I am grateful to Ronald L. Calabrese, Dieter Jaeger (Emory University) and Ward Yuhas (Axon Instruments) for useful technical discussions.

<ol>
 <li>Hypothalamic brain slices are prepared, incubated and transferred to the recording chamber as previously detailed (9) in animals whose GnRH neurons express green fluorescent protein (GFP) under the control of the GnRH peptide promoter (10). After removal of the brain from the animal, the brain is blocked using a razor blade to eliminate regions that are not of interest. In order to determine the regions to remove, the brain is placed on its dorsal surface so that the hypothalamus can be visualized. For both the coronal and sagittal slice orientations, the cerebellum is removed. For coronal slice preparation, the rostral portion of the cortex is removed. In this slice orientation, the lateral regions of the cortex provide support for slicing the brain and provide sufficient area of brain for the placement of silver wires to secure the brain in the recording chamber. For the sagittal slice preparation, the lateral regions of the brain are removed and the rostral portions of the cortex provide the support for brain slicing and for placement of silver wires to secure the brain in the recording well. Throughout the blocking procedure, the brain is continuously moistened with cold artificial cerebrospinal fluid (ACSF) using a glass pipette. </li>
 <li>A small amount of Superglue is placed on the cutting platform. The brain is lifted with a spatula. Excess ACSF is removed by bring a Kimwipe to the surface of the spatula and drawing excess ACSF away from the brain. The brain is gently slid forward onto the glue. It is important to not to touch the brain or push down on it in order to secure it. The weight of the brain itself is usually sufficient to ensure it is secure.</li>
 <li>The brain and the cutting platform is then secured to the vibrating microtome and the cutting well is filled with cold ACSF. The cold solution both firms the brain and may protect from damage during the cutting procedure. Different brain regions vary in the degree of coldness that optimizes quality slices. The temperature of the ACSF for preparation of hypothalamic slices is generally 0&deg;C.</li>
 <li>Slices should be cut slowly and each slice should float freely away from the brain. If slices begin to curl during cutting, the speed at which the blade is advancing should be slowed. One can use a small paint brush to uncurl slices but this does risk damaging the slices and thus, should be avoided. Slowly the speed of advance of the blade is preferred over using a paint brush.</li>
 <li>As each slice is cut from the brain, it is withdrawn from the cutting chamber and placed in a slice incubation chamber in a warm water bath (approximately 32&deg;C). The slice incubator is continuously supplied with oxygen using a mixture of 95% O2 and 5% CO2. Generally, pre-recording slice incubation ranges from 30 minutes to 2 hours.</li>
 <li>Glass pipettes (3-6 M&Omega;) are fabricated as previously described (9) using a vertical puller. The temperature and duration of pulling varies based on the type of pipette puller one uses, the type of glass one uses and the lifetime of the heating filament of the puller. The shape of the pipette tip can also vary depending on the user's preference. We have achieved good success with a uniformly tapered tip with an opening of 0.1 mm. Larger openings in the pipette tend to damage the neurons while smaller openings fail to result in long duration recordings.</li>
 <li>Pipettes coated with Sylgard to reduce capacitance. Pulled pipette tips are lightly coated with Sylgard 184 using a second pipette. Sylgarded tips are then heat-cured using a heat gun. These pipettes are then filled with the bath perfusate. To increase visualization of the pipette tips, a small amount of Alexa-568 is added to the portion of the bath solution to be used as the pipette solution before filling the pipettes. We do not weigh the Alexa-568 but rather use enough to turn the solution pink by eye. The pipette tips can then be visualized under the Texas red filter.</li>
 <li>We have developed an approach to obtain current-clamp recordings for detection of action potentials using low resistance seals (see Discussion). Recordings are performed at the cell's endogenous resting potential with no applied current using Axon Instrument's 2B Axoclamp. However, due to the low resistance seal, one could not detect action potentials with just the 2B amplifier. To increase the signal, a second amplifier (AM Systems 3000) is used in series with the Axoclamp 2B. This approach has the ease of loose seals and utility of long-term recordings. Technically, one can directly measure action potentials, thereby circumventing the problems with the approach in the voltage-clamp recording mode (see Discussion).</li>
 <li>One takes two pipettes to the surface of the two previously selected neurons at the same time, in a similar fashion as the approach for the whole cell recording configuration. Approaching both neurons at the same time has been more successful in obtaining the dual recordings than attempting each neuron of the pair independently. During this time, current injection pulses are passed from the 2B amplifier as is done during the approach for whole-cell recordings.</li>
 <li>Positive pressure must be maintained on both pipettes to prevent clogging the tip. Positive pressure for dual recordings is best generated by using an unfilled 3 ml empty syringe attached to the pipette holder by a length of piece of tubing. Using syringes allows one to maintain pressure on both pipettes. Positive pressure should be applied as soon as the pipette has been placed in the bath solution of the recording chamber. One approaches the selected neurons with the pipettes sequentially. After positioning the first pipette directly over the largest surface of the neuron, one maintains positive by leaving the syringe in place and leaves the pipette in this position. One then returns to the top of the recording chamber and lowers the second recording pipette into position. Selected neurons are often at different levels of the slice. The first pipette should be positioned over the neuron that is deepest in the slice and position the second pipette at the more superficial neuron. This prevents the positioning of the second pipette from shifting the positioning of the second pipette as the slice will move slightly in the vertical plane as pipettes enter the slice. </li>
 <li>Loose seals (18-30 M&Omega;) will be used. The positive pressure will induce a small dimple on the surface of each cell. One attempts to seal the first neuron by releasing positive pressure and applying very slight negative pressure. The positive pressure is released by removing the syringe. Suction by mouth is then applied to the free end of the tubing. A healthy neuron's membrane will quickly respond to the release of positive pressure and adhere to the pipette. Slight suction will form a seal with appropriate resistance in a healthy neuron.</li> 
 <li>When one attempts to seal a neuron and fails, the pipette cannot be re-used since membranes only seal to very clean glass. Instead, one raises the pipette (and microscope objective) away from the surface of the slice to the top of the perfusion well, removes the pipette from the bath, changes to a clean pipette and then returns to the slice to attempt to seal another cell. For these reason, it is important to select multiple potential neurons for recordings. For slices with less than 4-6 recording quality neurons from which to choose, the success in obtaining dual recordings is limited. </li>
 <li>After establishing a loose seal based on resistance, the current injection pulses from the 2B amplifier are terminated. The 2B amplifer is placed in series with the 3000 series amplifer. In this configuration, the output of the 2B amplifer serves as the input for the 3000 series amplifer. The latter amplifer provides signal for data acquisition.</li>
 <li>The seal (step 4) and reconfiguration of amplifers (step 5) is then repeated for the second neuron.</li>
</ol>
<img src="/files/ftp_upload/1678/1678fig1tmb.jpg" alt="Figure 1" border="0" /> Figure 1. A period of increased firing in two GnRH neurons using the loose seal attached recording configuration in a sagittal slice preparation. The slice was derived from a castrated male. Each upward deflection indicates an action potential. Note that one GnRH neuron (top traces) exhibits nearly continuous firing while the second GnRH neuron exhibits intermittent bursts. This pattern of activity from single GnRH neurons is similar to that of single units extracted from multi-unit recordings during hormone secretion in vivo (7). Please <a href="https://www.jove.com/files/ftp_upload/1678/1678fig1.jpg">click here</a> to see a larger version of figure 1.

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T., Knobil, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+unit+components+of+the+hypothalamic+multiunit+electrical+activity+associated+with+the+central+signal+generator+that+directs+the+pulsatile+secretion+of+gonadotropic+hormones.">Single unit components of the hypothalamic multiunit electrical activity associated with the central signal generator that directs the pulsatile secretion of gonadotropic hormones.</a> Proc Natl Acad Sci U S A. 90, 9630-9634 (1993).</li><li>Silverman, A. J., Wilson, R., Kesner, J. 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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dendrites+determine+the+contribution+of+after+depolarization+potentials+(ADPs)+to+generation+of+repetitive+action+potentials+in+hypothalamic+gonadotropin+releasing-hormone+(GnRH)+neurons.">Dendrites determine the contribution of after depolarization potentials (ADPs) to generation of repetitive action potentials in hypothalamic gonadotropin releasing-hormone (GnRH) neurons.</a> J Comput Neurosci. 26, 39-53 (2009).</li><li>Spergel, D. J., Kruth, U., Hanley, D. F., Sprengel, R., Seeburg, P. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GABA-+and+glutamate-activated+channels+in+green+fluorescent+protein-tagged+gonadotropin-releasing+hormone+neurons+in+transgenic+mice.">GABA- and glutamate-activated channels in green fluorescent protein-tagged gonadotropin-releasing hormone neurons in transgenic mice.</a> J Neurosci. 19, 2037-2050 (1999).</li><li>Nelson, J. F., Felicio, L. S., Randall, P. K., Sims, C., Finch, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+longitudinal+study+of+estrous+cyclicity+in+aging+C57BL/6J+mice:+I.+Cycle+frequency,+length+and+vaginal+cytology.">A longitudinal study of estrous cyclicity in aging C57BL/6J mice: I. Cycle frequency, length and vaginal cytology.</a> Biol Reprod. 27, 327-339 (1982).</li><li>Gee, D. M., Flurkey, K., Finch, C. 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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanosensitivity+of+N-type+calcium+channel+currents.">Mechanosensitivity of N-type calcium channel currents.</a> Biophys J. 83, 2560-2574 (2002).</li><li>Higure, Y., Katayama, Y., Takeuchi, K., Ohtubo, Y., Yoshii, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+killing+of+single+neurons+by+irradiation+of+intracellularly+injected+dye.">Rapid killing of single neurons by irradiation of intracellularly injected dye.</a> Science. 206, 702-704 (1979).</li><li>Higure, Y., Katayama, Y., Takeuchi, K., Ohtubo, Y., Yoshii, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lucifer+Yellow+slows+voltage-gated+Na++current+inactivation+in+a+light-dependent+manner+in+mice.">Lucifer Yellow slows voltage-gated Na+ current inactivation in a light-dependent manner in mice.</a> J Physiol. 550, 159-167 (2003).</li></ol>

The activity of interest in some neurons including GnRH neurons (based on hormone secretion) occurs on time scales of hours (5-7). Therefore, the whole cell configuration is not the best choice for some experimental goals due to the dialysis of intracellular messengers in the whole cell recording mode. Secondarily, whole-cell recordings are generally limited to animals less than about 120 days of age. With age, neuronal membranes appear to stiffen, making high resistance seals difficult to achieve. Additionally, if one obtains a high resistance seal, rupturing the patch disrupts the seal, leaving a hole between the pipette and the membrane. This leads to an unusable recording and a neuron that will quickly die due to ionic imbalances. Regular ovarian cycles, and thus, stable activity of the GnRH pulse generator does not occur until later in life (7-10 months of age in C57Bl6 females; 11, 12), beyond the age when one can reasonably anticipate obtaining whole-cell recordings reliably. Finally, whole-cell recordings destroy the endogenous ratios of internal and external ion concentrations. With whole cell recordings, the internal concentration of any ion equals the concentration of the ion in the pipette solution. This is because the pipette solution's relatively large volume rapidly reaches equilibrium with/replaces the relatively small endogenous volume of the cell.
The loose cell attached approach circumvents many of the limitations of whole-cell recordings. First, a low resistance seal (15-30 M&Omega;) can be used. These are relatively easy to form even in neurons from older animals. Secondly, one does not rupture the sealed patch of membrane. Therefore, loose cell attached recordings are technically much easier than whole-cell recordings. Additionally, since the cell membrane is intact, dialysis of intracellular components does not occur and endogenous ionic ratios are preserved. One can not use the loose cell attached approach for studying synaptic currents but it is ideal for long-term recordings from neurons in a relatively non-invasive manner. The cell-attached recordings can also be performed using any standard intracellular solution in the pipette. This offers the additional advantage of rupturing the membrane patch when the long term recording is completed and labeling the neuron with an intracellular marker.
The loose cell attached approach has been used in the voltage-clamp recording mode. However, voltage-clamp recording in the loose cell attached configuration has several methodological problems. First, the recorded signal is an indirect measure of activity. The signal that is measured (as the so-called action current) is the capacitive current that charges the membrane (13). This is an extremely important methodological issue. The capacitance and resistance of a recording pipette can filter the recorded signal. It is very likely that the small action currents are lost in charging the capacitance of the pipette which, cannot be properly compensated with most amplifiers, due to the high resistance of the headstage. When these signals go undetected, the apparent firing pattern of the neuron does not reflect the true firing pattern. Likewise, uncompensated pipette and seal resistances cause significant errors in measurements during changes such as when action currents are expressed (13). Some amplifiers provide capacitance and resistance "compensation" for the pipette and seal, which limits signal loss, but high resistance head stages of most amplifiers hinder optimal compensation. Secondly, an artificial situation is imposed on the cell. In voltage-clamp mode, the area around the cell membrane is held to a fixed potential, in these studies, 0 mV. This does not mean there is no current applied to the cell membrane. The signal measured in voltage-clamp is actually the amount of current applied to the membrane to maintain the fixed potential. Therefore, this applied current can alter the cell activity.
Dual recordings in the GnRH system are particularly challenging due to the limited number of GnRH neurons and their diffuse distribution. For dual recordings to be successful, the manipulator must be exceedingly stable. Even slight movement of the electrode can cause the pipette to slip off the neuron and end the recording. Moreover, movement of the pipette on the cell (e.g., re-positioning to compensate for movement) can alter firing patterns. Some ion channels such N-type calcium channels are mechanically sensitive: membrane stretch causes repetitive activity in both whole-cell and cell-attached recording configurations (14). Finally, the manipulator system must be capable of exceedingly fine and smooth motion. As noted above, with dual recordings, one takes two pipettes to the surface of the two previously selected neurons at the same time and attempts to seal one neuron. If successful, then one attempts to seal the second cell. Generally, one cannot expect to seal and have a high quality recording with every attempt. This, however, creates a particular problem with dual recordings. If one is successful with the first neuron but fails with the second, one must change the pipette and try a different cell. Therefore, one must be able to move both the immersion objective of the microscope and the pipette to the top of the perfusion well (to change the pipette) without disrupting the successfully sealed neuron.
Our development and use of the loose cell-attached approach for dual recordings is a major technical advance in studying GnRH neurons. It is likely to produce useful results that will help move the field forward in the context of the critical question of what mechanisms underlie the coordinated activity that results in pulsatile hormone secretion.

dual somatic recordings from gonadotropin-releasing hormone (gnrh) neurons identified by green fluorescent protein (gfp) in hypothalamic slices

Gonadotropin-Releasing Hormone (GnRH) is a small neuropeptide that regulates pituitary release of luteinizing hormone (LH) and follicle-stimulating hormone (FSH). These gonadotropins are essential for the regulation of reproductive function. The GnRH-containing neurons are distributed diffusely throughout the hypothalamus and project to the median eminence where they release GnRH from their axon terminals into the hypophysiotropic portal system (1). In the portal capillaries, GnRH travels to the anterior pituitary gland to stimulate release of gonadotropins into systemic circulation. GnRH release is not continuous but rather occurs in episodic pulses. It is well established that the intermittent manner of GnRH release is essential for reproduction (2, 3).
Coordination of activity of multiple GnRH neurons probably underlies GnRH pulses. Total peptide content in GnRH neurons is approximately 1.0 pg/cell (4), of which 30% likely comprises the releasable pool. Levels of GnRH during a pulse (5, 6), suggest multiple GnRH neurons are probably involved in neurosecretion. Likewise, single unit activity extracted from hypothalamic multi-unit recordings during LH release indicates changes in activity of multiple neurons (7). The electrodes with recorded activity during LH pulses are associated with either GnRH somata or fibers (8). Therefore, at least some of this activity arises from GnRH neurons.
The mechanisms that result in synchronized firing in hypothalamic GnRH neurons are unknown. Elucidating the mechanisms that coordinate firing in GnRH neurons is a complex problem. First, the GnRH neurons are relatively few in number. In rodents, there are 800-2500 GnRH neurons. It is not clear that all GnRH neurons are involved in episodic GnRH release. Moreover, GnRH neurons are diffusely distributed (1). This has complicated our understanding of coordination of firing and has made many technical approaches intractable. We have optimized loose cell-attached recordings in current-clamp mode for the direct detection of action potentials and developed a recording approach that allows for simultaneous recordings from pairs of GnRH neurons.

Watch this Scientific Journal Video about Dual Somatic Recordings from Gonadotropin-Releasing Hormone GnRH Neurons Identified by Green Fluorescent Protein GFP in Hypothalamic Slices at JoVE.com

Dual Somatic Recordings from Gonadotropin-Releasing Hormone GnRH Neurons Identified by Green Fluorescent Protein GFP in Hypothalamic Slices

Activity in neuronal systems often requires synchronous action potential discharges from neurons within a specific population. For example, pulses of gonadotropin-releasing hormone (GnRH) likely require coordinated activity between GnRH neurons. We present our methodological approach for reliably obtaining simultaneous electrophysiological recordings from the diffusely distributed GnRH neurons.

Dual Somatic Recordings from Gonadotropin-Releasing Hormone (GnRH) Neurons Identified by Green Fluorescent Protein (GFP) in Hypothalamic Slices

Gonadotropin-Releasing Hormone (GnRH) is a small neuropeptide that regulates pituitary release of luteinizing hormone (LH) and follicle-stimulating ...

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22.2K Views.  University of Texas San Antonio - UTSA. פעילות עצבית במערכות לעתים קרובות מחייב הפרשות סינכרוני פוטנציאל פעולה מתא עצב בתוך אוכלוסייה מסוימת. לדוגמה, פולסים של הורמון גונדוטרופין משחררים (GnRH) סביר לדרוש פעילות מתואמת בין הנוירונים GnRH. אנו מציגים גישה מתודולוגית שלנו אמין קבלת הקלטות אלקטרו סימולטני ...

Video: Dual Somatic Recordings from Gonadotropin-Releasing Hormone GnRH Neurons Identified by Green Fluorescent Protein GFP in Hypothalamic Slices

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

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 ...

Imaging Calcium Responses in GFP-tagged Neurons of Hypothalamic Mouse Brain Slices

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the precise molecular steps as well as the activity of specific neurons in multi-functional nuclei of brain areas such as the hypothalamus remain. This problem includes identification of the molecular components involved in the regulation of various neurohormone signal transduction cascades. Elevations of intracellular Ca2+ play an important role in regulating the sensitivity of neurons, both at the level of signal transduction and at synaptic sites.
New tools have emerged to help identify neurons in the myriad of brain neurons by expressing green fluorescent protein (GFP) under the control of a particular promoter. To monitor both spatially and temporally stimulus-induced Ca2+ responses in GFP-tagged neurons, a non-green fluorescent Ca2+ indicator dye needs to be used. In addition, confocal microscopy is a favorite method of imaging individual neurons in tissue slices due to its ability to visualize neurons in distinct planes of depth within the tissue and to limit out-of-focus fluorescence. The ratiometric Ca2+ indicator fura-2 has been used in combination with GFP-tagged neurons1. However, the dye is excited by ultraviolet (UV) light. The cost of the laser and the limited optical penetration depth of UV light hindered its use in many laboratories. Moreover, GFP fluorescence may interfere with the fura-2 signals2. Therefore, we decided to use a red fluorescent Ca2+ indicator dye. The huge Stokes shift of fura-red permits multicolor analysis of the red fluorescence in combination with GFP using a single excitation wavelength. We had previously good results using fura-red in combination with GFP-tagged olfactory neurons3. The protocols for olfactory tissue slices seemed to work equally well in hypothalamic neurons4. Fura-red based Ca2+ imaging was also successfully combined with GFP-tagged pancreatic &beta;-cells and GFP-tagged receptors expressed in HEK cells5,6. A little quirk of fura-red is that its fluorescence intensity at 650 nm decreases once the indicator binds calcium7. Therefore, the fluorescence of resting neurons with low Ca2+ concentration has relatively high intensity. It should be noted, that other red Ca2+-indicator dyes exist or are currently being developed, that might give better or improved results in different neurons and brain areas.

In this protocol, we update recent progress in imaging Ca2+ signals of GFP-tagged neurons in brain tissue slices using a red fluorescent Ca2+ indicator dye.

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the ...

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 ...

Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be performed in an intact brain preparation where the neural circuitry of the CNS remains intact. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with green fluorescent protein for identification in the intact brain. Fish have multiple populations of GnRH neurons, and their functions are dependent on their location in the brain and the GnRH gene that they express1 . We have focused our demonstration on GnRH3 neurons located in the terminal nerves (TN) associated with the olfactory bulbs using the intact brain of transgenic medaka fish (Figure 1B and C). Studies suggest that medaka TN-GnRH3 neurons are neuromodulatory, acting as a transmitter of information from the external environment to the central nervous system; they do not play a direct role in regulating pituitary-gonadal functions, as do the well-known hypothalamic GnRH1 neurons2, 3 .The tonic pattern of spontaneous action potential firing of TN-GnRH3 neurons is an intrinsic property4-6, the frequency of which is modulated by visual cues from conspecifics2 and the neuropeptide kisspeptin 15. In this video, we use a stable line of transgenic medaka in which TN-GnRH3 neurons express a transgene containing the promoter region of Gnrh3 linked to enhanced green fluorescent protein7 to show you how to identify neurons and monitor their electrical activity in the whole brain preparation6.

In this video, we will demonstrate how to record electrical activity from identified single neurons in a whole brain preparation, which preserves complex neural circuits. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with a fluorescent protein for identification in the intact brain preparation.

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be ...

Whole-cell Patch-clamp Recordings from Morphologically- and Neurochemically-identified Hippocampal Interneurons

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal population (10-20%), but show a high level of physiological, morphological, and neurochemical heterogeneity, reflecting their diverse functions. Therefore, investigation of interneurons provides important insights into the organization principles and function of neuronal circuits. This, however, requires an integrated physiological and neuroanatomical approach for the selection and identification of individual interneuron types. Whole-cell patch-clamp recording from acute brain slices of transgenic animals, expressing fluorescent proteins under the promoters of interneuron-specific markers, provides an efficient method to target and electrophysiologically characterize intrinsic and synaptic properties of specific interneuron types. Combined with intracellular dye labeling, this approach can be extended with post-hoc morphological and immunocytochemical analysis, enabling systematic identification of recorded neurons. These methods can be tailored to suit a broad range of scientific questions regarding functional properties of diverse types of cortical neurons.

Cortical networks are controlled by a small, but diverse set of inhibitory interneurons. Functional investigation of interneurons therefore requires targeted recording and rigorous identification. Described here is a combined approach involving whole-cell recordings from single or synaptically-coupled pairs of neurons with intracellular labeling, post-hoc morphological and immunocytochemical analysis.

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Subcellular Patch-clamp Recordings from the Somatodendritic Domain of Nigral Dopamine Neurons

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding these fundamental functions will shed light on the information transfer in these neurons. Dendritic patch-clamp recordings provide the possibility to directly examine the electrical properties of dendrites and underlying voltage-gated ion channels. However, these fine structures are not easily accessible to patch pipettes because of their small diameter. This report describes a step-by-step procedure to collect stable and reliable recordings from the dendrites of dopaminergic neurons in acute slices. Electrophysiological measurements are combined with post hoc recovery of cell morphology. Successful experiments rely on improved preparation of slices, solutions and pipettes, adequate adjustment of the optics and stability of the pipette in contact with the recorded structure. Standard principles of somatic patch-clamp recording are applied to dendrites but with a gentler approach of the pipette. These versatile techniques can be implemented to address various questions concerning the excitable properties of dendrites.

Subcellular patch-clamp recordings offer the possibility to investigate the functional properties of dendrites. However these fine structures are not easily accessible due to their small diameter. The protocol described here aims to facilitate the collection of stable and reliable recordings from the dendrites of dopamine neurons in slices in vitro.

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding ...