Research reported in this publication was supported by the National Institute of Neurological Disorders and Stroke of the National Institutes of Health under Award Number RO1 NS 102360. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

All animal procedures were performed with approval of the Institutional Animal Care and Use Committee (IACUC A334818).
1. Calibration of the force transducer
<ol>
	<li>Ensure that the computer is properly connected to the USB-6009 multifunctional I/O data acquisition (DAQ) device, which in turn should be connected to the force transducer. 
	NOTE: Other rat strains and species may require a different load-cell force transducer as higher forces are to be expected 44.</li>
	<li>Attach a custom clamp fashioned from a modified surgical hemostat to the force transducer that is mounted to a vacuum base adjustable lever arm. 
	NOTE: The custom-made clamp consists of a surgical hemostat modified with a tightening screw that allows for adjustment of the tension (Figure 1).</li>
	<li>Position the custom-made acrylic glass testing platform, which contains two wooden blocks for fixation of the rat hind limb, on the table. 
	NOTE: Other materials such as urethane can also be used instead of wood as long as the K-wires are able to penetrate and fixate.</li>
	<li>Attach the clamp, force transducer and adjustable lever arm combination vertically to the testing platform using its vacuum base.</li>
	<li>Fasten a hook or loop to the clamp for the calibration weights.</li>
	<li>Turn on the computer and open the software (e.g., LabVIEW).</li>
	<li>Once the software is opened, start the custom-made virtual instrument (VI) for ITF measurement (Figure 2). 
	NOTE: Figure 2 contains the LabVIEW code in a VI snippet. This VI snippet can be dragged onto the block diagram in LabVIEW. It will automatically be transformed into a graphical code. For this experiment the sampling rate was set at 2000 Hz with 25 samples to read for each iteration.</li>
	<li>Run the VI by pressing the white arrow in the left upper corner and select New calibration. A new window will open.</li>
	<li>Start the calibration process with zero weight (only the clamp with an attached hook or loop) and press OK.</li>
	<li>Consecutively, add 10, 20, 30 and 50 grams of weight and press OK in between each weight measurement.</li>
	<li>Once all five measurements are collected, click on Process.</li>
	<li>Only accept the values if the graph on the VI displays a positive linear curve (Figure 3).</li>
	<li>Reposition the clamp, force transducer and adjustable lever arm combination horizontally on the testing platform. This will be the position used for measuring the ITF.</li>
	<li>Click on Zero and the window will automatically close.</li>
</ol>
2. Animal subjects
<ol>
	<li>Use male Lewis rats weighing between 300-500 g. 
	NOTE: For comparison of nerve regeneration, it is imperative to use the same rat strain in both the control and experimental groups, since weight and incidence of autotomy are strain dependent and can tremendously influence the results of the ITF10,32,45,46,47.</li>
</ol>
3. Surgical preparation
<ol>
	<li>Prepare all required surgical instruments prior to surgery (Table of Materials).</li>
	<li>Weigh the animals to determine the required amount of anesthesia.</li>
	<li>Induce anesthesia by placing the rat in a chamber gassed with 3% isoflurane in oxygen.</li>
	<li>Deeply anesthetize the rat using a cocktail of ten-parts ketamine (100 mg/mL) and one-part xylazine (100 mg/mL) at a dosage of 1 mL/kg body weight via an intraperitoneal injection. Monitor the depth of anesthesia based on the response to a toe pinch and by observing the respiratory rate.</li>
	<li>Approximately 30 minutes after the initial dosage of the ketamine/xylazine cocktail, administer a supplementary dose of 0.3-0.6 mL/kg body weight of only ketamine (100 mg/mL) intraperitoneally to maintain adequate anesthesia throughout the entire procedure, which is defined as a low respiratory rate and an absent response to a toe pinch. 
	CAUTION: It is important to meticulously administer the required anesthesia as an overdose cannot be counteracted.</li>
	<li>Carefully shave the hind limbs of the rat using electric clippers.</li>
	<li>Place the rat in prone position on a heating pad to maintain the body temperature at 37 &#176;C. Optionally, the body temperature can be monitored using a rectal thermometer.</li>
	<li>Inject 5 mL of 0.9% sodium chloride (NaCl) subcutaneously into the loose skin over the neck of the rat to preserve an adequate hydration status throughout the procedure.</li>
	<li>Due to the non-survival nature of this procedure, the surgical field and instruments do not require to be sterile. The surgeon should use personal protective equipment (PPE) and surgical loupes are advised for proper visualization of the anatomical structures.</li>
</ol>
4. Surgical approach to the common peroneal nerve
<ol>
	<li>Place the rat in either the right or left lateral recumbent position depending on which side will be measured first.</li>
	<li>Create a 2-3 cm incision in the skin of the posterolateral thigh parallel to the femur starting at the greater trochanter using a surgical no. 15 blade.</li>
	<li>Identify the plane between the biceps femoris muscle and the gluteus maximus and vastus lateralis muscles and perform a blunt dissection using tenotomy scissors to separate these muscles and expose the underlying sciatic nerve.</li>
	<li>Locate the trifurcation of the sciatic nerve and place a retractor to acquire better access. The three branches of the sciatic nerve include the common peroneal nerve, the tibialis nerve and the sural nerve.</li>
	<li>Isolate the common peroneal nerve branch (usually the most ventral branch) of the sciatic nerve using a curved microsurgical forceps. 
	&#8203;NOTE: In case of uncertainty, gently stimulate the isolated nerve with a surgical nerve stimulator and observe the motor response. Stimulation of the common peroneal nerve results in dorsiflexion of the paw.</li>
</ol>
5. Dissection of the distal tibialis anterior muscle tendon
<ol>
	<li>In order to expose the TA muscle and its insertion, incise the skin at the anterolateral aspect of the lower leg, starting at the knee joint and descending to the mediodorsal side of the hind paw.</li>
	<li>Dissect the distal TA muscle tendon from the surrounding tissue using a scalpel with a surgical blade no. 15.</li>
	<li>Using a mosquito forceps, bluntly dissect the TA muscle tendon towards the insertion and cut the tendon as distal as possible. Leave the proximal TA muscle undisturbed, preserving the neurovascular pedicle. 
	&#8203;NOTE: Regularly (approximately every 5 minutes), moist the TA muscle with heated 0.9% NaCl (37 &#176;C) to prevent cooling and desiccation.</li>
</ol>
6. Isometric tetanic force measurement
<ol>
	<li>Connect the bipolar electrode cables and the ground cable according to their color to a bipolar stimulator device.</li>
	<li>Attach the other end of the bipolar electrode cables to a subminiature electrode. 
	NOTE: The reference electrode (red, anode) should be placed distal and the active electrode (black, cathode) proximal.</li>
	<li>Transfer the animal together with the heating pad to the testing platform.</li>
	<li>Fixate the hind limb of the rat to the wooden block using two 1 mm Kirschner wires through the ankle and the lateral condyle of the distal femur avoiding the posterior aspect of the knee. 
	CAUTION: Avoid vascular damage to the popliteal artery and vein which are located dorsally to the femur condyle.</li>
	<li>Attach a holder with a custom clamp to the testing platform using its vacuum base.</li>
	<li>Secure the distal TA muscle tendon to the clamp attached to the force transducer. 
	NOTE: The clamp and force transducer should be positioned parallel to the course of the TA muscle.</li>
	<li>Place the retractor at the posterolateral thigh of the rat in order to access the common peroneal nerve. 
	NOTE: The sciatic nerve and its branches should be kept moist with heated 0.9% NaCl (37 &#176;C) to prevent cooling and desiccation.</li>
	<li>Insert the ground cable in the surrounding muscles (e.g., the vastus lateralis muscle). 
	NOTE: The Grass SD9 stimulator requires a ground cable to reduce electrical artifacts. Newer stimulators might not require an extra ground cable.</li>
	<li>Hook the common peroneal nerve to the subminiature electrode and fix its position using the holder on the platform (Figure 4). 
	NOTE: Ensure that only the common peroneal nerve is hooked to the subminiature electrode.</li>
	<li>Optimization of the muscle length
	<ol>
		<li>Turn the bipolar stimulator device on and adjust the settings as follow: square monophasic pulse, delay 2 ms, stimulus pulse duration 0.4 ms, stimulus intensity 2 V. 
		NOTE: The delay determines the time between the sync out pulse and the delivery of the leading edge of the pulse.</li>
		<li>Select Parameter test and turn on Trigger collection in the VI.</li>
		<li>Increase the muscle length (preload) by adjusting the lever arm attached to the force transducer.</li>
		<li>Start at 10 g of preload and use increments of 10 g until the maximum active muscle force is determined.</li>
		<li>For each preload, apply two single twitches directly after each other using the button on the bipolar stimulator device. The output will be visible on the screen and the rat should show dorsiflexion of the paw. 
		NOTE: Before stimulating the nerve, always remove any excess 0.9% NaCl surrounding the nerve using cotton tipped applicators to ensure the signal is not conducted to the surrounding tissue.</li>
		<li>To stop the measurement, hit Trigger collection again in the VI.</li>
		<li>If the program automatically detects the two peak output forces click on Accept. In case the program does not automatically select these output forces, press Decline and select the peaks manually. The two peak output forces will be averaged to a mean peak output force (Figure 5).</li>
		<li>Calculate the active muscle force by subtracting the preload from the mean peak output force.</li>
		<li>Write down the active force for each preload to visualize the trend and recognize the maximum active force (Figure 6). A spreadsheet can also be used.</li>
	</ol>
	</li>
	<li>Measurement of isometric tetanic force
	<ol>
		<li>After determining the ideal muscle length, let the muscle rest at zero preload for 5 minutes prior to starting the tetanic muscle contractions.</li>
		<li>Meanwhile, switch from Parameter test to Frequency test on the VI and adjust the stimulus intensity to 10 V on the bipolar stimulator device.</li>
		<li>Keep the delay and stimulus pulse duration at 2 ms and 0.4 ms, respectively.</li>
		<li>Measure the isometric tetanic muscle force using increasing stimulus frequencies starting at 30 Hz with increments of 30 Hz until the maximum force plateau is observed.</li>
		<li>Click on Trigger collection and set to the predetermined optimal muscle length.</li>
		<li>Press the Repeat button on the bipolar stimulator device to induce a tetanic stimulation for a maximum of 5 seconds or until a force peak is clearly observed. 
		NOTE: Before stimulating the nerve, always remove any excess 0.9% NaCl surrounding the nerve using cotton tipped applicators to ensure the signal is not conducted to the surrounding tissue.</li>
		<li>To collect the data, press Trigger collection again and document the maximum output force. In case the program does not automatically detect the peak maximum output force, press Decline and select the peak manually.</li>
		<li>Let the muscle rest again at zero preload for 5 minutes prior to starting the next tetanic muscle contractions. 
		NOTE: Regularly (approximately every 5 min), moist the TA muscle with heated 0.9% NaCl (37 &#176;C) to prevent cooling and desiccation.</li>
		<li>Continue increasing the stimulus frequency until the maximum force plateau is reached. The force plateau will be defined as the maximum isometric tetanic force. 
		NOTE: After this step, remove the K-wires, staple or suture the skin and repeat the entire procedure to the contralateral hind limb, starting at step 4.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

This protocol describes a previously validated method for acquiring accurate maximum ITF measurements of the TA muscle in the rat model32. The recovery of maximum strength after experimental nerve reconstruction treatments is of primary interest in the clinical setting as it proves that the nerve not only regenerated, but also made working connections with the target muscle. The ITF can be used in a small nerve gap model, such as the rat sciatic nerve model32, and with a fe...

Loss of motor function following traumatic peripheral nerve injury has a significant impact on the quality of life and socioeconomic status of patients1,2,3. The prognosis of this patient population remains poor due to minimal improvements in surgical techniques over the years4. Direct end-to-end tension-free epineural repair forms the gold standard surgical reconstruction. However, in cases with extended nerve gaps interposition of an autologous nerve graft has proven to be superior5,6. The associated donor site morbidity and limited availability of autologous nerve grafts have imposed the need for alternative techniques7,8.
Experimental animal models have been used to elucidate the mechanism of peripheral nerve regeneration and to evaluate outcomes of a variety of reconstructive and pharmacological treatment options8,9. The rat sciatic nerve model is the most frequently used animal model10. Their small size makes them easy to handle and house. Due to their superlative neuroregenerative potential, the diminished time between intervention and evaluation of outcomes can result in relatively lower costs11,12. Other advantages of its use include morphological similarities to human nerve fibers and the high number of comparative/historic studies13. Although the latter should be approached cautiously, as a wide variety of different outcome measures between studies makes it difficult to compare results14,15,16,17,18.
Outcome measures to assess nerve regeneration range from electrophysiology to histomorphometry, but these methods imply a correlation but do not necessarily directly measure the return of motor function14,15. Regenerating nerve fibers might not make appropriate connections which can cause an overestimation of the number of functional connections14,15,19,20. The best and clinically most relevant measurement to demonstrate correct reinnervation of end organs remains assessment of muscle function21,22,23. Creating motor function assessment tools for animal models is, however, challenging. Medinaceli et al. first described the walking track analysis, which has since been the most frequently used method to evaluate functional recovery in experimental peripheral nerve studies21,24,25,26,27,28. The walking track analysis quantifies the sciatic functional index (SFI) based on measurements of pawprints from walking rats21,29. Major limitations of the walking track analysis, such as toe contractures, automutilation, smearing of the print and poor correlation with other measures of reinnervation, have necessitated the use of other parameters for quantification of functional recovery30,31.
In previous studies in Lewis rats32 and New Zealand rabbits33, we validated the isometric tetanic force (ITF) measurement for the tibialis anterior (TA) muscle and demonstrated its effectiveness in the evaluation of muscle recovery after different types of nerve repair34,35,36,37,38,39. The TA muscle is well suited because of its relatively large size, innervation by the peroneal branch of the sciatic nerve and well elucidated biochemical properties40,41,42,43. When muscle length (preload force) and electrical parameters are optimized the ITF provides a side-to-side variability of 4.4% and 7.5% in rats32 and rabbits33, respectively.
This article provides a detailed protocol of the ITF measurement in the rat sciatic nerve model, including a thorough description of the necessary pre-surgical planning, surgical approach and dissection of the common peroneal nerve and the distal TA muscle tendon. Using predetermined values for the stimulus intensity and duration, the optimal muscle length and stimulus pulse frequency will be defined. With these four parameters, the ITF can subsequently be consistently and accurately measured.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.9% Sodium Chloride</td>
			<td>Baxter Healthcare Corporation, Deerfield, IL, USA</td>
			<td>G130203</td>
			<td></td>
		</tr>
		<tr>
			<td>1 mm Kirshner wires</td>
			<td>Pfizer Howmedica, Rutherford, NJ</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Adson Tissue Forceps</td>
			<td>ASSI, Westbury, NY, USA</td>
			<td>MTK-6801226</td>
			<td></td>
		</tr>
		<tr>
			<td>Bipolar electrode cables</td>
			<td>Grass Instrument, Quincy, MA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Bipolar stimulator device</td>
			<td>Grass SD9, Grass Instrument, Quincy, MA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton-tip Applicators</td>
			<td>Cardinal Health, Waukegan, IL, USA</td>
			<td>C15055-006</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved Mosquito forceps</td>
			<td>ASSI, Westbury, NY, USA</td>
			<td>MTK-1201112</td>
			<td></td>
		</tr>
		<tr>
			<td>Force Transducer MDB-2.5</td>
			<td>Transducer Techniques, Temecula, CA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Gauze Sponges 4x4</td>
			<td>Covidien, Mansfield, MA, USA</td>
			<td>2733</td>
			<td></td>
		</tr>
		<tr>
			<td>Ground cable</td>
			<td>Grass Instrument, Quincy, MA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane chamber</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Custom-made</td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>Ketalar, Par Pharmaceutical, Chestnut, NJ</td>
			<td>42023-115-10</td>
			<td></td>
		</tr>
		<tr>
			<td>LabView Software</td>
			<td>National Instruments, Austin, TX</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Loop</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Custom-made</td>
		</tr>
		<tr>
			<td>Microsurgical curved forceps</td>
			<td>ASSI, Westbury, NY, USA</td>
			<td>JFA-5B</td>
			<td></td>
		</tr>
		<tr>
			<td>Microsurgical scissors</td>
			<td>ASSI, Westbury, NY, USA</td>
			<td>SAS-15R-8-18</td>
			<td></td>
		</tr>
		<tr>
			<td>Microsurgical straight forceps</td>
			<td>ASSI, Westbury, NY, USA</td>
			<td>JF-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Retractor</td>
			<td>ASSI, Westbury, NY, USA</td>
			<td>AG-124426</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Blade No. 15</td>
			<td>Bard-Parker, Aspen Surgical, Caledonia, MI, USA</td>
			<td>371115</td>
			<td></td>
		</tr>
		<tr>
			<td>Slim Body Skin Stapler</td>
			<td>Covidien, Mansfield, MA, USA</td>
			<td>8886803512</td>
			<td></td>
		</tr>
		<tr>
			<td>Subminiature electrode</td>
			<td>Harvard Apparatus, Holliston, MA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Nerve Stimulator</td>
			<td>Checkpoint Surgical LCC, Cleveland, OH, USA</td>
			<td>9094</td>
			<td></td>
		</tr>
		<tr>
			<td>Terrell Isoflurane</td>
			<td>Piramal Critical Care Inc., Bethlehem, PA, USA</td>
			<td>H961J19A</td>
			<td></td>
		</tr>
		<tr>
			<td>Testing platform</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Custom-made</td>
		</tr>
		<tr>
			<td>Tetontomy Scissors</td>
			<td>ASSI, Westbury, NY, USA</td>
			<td>ASIM-187</td>
			<td></td>
		</tr>
		<tr>
			<td>Traceable Big-Digit Timer/Stopwatch</td>
			<td>Fisher Scientific, Waltham, MA, USA</td>
			<td>S407992</td>
			<td></td>
		</tr>
		<tr>
			<td>USB-6009 multifunctional I/O data acquisition (DAQ) device</td>
			<td>National Instruments, Austin, TX</td>
			<td>779026-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum Base Holder</td>
			<td>Noga Engineering &#38; Technology Ltd., Shlomi, Isreal</td>
			<td>N/A</td>
			<td>Attached clamp is custom-made</td>
		</tr>
		<tr>
			<td>Weight (10 g)</td>
			<td>Denver Instruments, Denver, CO, USA</td>
			<td>820010.4</td>
			<td></td>
		</tr>
		<tr>
			<td>Weight (20 g)</td>
			<td>Denver Instruments, Denver, CO, USA</td>
			<td>820020.4</td>
			<td></td>
		</tr>
		<tr>
			<td>Weight (50 g)</td>
			<td>Denver Instruments, Denver, CO, USA</td>
			<td>820050.4</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>Xylamed, Bimeda MTC Animal Health, Cambridge, Canada</td>
			<td>1XYL002</td>
			<td></td>
		</tr>
	</tbody>
</table>

maximum isometric tetanic force measurement of the tibialis anterior muscle in the rat

Traumatic nerve injuries result in substantial functional loss and segmental nerve defects often necessitate the use of autologous interposition nerve grafts. Due to their limited availability and associated donor side morbidity, many studies in the field of nerve regeneration focus on alternative techniques to bridge a segmental nerve gap. In order to investigate the outcomes of surgical or pharmacological experimental treatment options, the rat sciatic nerve model is often used as a bioassay. There are a variety of outcome measurements used in rat models to determine the extent of nerve regeneration. The maximum output force of the target muscle remains the most relevant outcome for clinical translation of experimental therapies. Isometric force measurement of tetanic muscle contraction has previously been described as a reproducible and valid technique for evaluating motor recovery after nerve injury or repair in both rat and rabbit models. In this video, we will provide a step-by-step instruction of this invaluable procedure for assessment of functional recovery of the tibialis anterior muscle in a rat sciatic nerve defect model using optimized parameters. We will describe the necessary pre-surgical preparations in addition to the surgical approach and dissection of the common peroneal nerve and tibialis anterior muscle tendon. The isometric tetanic force measurement technique will be detailed. Determining the optimal muscle length and stimulus pulse frequency is explained and measuring the maximum tetanic muscle contraction is demonstrated.

Five parameters are used to measure the ITF measurement. These include muscle tension (preload force), stimulus intensity (voltage), stimulus pulse frequency, stimulus duration of 0.4 ms and a delay of 2 ms. Prior to measuring the ITF, the optimal muscle tension has to be determined using two single twitch muscle contractions at an intensity of 2 V during the parameter test. These stimuli cause dorsiflexion of the paw and produce an output signal on the graph in the VI (Figure 5). These sing...

Watch this Scientific Journal Video about Maximum Isometric Tetanic Force Measurement of the Tibialis Anterior Muscle in the Rat at JoVE.com

Maximum Isometric Tetanic Force Measurement of the Tibialis Anterior Muscle in the Rat

Evaluation of motor recovery remains the benchmark outcome measure in experimental peripheral nerve studies. The isometric tetanic force measurement of the tibialis anterior muscle in the rat is an invaluable tool to assess functional outcomes after reconstruction of sciatic nerve defects. The methods and nuances are detailed in this article.

Traumatic nerve injuries result in substantial functional loss and segmental nerve defects often necessitate the use of autologous interposition nerve ...

maximum-isometric-tetanic-force-measurement-tibialis-anterior-muscle

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Neuroscience

4.4K Views.  Mayo Clinic. Evaluation of motor recovery remains the benchmark outcome measure in experimental peripheral nerve studies. The isometric tetanic force measurement of the tibialis anterior muscle in the rat is an invaluable tool to assess functional outcomes after reconstruction of sciatic nerve defects. The methods and nuances are detailed in this article.

Video: Maximum Isometric Tetanic Force Measurement of the Tibialis Anterior Muscle in the Rat

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

An In Vitro Adult Mouse Muscle-nerve Preparation for Studying the Firing Properties of Muscle Afferents

Muscle sensory neurons innervating muscle spindles and Golgi tendon organs encode length and force changes essential to proprioception. Additional afferent fibers monitor other characteristics of the muscle environment, including metabolite buildup, temperature, and nociceptive stimuli. Overall, abnormal activation of sensory neurons can lead to movement disorders or chronic pain syndromes. We describe the isolation of the extensor digitorum longus (EDL) muscle and nerve for in vitro study of stretch-evoked afferent responses in the adult mouse. Sensory activity is recorded from the nerve with a suction electrode and individual afferents can be analyzed using spike sorting software. In vitro preparations allow for well controlled studies on sensory afferents without the potential confounds of anesthesia or altered muscle perfusion. Here we describe a protocol to identify and test the response of muscle spindle afferents to stretch. Importantly, this preparation also supports the study of other subtypes of muscle afferents, response properties following drug application and the incorporation of powerful genetic approaches and disease models in mice.

An In Vitro Adult Mouse Muscle-nerve Preparation for Studying the Firing Properties of Muscle Afferents

Muscle sensory neurons are involved in proprioceptor signaling and also report on metabolic state and injury related events. We describe an adult mouse in vitro muscle-nerve preparation for studies on stretch-activated muscle afferents.

Muscle sensory neurons innervating muscle spindles and Golgi tendon organs encode length and force changes essential to proprioception. Additional ...

Analysis of Gene Expression Changes in the Rat Hippocampus After Deep Brain Stimulation of the Anterior Thalamic Nucleus

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective treatment for several movement disorders that have failed to respond to medication. Recent progress in the field of DBS surgery has begun to extend the application of this surgical technique to other conditions as diverse as morbid obesity, depression and obsessive compulsive disorder. Despite these expanding indications, little is known about the underlying physiological mechanisms that facilitate the beneficial effects of DBS surgery. One approach to this question is to perform gene expression analysis in neurons that receive the electrical stimulation. Previous studies have shown that neurogenesis in the rat dentate gyrus is elicited in DBS targeting of the anterior nucleus of the thalamus1. DBS surgery targeting the ATN is used widely for treatment refractory epilepsy. It is thus of much interest for us to explore the transcriptional changes induced by electrically stimulating the ATN. In this manuscript, we describe our methodologies for stereotactically-guided DBS surgery targeting the ATN in adult male Wistar rats. We also discuss the subsequent steps for tissue dissection, RNA isolation, cDNA preparation and quantitative RT-PCR for measuring gene expression changes. This method could be applied and modified for stimulating the basal ganglia and other regions of the brain commonly clinically targeted. The gene expression study described here assumes a candidate target gene approach for discovering molecular players that could be directing the mechanism for DBS.

The mechanism underlying the therapeutic effects of Deep Brain Stimulation (DBS) surgery needs investigation. The methods presented in this manuscript describe an experimental approach to examine the cellular events triggered by DBS by analyzing the gene expression profile of candidate genes that can facilitate neurogenesis post DBS surgery.

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective ...

Generation of Local CA1 &#947; Oscillations by Tetanic Stimulation

Neuronal network oscillations are important features of brain activity in health and disease and can be modulated by a range of clinically used drugs. A protocol is provided to generate a model for studying CA1 &#947; oscillations (20 - 80 Hz). These &#947; oscillations are stable for at least 30 min&#160;and depend upon excitatory and inhibitory synaptic activity in addition to activation of pacemaker currents. Tetanically stimulated oscillations have a number of reproducible and easily quantifiable characteristics including spike count, oscillation duration, latency and frequency that report upon the network state. The advantages of the electrically stimulated oscillations include stability, reproducibility and episodic acquisition enabling robust characterization of network function. This model of CA1 &#947; oscillations can be used to study cellular mechanisms and to systematically investigate how neuronal network activity is altered in disease and by drugs. Disease state pharmacology can be readily incorporated by the use of brain slices from genetically modified or interventional animal models to enable selection of drugs that specifically target disease mechanisms.

Generation of Local CA1 &amp;#947; Oscillations by Tetanic Stimulation

Oscillations are fundamental network properties and are modulated by disease and drugs. Studying brain-slice oscillations allows characterization of isolated networks under controlled conditions. Protocols are provided for the preparation of acute brain slices for evoking CA1 &#947; oscillations.

Neuronal network oscillations are important features of brain activity in health and disease and can be modulated by a range of clinically used drugs. A ...

Visual Evoked Potential Recording in a Rat Model of Experimental Optic Nerve Demyelination

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to monitor the disease course in the follow-up period. Changes in the VEP parameters closely correlate with pathological damage in the optic nerve. This protocol provides a detailed description about the rodent model of optic nerve microinjection, in which a partial demyelination lesion is produced in the optic nerve. VEP recording techniques are also discussed. Using skull implanted electrodes, we are able to acquire reproducible intra-session and between-session VEP traces. VEPs can be recorded on individual animals over a period of time to assess the functional changes in the optic nerve longitudinally. The optic nerve demyelination model, in conjunction with the VEP recording protocol, provides a tool to investigate the disease processes associated with demyelination and remyelination, and can potentially be employed to evaluate the effects of new remyelinating drugs or neuroprotective therapies.

Focal demyelination is induced in the optic nerve using lysolecithin microinjection. Visual evoked potentials are recorded via skull electrodes implanted over the visual cortex to examine the signal conduction along the visual pathway in vivo. This protocol details the surgical procedures underlying electrode implantation and optic nerve microinjection.

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to ...

Sublimation of DAN Matrix for the Detection and Visualization of Gangliosides in Rat Brain Tissue for MALDI Imaging Mass Spectrometry

Sample preparation is key for optimal detection and visualization of analytes in Matrix-assisted Laser Desorption/Ionization (MALDI) Imaging Mass Spectrometry (IMS) experiments. Determining the appropriate protocol to follow throughout the sample preparation process can be difficult as each step must be optimized to comply with the unique characteristics of the analytes of interest. This process involves not only finding a compatible matrix that can desorb and ionize the molecules of interest efficiently, but also selecting the appropriate matrix deposition technique. For example, a wet matrix deposition technique, which entails dissolving a matrix in solvent, is superior for desorption of most proteins and peptides, whereas dry matrix deposition techniques are particularly effective for ionization of lipids. Sublimation has been reported as a highly efficient method of dry matrix deposition for the detection of lipids in tissue by MALDI IMS due to the homogeneity of matrix crystal deposition and minimal analyte delocalization as compared to many wet deposition methods 1,2. Broadly, it involves placing a sample and powdered matrix in a vacuum-sealed chamber with the samples pressed against a cold surface. The apparatus is then lowered into a heated bath (sand or oil), resulting in sublimation of the powdered matrix onto the cooled tissue sample surface. Here we describe a sublimation protocol using 1,5-diaminonaphthalene (DAN) matrix for the detection and visualization of gangliosides in the rat brain using MALDI IMS.

A protocol for the sublimation of DAN matrix onto rat brain tissue for the detection of gangliosides using MALDI Imaging Mass Spectrometry is presented.

Sample preparation is key for optimal detection and visualization of analytes in Matrix-assisted Laser Desorption/Ionization (MALDI) Imaging Mass ...

The Use of Trace Eyeblink Classical Conditioning to Assess Hippocampal Dysfunction in a Rat Model of Fetal Alcohol Spectrum Disorders

Neonatal rats were administered a relatively high concentration of ethyl alcohol (11.9% v/v) during postnatal days 4-9, a time when the fetal brain undergoes rapid organizational change and is similar to accelerated brain changes that occur during the third trimester in humans. This model of fetal alcohol spectrum disorders (FASDs) produces severe brain damage, mimicking the amount and pattern of binge-drinking that occurs in some pregnant alcoholic mothers. We describe the use of trace eyeblink classical conditioning (ECC), a higher-order variant of associative learning, to assess long-term hippocampal dysfunction that is typically seen in alcohol-exposed adult offspring. At 90 days of age, rodents were surgically prepared with recording and stimulating electrodes, which measured electromyographic (EMG) blink activity from the left eyelid muscle and delivered mild shock posterior to the left eye, respectively. After a 5 day recovery period, they underwent 6 sessions of trace ECC to determine associative learning differences between alcohol-exposed and control rats. Trace ECC is one of many possible ECC procedures that can be easily modified using the same equipment and software, so that different neural systems can be assessed. ECC procedures in general, can be used as diagnostic tools for detecting neural pathology in different brain systems and different conditions that insult the brain.

Trace eyeblink classical conditioning (ECC) was used to assess hippocampal-dependent associative learning in adult rats that were administered a high concentration (11.9% v/v) of alcohol during early neonatal brain development. In general, ECC procedures are sound diagnostic tools for detecting brain dysfunction across many psychological and biomedical settings.

Neonatal rats were administered a relatively high concentration of ethyl alcohol (11.9% v/v) during postnatal days 4-9, a time when the fetal brain ...

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and latency detect chronic axon loss and demyelination, respectively. Axonal excitability techniques &#34;by threshold tracking&#34; expand upon these measures by providing information regarding the activity of ion channels, pumps and exchangers that relate to acute function and may precede degenerative events. As such, the use of axonal excitability in animal models of neurological disorders may provide a useful in vivo measure to assess novel therapeutic interventions. Here we describe an experimental setup for multiple measures of motor axonal excitability techniques in the rat ulnar nerve.
The animals are anesthetized with isoflurane and carefully monitored to ensure constant and adequate depth of anesthesia. Body temperature, respiration rate, heart rate and saturation of oxygen in the blood are continuously monitored. Axonal excitability studies are performed using percutaneous stimulation of the ulnar nerve and recording from the hypothenar muscles of the forelimb paw. With correct electrode placement, a clear compound muscle action potential that increases in amplitude with increasing stimulus intensity is recorded. An automated program is then utilized to deliver a series of electrical pulses which generate 5 specific excitability measures in the following sequence: stimulus response behavior, strength duration time constant, threshold electrotonus, current-threshold relationship and the recovery cycle.
Data presented here indicate that these measures are repeatable and show similarity between left and right ulnar nerves when assessed on the same day. A limitation of these techniques in this setting is the effect of dose and time under anesthesia. Careful monitoring and recording of these variables should be undertaken for consideration at the time of analysis.

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Axonal excitability techniques provide a powerful tool to examine pathophysiology and biophysical changes that precede irreversible degenerative events. This manuscript demonstrates the use of these techniques on the ulnar nerve of anesthetized rats.

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and ...

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings provide a sensitive approach to measure nerve conduction in humans and rodent models. To broaden the technical possibilities for electromyography in mice, the measurement of compound muscle action potentials (CMAPs) from the brachial plexus nerve in the forelimb using needle electrodes is described here. CMAP recordings after stimulating the sciatic nerve in hindlimbs have been previously described. The newly introduced method here allows for the evaluation of the nerve conductivity at an additional site, and thus provides a more profound overview of the neuromuscular functionality. The technique provides information on both the relative number of functional axons and the myelination level. Thereby, this method can be applied to assess both axonal diseases as well as demyelinating conditions. This minimally invasive method does not require extraction of the nerve and therefore it is suitable for repeated measurements for longitudinal follow-up in the same animal. Similar recordings are performed in clinical setups to emphasize the translational relevance of the method.

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

The measurement of nerve conduction is a useful tool to assess mouse models of neurodegeneration but it is frequently only applied to stimulate the sciatic nerve in hindlimbs. Here, we describe a technique to measure compound muscle action potential (CMAP) in vivo in the mouse forelimb muscles innervated by the brachial plexus.

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings ...

Muscle Velocity Recovery Cycles to Examine Muscle Membrane Properties

Although conventional nerve conduction studies (NCS) and electromyography (EMG) are suitable for the diagnosis of neuromuscular disorders, they provide limited information about muscle fiber membrane properties and underlying disease mechanisms. Muscle velocity recovery cycles (MVRCs) illustrate how the velocity of a muscle action potential depends on the time after a preceding action potential. MVRCs are closely related to changes in membrane potential that follow an action potential, thereby providing information about muscle fiber membrane properties. MVRCs may be recorded quickly and easily by direct stimulation and recording from multi-fiber bundles in vivo. MVRCs have been helpful in understanding disease mechanisms in several neuromuscular disorders. Studies in patients with channelopathies have demonstrated the different effects of specific ion channel mutations on muscle excitability. MVRCs have been previously tested in patients with neurogenic muscles. In this prior study, muscle relative refraction period (MRRP) was prolonged, and early supernormality (ESN) and late supernormality (LSN) were reduced in patients compared to healthy controls. Thereby, MVRCs can provide in vivo evidence of membrane depolarization in intact human muscle fibers that underlie their reduced excitability. The protocol presented here describes how to record MVRCs and analyze the recordings. MVRCs can serve as a fast, simple, and useful method for revealing disease mechanisms across a broad range of neuromuscular disorders.

Presented here is a protocol for the recording of muscle velocity recovery cycles (MVRCs), a new method of examining muscle membrane properties. MVRCs enable in vivo assessment of muscle membrane potential and alterations in muscle ion channel function in relation to pathology, and it enables the demonstration of muscle depolarization in neurogenic muscles.

Although conventional nerve conduction studies (NCS) and electromyography (EMG) are suitable for the diagnosis of neuromuscular disorders, they provide ...