The bath recording technique was first used by Fricke (1984)8 and Beall et al. (1990)9 to measure electric fields generated during tail-flips. The technique was later modified and improved in the laboratory of Dr. Donald Edwards (Georgia State University) by his former graduate student Dr. Fadi A. Issa and his former postdoctoral associate Dr. Jens Herberholz. Further refinements have been made and new research applications have been tested in the laboratory of Dr. Jens Herberholz at the University of Maryland. I would like to thank my colleague Dr. David Yager for letting me use his high-speed video system and my research assistants David Rotstein and William Liden for help with the experiments.

Part 1: Recording chamber
<ol>
<li>The recording chamber is of rectangular shape and is made of thin walled glass. Chamber dimensions are 8.5 cm x 2 cm x 5 cm (length x width x height) for animals of 2.5 &ndash; 3.5 cm total lengths (measured from rostrum to telson). See Fig. 1 for an example of a chamber used in our experiments.</li>
<li>Alternatively, recording chambers can be made from other materials (e.g., non-toxic clear plastic). Chambers size may vary according to experimental procedure and chambers should be customized for each experimental series. For best results, chamber size should be as small as possible without restraining the animals in their natural behavior. As a rule of thumb, the lengths and width of the chamber should not be more than three times the size of the animal.</li>
</ol>
Part 2: Bath electrodes and ground wire
<ol>
<li>One pair of recording electrodes and a ground electrode are used. Electrodes are made of insulated copper wire (26 AWG with 0.25 mm insulation). One end of the bath electrodes is connected to an extracellular amplifier (A-M Systems 1700; see below) and 0.5 &ndash; 1.0 mm of insulation are stripped off the other ends. The ground wire is connected to the ground of the amplifier or any other grounded equipment and 2-3 cm of insulation is stripped off the other end. </li>
<li>Bath electrodes and ground wire are attached to the inside walls of the recording chamber with non-toxic glue. Recording electrodes are positioned centrally on both short sides of the chamber and opposite to each other (Fig. 1).</li>
<li>Ground electrode is positioned on one long side of the recording chamber perpendicular to the recording electrodes (Fig. 1). </li>
<li>Chamber is filled with deionized water. Best results are obtained with water of high resistance (~18 M&Omega;).</li>
</ol>
Part 3: Bath electrode recordings
<ol>
<li>Outputs from recording electrodes are amplified (1000x) by an extracellular amplifier (A-M Systems; Model 1700). Signal from the recording electrodes is filtered using a combination of low-frequency (&lt; 100 Hz) and high-frequency cut-offs (&gt; 5 KHz). Signal is then connected to a switch box and a data acquisition board (National Instruments). Digitized data is recorded, stored, and analyzed using data acquisition software (Photron Motion Tools).</li>
<li>Alternatively, amplified signal from recording electrodes can be digitized using other analog-digital converters (e.g., MDS Analytical Technologies; Digidata 1440) before digitized data is recorded using other commercially available data acquisition software (e.g., MDS Analytical Technologies; Axoscope).</li>
</ol>
Part 4: Stimulating probe
<ol>
<li>A stimulating probe is made of a glass pipette (14 cm length) equipped with a pair of fine wire electrodes. Electrode tips are exposed (0.2 mm) to produce an electronic signal when the animal is touched by the probe. This allows to measure the exact time of stimulation. </li>
<li>Outputs from stimulating probe are amplified (1000x) by an extracellular amplifier (A-M Systems; Model 1700). The signal is filtered using a combination of low-frequency (&lt; 100 Hz) and high-frequency cut-offs (&gt; 5 KHz). The signal is then connected to a switch box and a data acquisition board (National Instruments). Digitized data is recorded, stored, and analyzed using data acquisition software (Photron Motion Tools).</li>
</ol>
Part 5: Video recordings
<ol>
<li>A high-speed video camera (Fastcam-X 1280 PCI, Photron) is positioned perpendicular to the recording chamber to provide a side view. Level of brightness inside the recording chamber must be adjusted to provide best results, e.g. by using a goose-neck illuminator or other focusable light sources.</li>
<li>High-speed videography is combined and synchronized with electronic recordings using a breakout box &amp; data acquisition board (National Instruments). Amplified signal from the bath electrodes is connected to the breakout box and data acquisition board using BNC cables. An external hand-switch trigger starts the synchronized video and data acquisition.</li>
</ol>
Part 6: Experimental procedure
<ol>
<li>A single animal is introduced into the chamber and allowed to acclimate for 5 minutes. Escape tail-flips are elicited by single taps of different intensity to the head or the abdomen, respectively. The intensity of the taps is controlled by the experimenter. Each escape tail-flip is recorded with high-speed video at a frame rate of 1000 f/sec, and data points of electronic field potentials are recorded at 25 kHz. Start of the bath and video recording is initiated by manual switch on a trigger box. Recording time is determined by chosen frame rate (e.g., 1000 f/sec = 4 sec total recording time). Post- and pre-trigger recording times can be selected.</li>
<li>Note: Bath electrode recordings should be verified once for each tested species by combining bath electrode recordings with other available recordings methods. In crayfish, a pair of silver electrodes can be surgically implanted around the ventral nerve cord. Stimuli eliciting escape tail-flips can be applied and recorded electronic traces of implanted and bath electrodes can be compared.</li>
<li>Escape tail-flip behavior and corresponding neural activity can be recorded in a variety of different contexts, e.g. in response to visual threats, during predator attacks, or during agonistic encounters of two crayfish. Chamber size, recording mode, etc. has to be adjusted accordingly (see Discussion).</li>
</ol>
Part 7: Data analysis
<ol>
<li>Single video frames are analyzed using motion tool software (Photron). Electronic traces are used to identify type of neural circuit that was activated by each stimulus. Video data is compared to synchronized physiological recordings to determine the movement of the animal and the activated neural circuit. Each activated circuit produces a characteristic electronic signature (see Representative results).</li>
<li>Latency between probe contact and neural/muscular response are calculated for each experiment by measuring the delay between the onset of the probe signal and the onset of the signal recorded with the bath electrodes.</li>
</ol>
Part 8: Representative results
A series of single high-speed video frames and corresponding electric field recordings for escape tail-flip in response to a tactile stimulus delivered to the head or tail of a juvenile crayfish (Fig. 2).
Fig. 2A: Strong tactile stimulus to the head evoked a tail-flip controlled by the medial giant circuit. The recorded spike of the giant neuron (asterisks) and the large phasic deflection that follows enables non-ambiguous identification of the tail-flip as mediated by giant neuron activity. The backward movement shown in the video traces determines the identity of the activated neural circuit (MG).
Fig. 2B: Tail-flip mediated by the lateral giant circuit after a strong tactile stimulus was applied to the tail. Upward and forward motion seen in the video traces together with the synchronized electronic trace displaying the giant spike and the large, phasic initial deflection determines the identity of the activated neural circuit (LG).
Fig. 2C: Tail-flip controlled by non-giant circuitry. A more gradual tactile stimulus was delivered to the thorax of the animal. While the movement captured on video does not allow unambiguous identification of the activated circuit, the electronic recording lacks a giant spike and consists of much smaller deflections identifying the activated circuit (Non-G).
Fig. 3: Latency measurements for all three types of escape tail-flips. Time between probe contact and physiological response was measured for seven animals. Giant-mediated tail-flips are elicited significantly faster than non-giant tail-flips.
<img src="/files/ftp_upload/1297/Fig1.jpg" alt="" />&nbsp;
Figure 1: An example for a recording chamber used in our experiments for animals of 2.5-3.5 cm in total lengths. The bath electrodes are glued to opposite sides of the chamber while the ground wire is attached to the long side of the chamber and perpendicular to the bath electrodes.
Figure 2: Single video frames recorded at 1000 f/sec and corresponding electronic recordings for three different types of stimulation.
<img src="/files/ftp_upload/1297/Fig2A.jpg" alt="" />
A) A strong tactile stimulus was delivered to the head of the animal and elicited a medial giant (MG) mediated tail-flip. Six video-frames are displayed on the left. Recording trace from the probe used to touch the animal is shown in gray; the point of contact is indicated by the black arrowhead. Recording trace obtained with the bath electrodes is shown in blue. The inset shows the small giant axon spike that precedes the large phasic deflections. Grey bars correspond to the video-frames shown on the left. The first notable movement of the crayfish&rsquo;s tail occurred at frame #3, seven milliseconds after the contact with the probe.
<img src="/files/ftp_upload/1297/Fig2B.jpg" alt="" />
B) A strong tactile stimulus was delivered to the tail of the animal and elicited a lateral giant (LG) mediated tail-flip. Six video-frames are displayed on the left. Recording trace from the probe used to touch the animal is shown in gray; the point of contact is indicated by the black arrowhead. Recording trace obtained with the bath electrodes is shown in red. The inset shows the small giant axon spike that precedes the large phasic deflections. Grey bars correspond to the video-frames shown on the left. The first notable movement of the crayfish&rsquo;s tail occurred at frame #3, eight milliseconds after the contact with the probe.
<img src="/files/ftp_upload/1297/Fig2C.jpg" alt="" />
C) A weak and gradual tactile stimulus was delivered to the head of the animal and elicited a non-giant (Non-G) mediated tail-flip. Eight video-frames are displayed on the left. Recording trace from the probe used to touch the animal is shown in gray; the point of contact is indicated by the black arrowhead. Recording trace obtained with the bath electrodes is shown in black. The trace lacks the giant spike potential, the large initial deflections and consists of potentials of much smaller amplitude. Light gray bars correspond to the video-frames shown on the left. The first notable movement of the crayfish&rsquo;s tail occurred at frame #6, 115 milliseconds after the first contact with the probe.
<img src="/files/ftp_upload/1297/Fig3.jpg" alt="" />
Figure 3: Response latency measurements for seven different animals of both sexes and similar sizes (mean lengths &plusmn; stdv: 3.2 cm &plusmn; 0.2 cm, measured from rostrum to telson). MG (blue bar) and LG (red bar) tail-flips have significantly shorter response latencies than tail-flips mediated by Non-G (black bar) circuitry. Means and standard deviations are shown. Bars with the same letter did not differ significantly from each other (Wilcoxon Signed Rank test for pair-wise comparison, p &lt; 0.05).

<ol>
	<li>Herberholz, J., Issa, F. A., Edwards, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterns+of+neural+circuit+activation+and+behavior+during+dominance+hierarchy+formation+in+freely+behaving+crayfish.">Patterns of neural circuit activation and behavior during dominance hierarchy formation in freely behaving crayfish.</a> J. Neurosci. 21, 2759-2767 (2001).</li><li>Herberholz, J., Sen, M. M., Edwards, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Escape+behavior+and+escape+circuit+activation+in+juvenile+crayfish+during+prey-predator+interactions.">Escape behavior and escape circuit activation in juvenile crayfish during prey-predator interactions.</a> J. Exp. Biol. 207, 1855-1863 (2004).</li><li>Liden, W. H., Herberholz, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Behavioral+and+neural+responses+of+juvenile+crayfish+to+moving+shadows.J.">Behavioral and neural responses of juvenile crayfish to moving shadows.J.</a> Exp. Biol. 211, 1355-1361 (2008).</li><li>Finley, L. A., Macmillan, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+analysis+of+field+potentials+during+different+tailflip+behaviours+in+crayfish.">An analysis of field potentials during different tailflip behaviours in crayfish.</a> Mar. Freshw. Behav. Physiol. 35, 221-234 (2002).</li><li>Eaton, R. C., Lee, R. K. K., Foreman, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Mauthner+cell+and+other+identified+neurons+of+the+brainstem+escape+network+of+fish.">The Mauthner cell and other identified neurons of the brainstem escape network of fish.</a> Prog. Neurobiol. 63, 467-485 (2001).</li><li>Canfield, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Some+voluntary+C-bends+may+be+Mauthner+neuron+initiated.">Some voluntary C-bends may be Mauthner neuron initiated.</a> J. Comp. Physiol. A. 193, 1055-1064 (2007).</li><li>W&#246;hl, S., Schuster, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+predictive+start+of+hunting+archer+fish:+a+flexible+and+precise+motor+pattern+performed+with+the+kinematics+of+an+escape+C-start.">The predictive start of hunting archer fish: a flexible and precise motor pattern performed with the kinematics of an escape C-start.</a> J. Exp. Biol. 210, 311-324 (2007).</li><li>Fricke, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+habituation+in+the+crayfish+due+to+selective+weakening+of+electrical+synapses.">Development of habituation in the crayfish due to selective weakening of electrical synapses.</a> Brain Res. 322, 139-143 (1984).</li><li>Beall, S. P., Langley, D. J., Edwards, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhibition+of+escape+tailflip+in+crayfish+during+backward+walking+and+the+defense+posture.">Inhibition of escape tailflip in crayfish during backward walking and the defense posture.</a> J. Exp. Biol. 152, 577-582 (1990).</li></ol>

The authors have nothing to disclose.

Non-invasive recordings of single neuron activity or neural circuit activation are difficult to obtain in unrestrained animals. The method described here provides a means to identify patterns of neural activation underlying naturally occurring behavior.
In the past, we successfully used this technique to measure patterns of activity in neural escape circuits of juvenile crayfish during the formation of social dominance hierarchies1, during attacks from natural predators2,...

recordings of neural circuit activation in freely behaving animals

The relationship between patterns of neural activity and corresponding behavioral expression is difficult to establish in unrestrained animals. Traditional non-invasive methods require at least partially restrained research subjects, and they only allow identification of large numbers of simultaneously activated neurons. On the other hand, small ensembles of neurons or individual neurons can only be measured using single-cell recordings obtained from largely reduced preparations. Since the expression of natural behavior is limited in restrained and dissected animals, the underlying neural mechanisms that control such behavior are difficult to identify. 
Here, I present a non-invasive physiological technique that allows measuring neural circuit activation in freely behaving animals. Using a pair of wire electrodes inside a water-filled chamber, the bath electrodes record neural and muscular field potentials generated by juvenile crayfish during natural or experimentally evoked escape responses. The primary escape responses of crayfish are mediated by three different types of tail-flips which move the animals away from the point of stimulation. Each type of tail-flip is controlled by its own neural circuit; the two fastest and most powerful escape responses require activation of different sets of large &#8220;command&#8221; neurons. In combination with behavioral observations, the bath electrode recordings allow unambiguous identification of these neurons and the associated neural circuits. Thus activity of neural circuitry underlying naturally occurring behavior can be measured in unrestrained animals and in different behavioral contexts.

Watch this Scientific Journal Video about Recordings of Neural Circuit Activation in Freely Behaving Animals at JoVE.com

Recordings of Neural Circuit Activation in Freely Behaving Animals

Non-invasive measurements of neural activity patterns in freely behaving animals are obtained by combining neurophysiological recordings with high speed videography.

The relationship between patterns of neural activity and corresponding behavioral expression is difficult to establish in unrestrained animals. ...

recordings-of-neural-circuit-activation-in-freely-behaving-animals

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11.5K Views.  University of Maryland. לא פולשני מדידות של דפוסי פעילות עצבית להתנהג בחופשיות בעלי חיים מתקבלים על ידי שילוב של הקלטות neurophysiological עם videography במהירות גבוהה.

Video: Recordings of Neural Circuit Activation in Freely Behaving Animals

Whole Cell Recordings from Brain of Adult Drosophila

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by describing the dissecting solution and capture of the adult females used in our studies. The procedure for removing the whole brain intact, including both optic lobes, is illustrated. Dissection of the overlying trachea is also shown.  The isolated brain is not only small but needs special care in handling at this stage to prevent damage to the neurons, many of which are close to the outer surface of the tissue. We show how a special holder we developed is used to stabilize the brain in the recording chamber. A standard electrophysiology set up is used for recording from single neurons or pairs of neurons. A fluorescent image, viewed through the recording microscope, from a GAL4 line driving GFP expression (GH146) illustrates how projection neurons (PNs) are identified in the live brain. A high power Nomarski image shows a view of a single neuron that is being targeted for whole cell recording. When the brain is successfully removed without damage, the majority of the neurons are spontaneously active, firing action potentials and/or exhibiting spontaneous synaptic input. This in situ preparation, in which whole cell recording of identified neurons in the whole brain can be combined with genetic and pharmacological manipulations, is a useful model for exploring cellular physiology and plasticity in the adult CNS.

This video demonstrates the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons using standard whole cell technology. It includes images of GFP labeled cells and neurons viewed during recording.

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by ...

Whole-cell Recordings of Light Evoked Excitatory Synaptic Currents in the Retinal Slice

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we will guide you through the process of performing whole-cell recordings of light evoked currents of individual cells in the retinal slice preparation. We use the aquatic tiger salamander as an animal model. We begin by describing the dissection of the eye and show how slices are mounted for electrophysiological recordings. Once the slice is placed in the recording chamber, we demonstrate how to perform whole-cell voltage clamp recordings. We then project visual stimuli onto the photoreceptors in the slice to elicit light-evoked current responses. During the recording we perfuse the slice with pharmacological agents, whereby an 8-channel perfusion system allows us to quickly switch between different agents.  The retinal slice preparation is widely used for patch clamp recordings in the retina, in particular to study amacrine or bipolar cells, which are not accessible in a whole-mount preparation.

This video shows the process of whole-cell voltage clamp recordings in the retinal slice of the aquatic tiger salamander. We demonstrate the preparation of the slice as well as how to perform patch clamp recordings during visual stimulation of the retina.

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we ...

Assay for Neural Induction in the Chick Embryo

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying the formation and initial patterning of the nervous system. This video demonstrates how to graft organizer tissue into a host, a method by which Hensen s node (the organizer in the chick embryo) is grafted to a host competent ectoderm. The organizer graft instructs overlying na ve tissue to adopt a neural fate via neural inducing signals. This mechanism is referred to as neural induction, and constitutes the initial step in the formation of brain and spinal cord in amniotes. This method is essentially used for the characterization of putative neural inducing molecules in chick. This video demonstrates the different steps in the assay for neural induction; First, the donnor embryo is explanted and pinned on a dish. Then, the host embryo is prepared for New culture. The graft is excised and transplanted to the host area pellucida margin. The host is cultured for 18-22 hrs. The assembly is fixed and processed for further applications (e.g. in situ hybridization). This method was originally devised by Waddington 1,2 and Gallera 3,4.

Neural induction is the first step in the formation of the brain. It is a mechanism by which Hensen's node (organizer), instructs adjacent tissue to adopt a neural fate, i.e. to give rise to the nervous system. This video demonstrates an assay for neural induction in chick embryo.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Tracking Dynamics of Muscle Engraftment in Small Animals by In Vivo Fluorescent Imaging

Muscular dystrophies are a group of degenerative muscle diseases characterized by progressive loss of contractile muscle cells. Currently, there is no curative treatment available. Recent advances in stem cell biology have generated new hopes for the development of effective cell based therapies to treat these diseases. Transplantation of various types of stem cells labeled with fluorescent proteins into muscles of dystrophic animal models has been used broadly in the field. A non-invasive technique with the capability to track the transplanted cell fate longitudinally can further our ability to evaluate muscle engraftment by transplanted cells more accurately and efficiently.
In the present study, we demonstrate that in vivo fluorescence imaging is a sensitive and reliable method for tracking transplanted GFP (Green Fluorescent Protein)-labeled cells in mouse skeletal muscles. Despite the concern about background due to the use of an external light necessary for excitation of fluorescent protein, we found that by using either nude mouse or eliminating hair with hair removal reagents much of this problem is eliminated. Using a CCD camera, the fluorescent signal can be detected in the tibialis anterior (TA) muscle after injection of 5 x 105 cells from either GFP transgenic mice or eGFP transduced myoblast culture. For more superficial muscles such as the extensor digitorum longus (EDL), injection of fewer cells produces a detectable signal. Signal intensity can be measured and quantified as the number of emitted photons per second in a region of interest (ROI). Since the acquired images show clear boundaries demarcating the engrafted area, the size of the ROI can be measured as well. If the legs are positioned consistently every time, the changes in total number of photons per second per muscle and the size of the ROI reflect the changes in the number of engrafted cells and the size of the engrafted area. Therefore the changes in the same muscle over time are quantifiable. In vivo fluorescent imaging technique has been used primarily to track the growth of tumorogenic cells, our study shows that it is a powerful tool that enables us to track the fate of transplanted stem cells.

Tracking Dynamics of Muscle Engraftment in Small Animals by In Vivo Fluorescent Imaging

We describe an in vivo fluorescence imaging protocol to monitor muscle regeneration by GFP-labeled myoblasts after transplantation into skeletal muscles of both healthy and dystrophic mice. This protocol can be adapted to study muscle regeneration by transplantation of other types of cells and in other muscular conditions as well.

Muscular dystrophies are a group of degenerative muscle diseases characterized by progressive loss of contractile muscle cells. Currently, there is no ...

Measures of Heart and Ventilatory Rates in Freely Moving Crayfish

The fear, flight or fight response serves as the fundamental physiological basis for examining an organism's awareness of its environment under an impending predator attack.  Although it is not known whether invertebrates posses an autonomic nervous system identical to that of vertebrates, evidence shows invertebrates have a  sympathetic-like  response to regulate the internal environment and ready the organism to act behaviorally to a given stimuli.  Furthermore, this physiological response can be feasibly measured and it acts as a biological index for the animal's internal state.  Measurements of the physiological response can be directly related to internal and external stressors through changes in the central nervous system controlled coordination of the cardio-vascular and respiratory systems.   More specifically, monitoring heart and ventilation rates provide quantifiable measures of the stress response not always behaviorally observed. Crayfish are good model organisms for heart and ventilatory rate measurements due to the feasibility of recording, as well as the rich history known of the morphology of the crayfish, dating back to Huxley in 1888, and the well-studied typical behaviors.

Invertebrates show an autonomic sympathetic-like response similar to that described for vertebrates. The coordination of the cardio-vascular and ventilatory systems allows for measurement of a biological index in which to assess an organism internal state.

The fear, flight or fight response serves as the fundamental physiological basis for examining an organism's awareness of its environment under an ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

RhoC GTPase Activation Assay

RhoC GTPase has 91% homology to RhoA GTPase. Because of its prevalence in cells, many reagents and techniques for RhoA GTPase have been developed. However, RhoC GTPase is expressed in metastatic cancer cells at relatively low levels. Therefore, few RhoC-specific reagents have been developed. We have adapted a Rho activation assay to detect RhoC GTPase. This technique utilizes a GST-Rho binding domain fusion protein to pull out active RhoC GTPase. In addition, we can harvest total protein at the beginning of the assay to determine levels of total (GTP and GDP bound) RhoC GTPase. This allows for the determination of active versus total RhoC GTPase in the cell. Several commercial versions of this procedure have been developed however, the commercial kits are optimized for RhoA GTPase and typically do not work well for RhoC GTPase. Parts of the assay have been modified as well as development of a RhoC-specific antibody.

This protocol utilizes a pull down assay to determine the levels of active RhoC GTPase within cells.

RhoC GTPase has 91% homology to RhoA GTPase.  Because of its prevalence in cells, many reagents and techniques for RhoA GTPase have been developed.  ...

Modeling Biological Membranes with Circuit Boards and Measuring Electrical Signals in Axons: Student Laboratory Exercises

This is a demonstration of how electrical models can be used to characterize biological membranes. This exercise also introduces biophysical terminology used in electrophysiology. The same equipment is used in the membrane model as on live preparations. Some properties of an isolated nerve cord are investigated: nerve action potentials, recruitment of neurons, and responsiveness of the nerve cord to environmental factors.

This is a demonstration of how biological membranes can be understood using electrical models. We also demonstrate procedures for recording action potentials from the ventral nerve cord of the crayfish for student orientated laboratories.

This is a demonstration of how electrical models can be used to characterize biological membranes. This exercise also introduces biophysical terminology ...

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and hermaphrodites. In the earlier phase of gametogenesis, the germ cells of hermaphrodites differentiate into limited number of sperm - around 300 - and are stored in a small 'bag' called the spermatheca. Later on, hermaphrodites continually produce oocytes1. In contrast, males produce exclusively sperm throughout their adulthood. The males produce so much sperm that it accounts for &gt;50% of the total cells in a typical adult worm2. Therefore, isolating sperm from males is easier than from that of hermaphrodites.
Only a small proportion of males are naturally generated due to spontaneous non-disjunction of X chromosome3. Crossing hermaphrodites with males or more conveniently, the introduction of mutations to give rise to Him (High Incidence of Males) phenotype are some of strategies through which one can enrich the male population3.
Males can be easily distinguished from hermaphrodites by observing the tail morphology4. Hermaphrodite's tail is pointed, whereas male tail is rounded with mating structures.
Cutting the tail releases vast number of spermatids stored inside the male reproductive tract. Dissection is performed under a stereo microscope using 27 gauge needles. Since spermatids are not physically connected with any other cells, hydraulic pressure expels internal contents of male body, including spermatids2.
Males are directly dissected on a small drop of 'Sperm Medium'. Spermatids are sensitive to alteration in the pH. Hence, HEPES, a compound with good buffering capacity is used in sperm media. Glucose and other salts present in sperm media help maintain osmotic pressure to maintain the integrity of sperm.
Post-meiotic differentiation of spermatids into spermatozoa is termed spermiogenesis or sperm activation. Shakes5, and Nelson6 previously showed that round spermatids can be induced to differentiate into spermatozoa by adding various activating compounds including Pronase E. Here we demonstrate in vitro spermiogenesis of C. elegans spermatids using Pronase E.
Successful spermiogenesis is pre-requisite for fertility and hence the mutants defective in spermiogenesis are sterile. Hitherto several mutants have been shown to be defective specifically in spermiogenesis process7. Abnormality found during in vitro activation of novel Spe (Spermatogenesis defective) mutants would help us discover additional players participating in this event.

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

A protocol for isolating and activating spermatids from male C. elegans is described here. Cutting the posterior end of male releases spermatids. The spermatids can be activated by addition of protease.

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and ...

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1. Apoptosis can be triggered when cells encounter a wide range of cytotoxic stresses. These insults initiate signaling cascades that ultimately cause the release of cytochrome c from the mitochondrial intermembrane space to the cytoplasm2. The release of cytochrome c from mitochondria is a key event that triggers the rapid activation of caspases, the key cellular proteases which ultimately execute cell death3-4.
The pathway of apoptosis is regulated at points upstream and downstream of cytochrome c release from mitochondria5. In order to study the post-mitochondrial regulation of caspase activation, many investigators have turned to direct cytoplasmic microinjection of holocytochrome c (heme-attached) protein into cells6-9. Cytochrome c is normally localized to the mitochondria where attachment of a heme group is necessary to enable it to activate apoptosis10-11. Therefore, to directly activate caspases, it is necessary to inject the holocytochrome c protein instead of its cDNA, because while the expression of cytochrome c from cDNA constructs will result in mitochondrial targeting and heme attachment, it will be sequestered from cytosolic caspases. Thus, the direct cytosolic microinjection of purified heme-attached cytochrome c protein is a useful tool to mimic mitochondrial cytochrome c release and apoptosis without the use of toxic insults which cause cellular and mitochondrial damage.
In this article, we describe a method for the microinjection of cytochrome c protein into cells, using mouse embryonic fibroblasts (MEFs) and primary sympathetic neurons as examples. While this protocol focuses on the injection of cytochrome c for investigations of apoptosis, the techniques shown here can also be easily adapted for microinjection of other proteins of interest.

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

In this protocol, we describe the direct cytoplasmic microinjection of cytochrome c protein into fibroblasts and primary sympathetic neurons. This technique allows for the introduction of cytochrome c protein into the cytoplasm of cells and mimics the release of cytochrome c from mitochondria, which occurs during apoptosis.

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1.  Apoptosis can be triggered when cells encounter a ...