).</li>
	<li>In most cases, the previous step removes entirely the sustained K+-current. If this is the case, the isolation of the fSTC is complete and you can proceed with the deconvolution analysis and derive the time course of glutamate clearance from astrocytes. In some cases, however, a small sustained K+-current is still present (Figure 2d) and further analysis is required (see section 5). 
	Note: remember that the analysis described in 4.7-4.10 must be performed on the average traces obtained in control conditions and in TBOA (10 &mu;M).</li>
</ol>
5. Subtraction of the Residual zdustained K+-current from fSTCs
<ol>
	<li>Measure the amplitude of the sustained K+-current remaining after performing the analysis described in section 4.</li>
	<li>Scale the amplitude of the mono-exponential function described in 3.3 to the amplitude of the residual sustained K+-current measured in 5.1 (Figure 2d left and middle). To do this, in equation 3.3, set the term A equals to the amplitude of the sustained K+-current measured in 5.1 and the term &Tau;rise equals to the average value of &Tau;rise estimated in 3.4.</li>
	<li>Subtract the resulting mono-exponential function from the fSTC and sustained K+-current obtained in 4.10 (Figure 2d right). This step completes the isolation of the fSTC.</li>
</ol>
6. Isolation of Flash-activated Transporter Currents (FTCs)
<ol>
	<li>Average at least 20 sweeps obtained with the light path open, in control conditions and in TBOA (10 &mu;M).</li>
	<li>Average an identical number of sweeps obtained with the light path closed, in control conditions and in TBOA (10 &mu;M).</li>
	<li>Compare the amplitude of the current response to the -3 mV voltage step in the four averaged traces (control single pulse, control paired-pulses, TBOA single pulse, TBOA paired-pulses).</li>
	<li>If the amplitude of the current response is the same in all four traces, proceed to step 6.7.</li>
	<li>If the amplitude of the average current response is different in any of the four traces, check that all individual traces were appropriate to be included in the average.</li>
	<li>If the amplitude of the average current is different in any of the four traces, but all individual traces are appropriate to be included in the average, verify that in each trace with the light path open the size of the FTC scales linearly with the size of the current response to the test voltage step. If this is the case, scale all traces with respect to each other so that the current responses to the test voltage step are all equal in amplitude. 
	Note: in the recording conditions described here (i.e. intracellular CH3SO3- or C6H11O7-), flash stimuli generate astrocytic currents consisting only of the fast-rising transient stoichiometric inward current.</li>
	<li>Subtract the average trace obtained with the light path blocked from the average trace obtained with the light path open. This step allows removing the stimulus artifact and isolating the FTCs. 
	Note: the analysis described in 6.7 must be performed on the average traces obtained in control conditions and in TBOA (10 &mu;M).</li>
</ol>
7. Deconvolution analysis
<ol>
	<li>Fit the transporter current (fSTC or FTC) recorded in control conditions (Figure 3a left) and in TBOA (10 &mu;M) (Figure 3a right) and isolated as described in sections 4 - 6, with the multi-exponential function: 
	<img alt="Equation 2" fo:src="/files/ftp_upload/50708/50708eq2highres.jpg" src="/files/ftp_upload/50708/50708eq2.jpg" /></li>
	<li>Create an instantaneously rising function that decays mono-exponentially according to the function: 
	<img alt="Equation 3" fo:src="/files/ftp_upload/50708/50708eq3highres.jpg" src="/files/ftp_upload/50708/50708eq3.jpg" /> 
	and that best describes the decaying phase of the transporter current recorded in TBOA (10 &mu;M) (Figure 3b). 
	Note: the function described in 7.2 (i.e. H(t)) represents the time course of glutamate clearance from astrocytes in the presence of TBOA (10 &mu;M).</li>
	<li>Deconvolve the mono-exponential function obtained in 7.2 (Figure 3b and 3c middle) from the fit of the transporter current recorded in TBOA (10 &mu;M) obtained in 7.1 (Figure 3a right, c left). 
	Note: deconvolution is a mathematical operation included in many analysis software packages like (e.g. Matlab, Python). In IgorPro, the programming code that we typically use to perform this kind of analysis, the deconvolution operation can be efficiently computed as described below, by using discrete fast Fourier transforms. First, use the FFT operation to compute the discrete fast Fourier transform of the mono-exponential function obtained in 7.2 and of the fit of the transporter current recorded in TBOA obtained in 7.1. Next, use the IFFT operation to compute the inverse discrete fast Fourier transform of the ratio of the two FFT functions. The resulting function I(t) can be described as follows: 
	<img alt="Equation 4" fo:src="/files/ftp_upload/50708/50708eq4highres.jpg" src="/files/ftp_upload/50708/50708eq4.jpg" /> 
	The step described in 7.3 allows deriving the filter of the transporter current in TBOA (10 &mu;M) (Figure 3c right). The filter represents the distortion factors that convert the lifetime of glutamate at astrocytic membranes into the transporter current isolated in sections 4 - 6 (i.e. the fSTC or FTC).</li>
	<li>Deconvolve the filter (Figure 3c right, Figure 3d middle) from the fit of the transporter current in control conditions (Figure 3a left, Figure 3d left). 
	Note: this step allows deriving the time course of glutamate clearance from astrocytes in control conditions (Figure 3d right). The assumption underlying this deconvolution step is that the temporal profile of the filter remains unaltered in control conditions and in TBOA (10 &mu;M).</li>
	<li>To obtain a quantitative estimate of the overall time course of glutamate clearance, calculate the centroid (&lt;t&gt;) of the waveform obtained in step 7.4. This is done by calculating &lt;t&gt; as: 
	<img alt="Equation 5" fo:src="/files/ftp_upload/50708/50708eq5highres.jpg" src="/files/ftp_upload/50708/50708eq5.jpg" /> 
	where G(t) is the waveform obtained in step 7.1. In the equation described above, the term t corresponds to the time window over which the integral is calculated. 
	Note: when the method was first established, &lt;t&gt; was calculated over time windows that were left unconstrained 13 . This approach can be used as long as the integration window is set to be wider than the duration of glutamate clearance and the clearance waveform decays completely back to baseline. If this is not the case, however, this method of estimating &lt;t&gt; encounters some limitations. For example, if the clearance waveform does not decay exactly back to baseline, then &lt;t&gt; increases with the width of the integration window. To avoid any potential source of inaccuracy for the estimate of &lt;t&gt;, we now calculate it over a time window corresponding to 10% of the peak of the transporter current, before and after its onset 14 . The latter approach improves the consistency with which &lt;t&gt; is measured across cells. This comes in handy particularly when analyzing small effects of pharmacological treatments on the time course of clearance.</li>
</ol>

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The authors declare no conflict of interest.

Here we describe an experimental approach to obtain electrophysiological recordings from astrocytes, an analytical protocol to isolate glutamate transporter currents in astrocytes and a mathematical method to derive the time course of glutamate clearance from astrocytic transporter currents.
The success of the analysis relies on the ability to obtain high-quality patch clamp recordings from astrocytes and on the accuracy of the fitting algorithms used to describe the transporter currents. The ...

Astrocytes are one of the most abundant cell types in the brain with star-shaped morphology and fine membrane protrusions that extend throughout the neuropil and reach neighboring synaptic contacts 1,2. The astrocytes' cell membrane is densely packed with glutamate transporter molecules 3. Under physiological conditions, glutamate transporters rapidly bind glutamate at the extracellular side of the membrane and transfer it to the cell cytoplasm. By doing so, the transporters maintain low the basal concentration of glutamate in the extracellular space 4. Glutamate transporters in fine astrocytic processes adjacent to excitatory synapses are ideally positioned to bind glutamate released during synaptic events as it diffuses away from the synaptic cleft. By doing so, the transporters also limit glutamate spillover towards peri- and extra-synaptic regions and onto neighboring synapses, reducing the spatial spread of excitatory signals in the brain 5-7.
Glutamate transport is an electrogenic process stoichiometrically coupled to the movement of 3 Na+ and 1 H+ along their electrochemical gradient and to the counter-transport of 1 K+ 8. Glutamate transport is associated with (but not stoichiometrically coupled to) an anionic conductance permeable to SCN- (thiocyanate) &gt; NO3- (nitrate) &asymp; ClO4- (perchlorate) &gt; I- &gt; Br- &gt; Cl- &gt; F-, not to CH3SO3- (methane sulfonate) and C6H11O7- (gluconate) 9-11. Both currents (stoichiometric and non-stoichiometric) can be recorded by obtaining whole-cell patch-clamp recordings from astrocytes, visually identified under Dodt illumination or infra-red differential interference contrast (IR-DIC) in acute brain slices 12. The stoichiometric component of the current associated with glutamate transport across the membrane can be isolated by using CH3SO3-, or C6H11O7- based intracellular solutions and can be evoked by flash-uncaging glutamate on astrocytes 13,14, or by activating glutamate release from neighboring synapses, either electrically 12 or with a targeted optogenetic control.
The time course of the stoichiometric component of the transporter current is shaped by the lifetime of the glutamate concentration profile at astrocytic membranes (i.e. glutamate clearance), the kinetics of glutamate transporters, the passive membrane properties of astrocytes, and during synaptic stimulations, by the synchronicity of glutamate release across the activated synapses 13. Here we describe in full detail: (1) an experimental approach to isolate the stoichiometric component of glutamate transporter currents from whole-cell patch-clamp recordings from astrocytes using acute mouse hippocampal slices as an example experimental preparation; (2) an analytical approach to derive the time course of glutamate clearance from these recordings 13,14. These methods can be used to record and analyze glutamate transporter currents from astrocytes in any region of the central nervous system.

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deriving the time course of glutamate clearance with a deconvolution analysis of astrocytic transporter currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

The success of the analytical approach described here critically depends on obtaining high-quality electrophysiological recordings of transporter currents from astrocytes in any region of the central nervous system. In acute mouse hippocampal slices, astrocytes can be readily identified under Dodt illumination or IR-DIC because of their small cell body (&Oslash; = 10 &mu;m) and prominent nucleus (Figure 1). Their distinctive star-shaped morphology can be appreciated with epifluorescence, confocal, or two...

Watch this Scientific Journal Video about Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents at JoVE.com

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

deriving-time-course-glutamate-clearance-with-deconvolution-analysis

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JoVE Journal

Neuroscience

10.3K Views.  National Institutes of Health. We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

Video: Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

The marine gastropod mollusk Aplysia californica has a venerable history as a model of nervous system function, with particular significance in studies of learning and memory. The typical preparations for such studies are ones in which the sensory and motoneurons are left intact in a minimally dissected animal, or a technically elaborate neuronal co-culture of individual sensory and motoneurons. Less common is the isolated neuronal preparation in which small clusters of nominally homogeneous neurons are dissociated into single cells in short term culture. Such isolated cells are useful for the biophysical characterization of ion currents using patch clamp techniques, and targeted modulation of these conductances. A protocol for preparing such cultures is described. The protocol takes advantage of the easily identifiable glutamatergic sensory neurons of the pleural and buccal ganglia, and describes their dissociation and minimal maintenance in culture for several days without serum.

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

We describe the dissection of the nervous system of the marine sea hare Aplysia after anesthesia, the isolation of neurons for short term-tissue culture, and recordings of single cell ion currents via the patch clamp technique.

The marine gastropod mollusk Aplysia californica has a venerable  history as a model of nervous system function, with particular significance in  studies ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Multi-photon Intracellular Sodium Imaging Combined with UV-mediated Focal Uncaging of Glutamate in CA1 Pyramidal Neurons

Multi-photon fluorescence microscopy has enabled the analysis of morphological and physiological parameters of brain cells in the intact tissue with high spatial and temporal resolution. Combined with electrophysiology, it is widely used to study activity-related calcium signals in small subcellular compartments such as dendrites and dendritic spines. In addition to calcium transients, synaptic activity also induces postsynaptic sodium signals, the properties of which are only marginally understood. Here, we describe a method for combined whole-cell patch-clamp and multi-photon sodium imaging in cellular micro domains of central neurons. Furthermore, we introduce a modified procedure for ultra-violet (UV)-light-induced uncaging of glutamate, which allows reliable and focal activation of glutamate receptors in the tissue. To this end, whole-cell recordings were performed on Cornu Ammonis subdivision 1 (CA1) pyramidal neurons in acute tissue slices of the mouse hippocampus. Neurons were filled with the sodium-sensitive fluorescent dye SBFI through the patch-pipette, and multi-photon excitation of SBFI enabled the visualization of dendrites and adjacent spines. To establish UV-induced focal uncaging, several parameters including light intensity, volume affected by the UV uncaging beam, positioning of the beam as well as concentration of the caged compound were tested and optimized. Our results show that local perfusion with caged glutamate (MNI-Glutamate) and its focal UV-uncaging result in inward currents and sodium transients in dendrites and spines. Time course and amplitude of both inward currents and sodium signals correlate with the duration of the uncaging pulse. Furthermore, our results show that intracellular sodium signals are blocked in the presence of blockers for ionotropic glutamate receptors, demonstrating that they are mediated by sodium influx though this pathway. In summary, our method provides a reliable tool for the investigation of intracellular sodium signals induced by focal receptor activation in intact brain tissue.

We describe the combination of focal UV-induced photo-activation of neuro-active compounds with whole-cell patch-clamp and multi-photon imaging of intracellular sodium transients in dendrites and spines of hippocampal neurons in acute tissue slices of the mouse brain.

Multi-photon fluorescence microscopy has enabled the analysis of morphological and physiological parameters of brain cells in the intact tissue with high ...

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma establishes a lifelong persistent infection in the brain. While this brain infection is asymptomatic in most immunocompetent people, in the developing fetus or immunocompromised individuals such as acquired immune deficiency syndrome (AIDS) patients, this predilection for and persistence in the brain can lead to devastating neurologic disease. Thus, it is clear that the brain-Toxoplasma interaction is critical to the symptomatic disease produced by Toxoplasma, yet we have little understanding of the cellular or molecular interaction between cells of the central nervous system (CNS) and the parasite. In the mouse model of CNS toxoplasmosis it has been known for over 30 years that neurons are the cells in which the parasite persists, but little information is available about which part of the neuron is generally infected (soma, dendrite, axon) and if this cellular relationship changes between strains. In part, this lack is secondary to the difficulty of imaging and visualizing whole infected neurons from an animal. Such images would typically require serial sectioning and stitching of tissue imaged by electron microscopy or confocal microscopy after immunostaining. By combining several techniques, the method described here enables the use of thick sections (160 &#181;m) to identify and image whole cells that contain cysts, allowing three-dimensional visualization and analysis of individual, chronically infected neurons without the need for immunostaining, electron microscopy, or serial sectioning and stitching. Using this technique, we can begin to understand the cellular relationship between the parasite and the infected neuron.

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Using this protocol, we were able to image 160 &#181;m thick brain sections from mice infected with the parasite Toxoplasma gondii, which enables visualization and analysis of the spatial relationship between the encysting parasite and the infected neuron.

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma ...

Peering at Brain Polysomes with Atomic Force Microscopy

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, formed by ribosomes, mRNAs, several proteins and non-coding RNAs, represent integrated platforms where translational controls take place. However, while the ribosome has been widely studied, the organization of polysomes is still lacking comprehensive understanding. Thus much effort is required in order to elucidate polysome organization and any novel mechanism of translational control that may be embedded. Atomic force microscopy (AFM) is a type of scanning probe microscopy that allows the acquisition of 3D images at nanoscale resolution. Compared to electron microscopy (EM) techniques, one of the main advantages of AFM is that it can acquire thousands of images both in air and in solution, enabling the sample to be maintained under near physiological conditions without any need for staining and fixing procedures. Here, a detailed protocol for the accurate purification of polysomes from mouse brain and their deposition on mica substrates is described. This protocol enables polysome imaging in air and liquid with AFM and their reconstruction as three-dimensional objects. Complementary to cryo-electron microscopy (cryo-EM), the proposed method can be conveniently used for systematically analyzing polysomes and studying their organization.

While ribosome structure has been extensively characterized, the organization of polysomes is still understudied. To overcome this lack of knowledge, we present here a detailed preparation protocol for accurate imaging of mammalian polysomes by atomic force microscopy (AFM) in air and liquid.

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, ...

A Protocol for the Administration of Real-Time fMRI Neurofeedback Training

Neurologic disorders are characterized by abnormal cellular-, molecular-, and circuit-level functions in the brain. New methods to induce and control neuroplastic processes and correct abnormal function, or even shift functions from damaged tissue to physiologically healthy brain regions, hold the potential to dramatically improve overall health. Of the current neuroplastic interventions in development, neurofeedback training (NFT) from functional Magnetic Resonance Imaging (fMRI) has the advantages of being completely non-invasive, non-pharmacologic, and spatially localized to target brain regions, as well as having no known side effects. Furthermore, NFT techniques, initially developed using fMRI, can often be translated to exercises that can be performed outside of the scanner without the aid of medical professionals or sophisticated medical equipment. In fMRI NFT, the fMRI signal is measured from specific regions of the brain, processed, and presented to the participant in real-time. Through training, self-directed mental processing techniques, that regulate this signal and its underlying neurophysiologic correlates, are developed. FMRI NFT has been used to train volitional control over a wide range of brain regions with implications for several different cognitive, behavioral, and motor systems. Additionally, fMRI NFT has shown promise in a broad range of applications such as the treatment of neurologic disorders and the augmentation of baseline human performance. In this article, we present an fMRI NFT protocol developed at our institution for modulation of both healthy and abnormal brain function, as well as examples of using the method to target both cognitive and auditory regions of the brain.

The ability to induce and/or control neural plasticity may be critical in future treatments for neurologic disorders and the recovery from brain injury. In this paper, we present a protocol on the use of neurofeedback training with functional magnetic resonance imaging to modulate human brain function.

Neurologic disorders are characterized by abnormal cellular-, molecular-, and circuit-level functions in the brain. New methods to induce and control ...

Live Images of GLUT4 Protein Trafficking in Mouse Primary Hypothalamic Neurons Using Deconvolution Microscopy

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral tissues. The hypothalamus in the central nervous system (CNS) is the control center for energy and insulin signal response regulation. Chronic inflammation in peripheral tissues and imbalances of certain chemokines (such as CCL5, TNF&#945;, and IL-6) contribute to diabetes and obesity. However, the functional mechanism(s) connecting chemokines and hypothalamic insulin signal regulation still remain unclear.
In vitro primary neuron culture models are convenient and simple models which can be used to investigate insulin signal regulation in hypothalamic neurons. In this study, we introduced exogeneous GLUT4 protein conjugated with GFP (GFP-GLUT4) into primary hypothalamic neurons to track GLUT4 membrane translocation upon insulin stimulation. Time-lapse images of GFP-GLUT4 protein trafficking were recorded by deconvolution microscopy, which allowed users to generate high-speed, high-resolution images without damaging the neurons significantly while conducting the experiment. The contribution of CCR5 in insulin regulated GLUT4 translocation was observed in CCR5 deficient hypothalamic neurons, which were isolated and cultured from CCR5 knockout mice. Our results demonstrated that the GLUT4 membrane translocation efficiency was reduced in CCR5 deficient hypothalamic neurons after insulin stimulation.

This protocol describes a technique for observation of real-time Green Fluorescence Protein (GFP) tagged Glucose Transporter 4 (GLUT4) protein trafficking upon insulin stimulation and characterization of the biological role of CCR5 in the insulin&#8211;GLUT4 signaling pathway with Deconvolution Microscopy.

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral ...

Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

Drosophila neuromuscular junction (NMJ) is an excellent model system to study glutamatergic synaptic transmission. We describe the technique of focal macropatch recordings of synaptic currents from visualized boutons at the Drosophila larval NMJ. This technique requires customized fabrication of recording micropipettes, as well as a compound microscope equipped with a high magnification, long-distance water immersion objective, differential interference contrast (DIC) optics, and a fluorescent attachment. The recording electrode is positioned on the top of a selected synaptic bouton visualized with DIC optics, epi-fluorescence, or both. The advantage of this technique is that it allows monitoring the synaptic activity of a limited number of sites of release. The recording electrode has&#160;a diameter of several microns, and the release sites positioned outside of the electrode rim do&#160;not significantly affect the recorded currents. The recorded synaptic currents have fast kinetics and can be readily resolved. These advantages are especially important for the studies of mutant fly lines with enhanced spontaneous or asynchronous synaptic activity.

Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

Synaptic currents can be recorded focally from visualized synaptic boutons at the Drosophila third instar larvae neuromuscular junction. This technique enables monitoring the activity of a single synaptic bouton.

Drosophila neuromuscular junction (NMJ) is an excellent model system to study glutamatergic synaptic transmission. We describe the technique of focal ...