This work was funded by the National Institutes of Health (NIH) K12 for Physical and Occupational Therapists-Comprehensive Opportunities in Rehabilitation Research Training (K12 HD055931), the Foundation for Physical Therapy and the Pittsburgh Claude D. Pepper Older Americans Independence Center (P30 AG024827).

1. Electrode Preparation
<ul>
	<li>Cut a 1 cm x 1 cm section of the vector circuit board.</li>
	<li>Stabilize 2 wirewrap pins using a vice grip, with pins spaced approximately 3.5 mm apart. The bifurcated ends of the pins should be facing up.</li>
	<li>Bend wirewrap pins at a 90&deg; angle, approximately halfway along their length, using needlenose pliers.</li>
	<li>Expose the copper wires of connectors, approximately 7 cm from lead connection, by stripping the wire insulation.</li>
	<li>Solder one of the copper wires to the bifurcated end of one the wirewrap pins. Repeat soldering step for the other wirewrap pin.</li>
	<li>Pull shrink tubing at soldered connection between copper wires and gold pins in order to insulate and stabilize. Heat the tubing using the soldering tip.</li>
	<li>Insert the unsoldered tips of the wirewrap pins into adjacent openings of the breadboard circuit board.</li>
	<li>Ensure that the wirewrap pins are parallel to one another and stick through the breadboard such that the tips are located the same distance when placed through the board.</li>
	<li>Embed soldered ends of the pins in clear epoxy. Allow the epoxy to cure for approximately 10 minutes, or until completely dry.</li>
	<li>Repeat the application of epoxy until the pins and vector circuit board are completely embedded in order to provide structural integrity and strain relief for the lead wires.</li>
	<li>Sand down the epoxy using a metal file to expose the tips of the gold pins.</li>
	<li>Use a multimeter to ensure the two leads are not electrically shorted and that the lead wires have proper continuity</li>
	<li>Attach the electrode leads from the NMES device to female 2-way wire connector (Pomona patch cord).</li>
</ul>
2. Animal Preparation
<ul>
	<li>Animals are anesthetized via inhalation using 2% isofluorane (Abbott Laboratories, North Chicago, IL) in 100% O2 gas. The animal should remain anesthetized throughout the NMES session.</li>
	<li>Before starting the NMES protocol perform a toe pinch to ensure that the animal is fully anesthetized.</li>
	<li>Animal should be regularly checked for response to stimulation in order to ensure the anesthetic level is sufficient. The anesthesia should be adjusted as needed. Movement other than that of the stimulated leg, changes in respiratory depth, and color of mucous membranes should all be assessed regularly to monitor depth of anesthesia.</li>
	<li>Position the animal in lateral recumbency.</li>
	<li>Apply ophthalmic lubricant to the eyes in order to prevent corneal drying during the procedure.</li>
	<li>Shave the hindlimb area to be treated, and wipe with alcohol.</li>
	<li>Wipe the electrode with 3% bleach, then rinse with water.</li>
	<li>Apply a layer of conducting gel over the site where the electrode will be applied. Reapply as necessary throughout the NMES protocol to ensure current flow between the electrode and the skin.</li>
	<li>A heat lamp or circulating water blanket should be used to maintain normal core body ranges.</li>
	<li>Note: All procedures have been reviewed and approved by the University of Pittsburgh Institutional Animal Care and Use Committee and performed in PHS assured and AAALAC Int. accredited program and facilities.</li>
</ul>
3. Stimulation
<ul>
	<li>Electrode placement:
	<ul>
		<li>For stimulation of anterior compartment muscles, including the tibialis anterior and the extensor digitorum longus muscle, place the surface electrode directly over the animal's deep fibular nerve, which is a distal branch off of the common peroneal nerve, and is located just anterior to the the fibular head.</li>
		<li>For stimulation of anterior compartment muscles, placement of the electrode is confirmed when stimulation elicits full ankle dorsiflexion and full extension of the digits. On the other hand, dorsiflexion, in the absence of digit extension suggests that only the tibialis anterior muscle is being stimulated. Such a targeted contractile response may be desirable, depending on study design.</li>
		<li>The muscle contraction is divided into 2 phases of stimulation: a free moving, concentric phase, which occurs during the initial ~0.5 seconds. During this first phase, the paw moves from a resting position to maximal dorsiflexion and digit extension. The second phase of stimulation is an isometric contraction sustained at end-range dorsiflexion and digit extension.</li>
	</ul>
	</li>
	<li>Stimulation parameters:
	<ul>
		<li>This NMES model utilizes the 300 PV Empi multifunction electrotherapy device, which provides 2 channels of conventional NMES (see Table). For the purposes of this model, only 1 channel is used.</li>
		<li>Parameters used include symmetric waveform, a pulse duration of 150 &mu;s and a frequency of 50Hz. Stimulation time is set to 5 seconds, with a 0.5 second ramp up and a 0.5 second ramp down. This will allow the muscle to more gradually acclimate to the stimulation. In the current protocol, off time in between contractions was set to 10 seconds, but this may be adjusted depending on the desired effect. Decreased off times will result in a more rapid initiation of muscle fatigue. These stimulation parameters were based on clinical protocols designed to enhance muscle strength using neuromuscular electrical stimulation, without inducing significant skeletal muscle damage 1, 15, 16.</li>
		<li>Mice complete two sets of 10 contractions, with a 5 minute rest in between sets.</li>
		<li>For adult animals, NMES intensity is typically initiated at 9 mA. From our experience, this is approximate to the maximal starting intensity that does not induce a noticeable gait impairment immediately following stimulation. For subsequent NMES sessions, the intensity is increased by 1 mA every time the animals are able to complete 20 full dorsiflexions.</li>
	</ul>
	</li>
	<li>Following completion of the stimulation protocol and recovery from anesthesia, animals typically demonstrate a normal gait and posture. Gait should remain unimpaired throughout the duration of the NMES program.</li>
</ul>
4. Representative Results
The NMES protocol described in this article has been implemented in several mouse strains, including: Wild Type (B6/10), B6.SCID and mdx/SCID mice. Representative results of NMES performed over 4 weeks (20 sessions) in 3-5 month old mdx/SCID are presented.
Animals were humanely euthanized by cervical dislocation while under anesthesia. The tibialis anterior muscle was harvested and immediately frozen in 2 methyl-butane pre-cooled in liquid nitrogen, and stored at -80 &deg;C. Serial cross-sections (10 &mu;m) were obtained and mounted onto slides. Statistical analyses were performed using standard statistical software packages (SPSS, v19.0 software). First, Levene's test was used to assess whether there was equality of variances. Independent samples t-test was then performed to investigate differences between NMES and Control groups.
Hematoxylin and Eosin (H&amp;E) stains were performed to investigate whether the NMES protocol would result in increased injury in the dystrophic muscle. One section was selected, and pictures were obtained using a light microscope (Nikon Eclipse E800; Nikon, Japan). The total number of fibers and the number of fibers with centrally located nuclei were manually counted using the National Institutes of Health (NIH) - developed image analysis software, Image J. There was no significant increase in the regeneration index (number of centrally nucleated fibers/total number of fibers) in the animals submitted to NMES, as compared to controls (p=0.802; Figure 1). This suggests that the application of NMES does not increase the degeneration-regeneration cascade observed in dystrophic animals, and is therefore not likely to be inducing an increased muscle injury.
Immunofluorescent staining for CD31 was performed to investigate the effect of the 4-week NMES protocol on muscle vascularity. Briefly, muscle sections were fixed in 4% formalin and blocked using 5% Horse Serum (HS). Sections were incubated with a rat anti-mouse primary antibody (1:300 dilution in 5% HS) and treated with a 555-labeled goat anti-rat secondary antibody (1:300 dilution in 5% HS). To quantify the total number of CD31 positive cells, one section was selected, and photographed using fluorescent microscopy (Nikon Eclipse E800; Japan). The total number of capillaries was manually counted using Northern Eclipse Software (Empix Imaging Inc.). There was a significant increase in the number of CD31 positive cells in the animals submitted to NMES when compared to controls (p&lt;0.01; Figure 2), indicating that the NMES protocol as described promotes skeletal muscle angiogenesis.
In situ contractile testing: Four weeks after completion of an NMES protocol, contractile testing of the anterior compartment muscles was performed using an in situ testing apparatus (Model 809B, Aurora Scientific Inc, Canada), stimulator (Model 701C, Aurora Scientific Inc, Canada), and force transducer (Aurora Scientific Inc, Canada). Briefly, the peroneal nerve of anesthetized animals was isolated through a small incision lateral to the knee. Mice were then placed supine on a platform and the foot being tested was positioned on the footplate. The hindlimb used for testing was stabilized with cloth tape on the knee and foot. Muscles were stimulated using a hook electrodes inserted beneath the skin. Muscle tetanic contraction was tested at a frequency of 150Hz for 350ms to investigate whether the 4-week NMES protocol would induce improvements in muscle force. Results were normalized to wet muscle weight. There was an approximately 30% increase in the tetanic contraction of animals submitted to NMES, when compared to controls (p=0.005) (Figure 3), suggesting the protocol described improves skeletal muscle strength in mdx/SCID animals.
<img alt="Figure 1" src="/files/ftp_upload/3914/3914fig1.jpg" /> 
Figure 1. Hematoxylin &amp; Eosin skeletal muscle staining of dystrophic (mdx)/ immunodeficient mice (4-5 months)(10x magnification): A) untreated controls, B) after 4 weeks of NMES.
<img alt="Figure 2" src="/files/ftp_upload/3914/3914fig2.jpg" /> 
Figure 2. CD31 immunofluorescence (red), as a marker of skeletal muscle vascularity, in the skeletal muscle of dystrophic (mdx)/ immunodeficient mice (4-5 months)(20x magnification) A) untreated controls, B) after 4 weeks of NMES.
<img alt="Figure 3" src="/files/ftp_upload/3914/3914fig3.jpg" /> 
Figure 3. Contractile testing of the anterior compartment muscles in control and animals treated with NMES for 4 weeks. mNm= milli Newton meters. <a href="https://www.jove.com/files/ftp_upload/3914/3914fig3large.jpg">Click here to view larger figure</a>.

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We have nothing to disclose.

These results suggest that our developed murine model of NMES promotes skeletal muscle angiogenesis (Figure 2) and strengthening (Figure 3), but does not induce skeletal muscle damage (Figure 1).
It should be noted that the stimulation parameters described here have been designed to induce a muscle overloading to the anterior compartment muscles. Just as is the case for clinical scenarios, electrode placement may be adjusted to stimulate other muscle groups, although the muscle response to NMES may be different, depending on fiber type composition. NMES duration, total number of treatment sessions, and total number of repetitions may be modified according to study design.
As with any protocol, this method has limitations that should be noted. In the present study, stimulation was performed over the deep fibular nerve. However, in the clinic, stimulation is typically administered at the motor unit. In the animal model, stimulation of the nerve would require a lower intensity in order to elicit full muscle contraction. Another limitation of the model as presented is that we are unable to obtain information regarding comfort level during the application of NMES, given that the animals are anesthetized. Therefore, the tolerability of comparable intensities in a clinical population is difficult to assess. However, our histological results suggest our NMES model, as described, does not induce muscle injury.
The development of animals models that mimic modalities commonly implemented in the clinic provide us with useful laboratory tools that allows for an improved understanding of cellular and molecular responses to treatment interventions. In addition, such models are useful to conduct pre-clinical studies to both refine existing rehabilitation protocols and develop novel indications.

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Vector wire wrap posts with bifurcated solder terminal</td>
 <td>Newark</td>
 <td>T68</td>
 </tr>
 <tr>
 <td>Vector circuit board</td>
 <td>Newark</td>
 <td>3662-2</td>
 </tr>
 <tr>
 <td>Pomona patch cord</td>
 <td>Newark</td>
 <td>P-36-0</td>
 </tr>
 <tr>
 <td>5 minute epoxy</td>
 <td>VO Baker Co.</td>
 <td>4001</td>
 </tr>
 <tr>
 <td>Spectra 360 Electrode Gel</td>
 <td>Milliken Medical</td>
 <td>MR41217</td>
 </tr>
 <tr>
 <td>Portable neuromuscular stimulation device</td>
 <td>EMPI</td>
 <td>300pv</td>
 </tr>
 </tbody>
</table>

a murine model of muscle training by neuromuscular electrical stimulation

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional capacity. Transcutaneous surface stimulation of skeletal muscle involves a current flow between a cathode and an anode, thereby inducing excitement of the motor unit and the surrounding muscle fibers. 
NMES is an attractive modality to evaluate skeletal muscle adaptive responses for several reasons. First, it provides a reproducible experimental model in which physiological adaptations, such as myofiber hypertophy and muscle strengthening6, angiogenesis7-9, growth factor secretion9-11, and muscle precursor cell activation12 are well documented. Such physiological responses may be carefully titrated using different parameters of stimulation (for Cochrane review, see 13). In addition, NMES recruits motor units non-selectively, and in a spatially fixed and temporally synchronous manner14, offering the advantage of exerting a treatment effect on all fibers, regardless of fiber type. Although there are specified contraindications to NMES in clinical populations, including peripheral venous disorders or malignancy, for example, NMES is safe and feasible, even for those who are ill and/or bedridden and for populations in which rigorous exercise may be challenging. 
Here, we demonstrate the protocol for adapting commercially available electrodes and performing a NMES protocol using a murine model. This animal model has the advantage of utilizing a clinically available device and providing instant feedback regarding positioning of the electrode to elicit the desired muscle contractile effect. For the purpose of this manuscript, we will describe the protocol for muscle stimulation of the anterior compartment muscles of a mouse hindlimb.

Watch this Scientific Journal Video about A Murine Model of Muscle Training by Neuromuscular Electrical Stimulation at JoVE.com

A Murine Model of Muscle Training by Neuromuscular Electrical Stimulation

A murine model of neuromuscular electrical stimulation (NMES), a safe and inexpensive clinical modality, to the anterior compartment muscles is described. This model has the advantage of modifying a readily available clinical device for the purpose of eliciting targeted and specific muscle contractions in mice.

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional ...

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20.7K ビュー.  University of Pittsburgh. 前部コンパートメントの筋肉への神経筋電気刺激（NMES）、安全かつ安価な治療モダリティのマウスモデルは、記載されています。このモデルは、マウスでターゲットを絞った特定の筋肉の収縮を誘発する目的のために容易に利用可能な臨床デバイスを変更できるという利点があります。

ビデオ: A Murine Model of Muscle Training by Neuromuscular Electrical Stimulation

A murine model of neuromuscular electrical stimulation (NMES), a safe and inexpensive clinical modality, to the anterior compartment muscles is described.  This model has the advantage of modifying a readily available clinical device for the purpose of eliciting targeted and specific muscle contractions in...

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Dissection of the Transversus Abdominis Muscle for Whole-mount Neuromuscular Junction Analysis

In this video we demonstrate a protocol for dissection of the transversus abdominis muscle of the mouse and use immunofluorescence and microscopy to visualize neuromuscular...

Dissection of the Transversus Abdominis Muscle for Whole-mount Neuromuscular Junction Analysis

Genetically-modified mice have emerged as a preferable animal model to study the molecular mechanisms underlying conduction abnormalities, atrial and ventricular arrhythmias, and sudden cardiac death.1 Intracardiac pacing studies can be performed in mice using a 1.1F octapolar catheter inserted into the jugular vein, and advanced into the right atrium and ventricle. Here, we illustrate the steps involved in performing programmed electrical stimulation in mice. Surface ECG and intracardiac electrograms are recorded simultaneously in the atria, atrioventricular junction, and ventricular myocardium, whereas intracardiac pacing of the atrium is performed using an external stimulator. Thus, programmed electrical stimulation in mice provides unique opportunities to explore molecular mechanisms underlying conduction defects and cardiac arrhythmias.

Programmed electrical stimulation provides the ability to determine conduction properties of the heart, and the possibility to induce and terminate cardiac arrhythmias using various pacing protocols. Using a transvenous catheter, intracardiac electrogram recordings can be obtained in mice following programmed electrical stimulation protocols to identify arrhythmogenic substrates.

Genetically-modified mice have emerged as a preferable animal model to study the molecular mechanisms underlying conduction abnormalities, atrial and ...

Transcranial magnetic stimulation (TMS) has been in use for more than 20 years 1, and has grown exponentially in popularity over the past decade. While the use of TMS has expanded to the study of many systems and processes during this time, the original application and perhaps one of the most common uses of TMS involves studying the physiology, plasticity and function of the human neuromuscular system. Single pulse TMS applied to the motor cortex excites pyramidal neurons transsynaptically 2 (Figure 1) and results in a measurable electromyographic response that can be used to study and evaluate the integrity and excitability of the corticospinal tract in humans 3. Additionally, recent advances in magnetic stimulation now allows for partitioning of cortical versus spinal excitability 4,5. For example, paired-pulse TMS can be used to assess intracortical facilitatory and inhibitory properties by combining a conditioning stimulus and a test stimulus at different interstimulus intervals 3,4,6-8. In this video article we will demonstrate the methodological and technical aspects of these techniques. Specifically, we will demonstrate single-pulse and paired-pulse TMS techniques as applied to the flexor carpi radialis (FCR) muscle as well as the erector spinae (ES) musculature. Our laboratory studies the FCR muscle as it is of interest to our research on the effects of wrist-hand cast immobilization on reduced muscle performance6,9, and we study the ES muscles due to these muscles clinical relevance as it relates to low back pain8. With this stated, we should note that TMS has been used to study many muscles of the hand, arm and legs, and should iterate that our demonstrations in the FCR and ES muscle groups are only selected examples of TMS being used to study the human neuromuscular system.

Transcranial magnetic stimulation (TMS) is a non-invasive tool to gain insight on the physiology and function of the human nervous system. Here, we present our TMS techniques to study cortical excitability of the upper limb and lumbar musculature.

Transcranial magnetic stimulation (TMS) has been in use for more than 20 years 1, and has grown exponentially in popularity over the past decade.  While ...

Murine Model of Wound Healing

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. Impaired wound healing, which occurs in diabetic patients for example, can lead to severe unfavorable outcomes such as amputation. There is, therefore, an increasing impetus to develop novel agents that promote wound repair. The testing of these has been limited to large animal models such as swine, which are often impractical. Mice represent the ideal preclinical model, as they are economical and amenable to genetic manipulation, which allows for mechanistic investigation. However, wound healing in a mouse is fundamentally different to that of humans as it primarily occurs via contraction. Our murine model overcomes this by incorporating a splint around the wound. By splinting the wound, the repair process is then dependent on epithelialization, cellular proliferation and angiogenesis, which closely mirror the biological processes of human wound healing. Whilst requiring consistency and care, this murine model does not involve complicated surgical techniques and allows for the robust testing of promising agents that may, for example, promote angiogenesis or inhibit inflammation. Furthermore, each mouse acts as its own control as two wounds are prepared, enabling the application of both the test compound and the vehicle control on the same animal. In conclusion, we demonstrate a practical, easy-to-learn, and robust model of wound healing, which is comparable to that of humans.

A murine model of cutaneous wound healing that can be used to assess therapeutic compounds in physiological and pathophysiological settings.

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. ...

A Murine Model of Subarachnoid Hemorrhage

In this video publication a standardized mouse model of subarachnoid hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of Willis perforation (CWp) and proven by intracranial pressure (ICP) monitoring. Thereby a homogenous blood distribution in subarachnoid spaces surrounding the arterial circulation and cerebellar fissures is achieved. Animal physiology is maintained by intubation, mechanical ventilation, and continuous on-line monitoring of various physiological and cardiovascular parameters: body temperature, systemic blood pressure, heart rate, and hemoglobin saturation. Thereby the cerebral perfusion pressure can be tightly monitored resulting in a less variable volume of extravasated blood. This allows a better standardization of endovascular filament perforation in mice and makes the whole model highly reproducible. Thus it is readily available for pharmacological and pathophysiological studies in wild type and genetically altered mice.

A standardized mouse model of subarachnoid hemorrhage by intraluminal Circle of Willis perforation is described. Vessel perforation and subarachnoid bleeding are monitored by intracranial pressure monitoring. In addition various vital parameters are recorded and controlled to maintain physiologic conditions.

In this video publication a standardized mouse model of subarachnoid  hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of  Willis ...

Reproducable Paraplegia by Thoracic Aortic Occlusion in a Murine Model of Spinal Cord Ischemia-reperfusion

Background 
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered advancements to decrease the occurrence this injury.&#160;The current model demonstrates reproducible lower&#160;extremity paralysis following thoracic aortic occlusion.
Methods 
Adult male C57BL6 mice were anesthetized with isoflurane. Through a cervicosternal incision the aorta was exposed. The descending thoracic aorta and left subclavian arteries were identified without entrance into pleural space. Skeletonization of these arteries was followed by immediate closure (Sham) or occlusion for 4 min (moderate ischemia) or 8 min (prolonged ischemia). The sternotomy and skin were closed and the mouse was transferred to warming bed for recovery.&#160; Following recovery, functional analysis was obtained at 12 hr intervals until 48 hr.
Results 
Mice that underwent sham surgery showed no observable hind&#160;limb deficit. Mice subjected to moderate ischemia for 4 min had minimal functional deficit at 12 hr followed by progression to complete paralysis at 48 hr. Mice subjected to prolonged ischemia had an immediate paralysis with no observable hind-limb movement at any point in the postoperative period. There was no observed intraoperative or post&#160;operative mortality.
Conclusion 
Reproducible lower extremity paralysis whether immediate or delayed can be achieved in a murine model. Additionally, by using a median sternotomy and careful dissection, high survival rates, and reproducibility can be achieved.

The lack of mechanistic understanding of spinal cord ischemia-reperfusion injury has hindered further adjuncts to prevent paraplegia following high risk aortic operations. Thus, the development of animal models is imperative. This manuscript demonstrates reproducible lower extremity paralysis following thoracic aortic occlusion in a murine model.

Background
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered ...

Assessment of Right Ventricular Structure and Function in Mouse Model of Pulmonary Artery Constriction by Transthoracic Echocardiography

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, the RV is significantly affected in pulmonary diseases such as pulmonary artery hypertension (PAH). In addition, the RV is remarkably sensitive to cardiac pathologies, including left ventricular (LV) dysfunction, valvular disease or RV infarction4. To understand the role of RV in the pathogenesis of cardiac diseases, a reliable and noninvasive method to access the RV structurally and functionally is essential.
A noninvasive trans-thoracic echocardiography (TTE) based methodology was established and validated for monitoring dynamic changes in RV structure and function in adult mice. To impose RV stress, we employed a surgical model of pulmonary artery constriction (PAC) and measured the RV response over a 7-day period using a high-frequency ultrasound microimaging system. Sham operated mice were used as controls. Images were acquired in lightly anesthetized mice at baseline (before surgery), day 0 (immediately post-surgery), day 3, and day 7 (post-surgery). Data was analyzed offline using software.
Several acoustic windows (B, M, and Color Doppler modes), which can be consistently obtained in mice, allowed for reliable and reproducible measurement of RV structure (including RV wall thickness, end-diastolic&#160;and end-systolic dimensions), and function (fractional area change, fractional shortening, PA peak velocity, and peak pressure gradient) in normal mice and following PAC.
Using this method, the pressure-gradient resulting from PAC was accurately measured in real-time using Color Doppler mode and was comparable to direct pressure measurements performed with a Millar high-fidelity microtip catheter. Taken together, these data demonstrate that RV measurements obtained from various complimentary views using echocardiography are reliable, reproducible and can provide insights regarding RV structure and function. This method will enable a better understanding of the role of RV cardiac dysfunction.

Right ventricle (RV) dysfunction is critical to the pathogenesis of cardiovascular disease, yet limited methodologies are available for its evaluation. Recent advances in ultrasound imaging provide a noninvasive and accurate option for longitudinal RV study. Herein, we detail a step-by-step echocardiographic method using a murine model of RV pressure overload.

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, ...

Excitotoxic Stimulation of Brain Microslices as an In vitro Model of Stroke

Examining molecular mechanisms involved in neuropathological conditions, such as ischemic stroke, can be difficult when using whole animal systems. As such, primary or &#39;neuronal-like&#39; cell culture systems are commonly utilized. While&#160;these systems are relatively easy to work with, and are useful model systems in which various functional outcomes (such as cell death) can be readily quantified, the examined outcomes and pathways in cultured immature neurons (such as excitotoxicity-mediated cell death pathways) are not necessarily the same as those observed in mature brain, or in intact tissue. Therefore, there is the need to develop models in which cellular mechanisms in mature neural tissue can be examined. We have developed an in vitro technique that can be used to investigate a variety of molecular pathways in intact nervous tissue. The technique described herein utilizes rat cortical tissue, but this technique can be adapted to use tissue from a variety of species (such as mouse, rabbit, guinea pig, and chicken) or brain regions (for example, hippocampus, striatum, etc.). Additionally, a variety of stimulations/treatments can be used (for example, excitotoxic, administration of inhibitors, etc.). In conclusion, the brain slice model described herein can be used to examine a variety of molecular mechanisms involved in excitotoxicity-mediated brain injury.

Excitotoxic Stimulation of Brain Microslices as an In vitro Model of Stroke

We have developed a brain slice model which can be used to examine molecular mechanisms involved in excitotoxicity-mediated brain injury.&#160;This technique generates viable mature brain tissue and reduces animal numbers required for experimentation, whilst keeping the neuronal circuitry, cellular interactions, and postsynaptic compartments partly intact.

Examining molecular mechanisms involved in neuropathological conditions, such as ischemic stroke, can be difficult when using whole animal systems. As ...

Clinical Assessment of Spatiotemporal Gait Parameters in Patients and Older Adults

Spatial and temporal characteristics of human walking are frequently evaluated to identify possible gait impairments, mainly in orthopedic and neurological patients1-4, but also in healthy older adults5,6. The quantitative gait analysis described in this protocol is performed with a recently-introduced photoelectric system (see&#160;Materials table) which has the potential to be used in the clinic because it is portable, easy to set up (no subject preparation is required before a test), and does not require maintenance and sensor calibration. The photoelectric system consists of series of high-density floor-based photoelectric cells with light-emitting and light-receiving diodes that are placed parallel to each other to create a corridor, and are oriented perpendicular to the line of progression7. The system simply detects interruptions in light signal, for instance due to the presence of feet within the recording area. Temporal gait parameters and 1D spatial coordinates of consecutive steps are subsequently calculated to provide common gait parameters such as step length, single limb support and walking velocity8, whose validity against a criterion instrument has recently been demonstrated7,9. The measurement procedures are very straightforward; a single patient can be tested in less than 5 min&#160;and a comprehensive report can be generated in less than 1&#160;min.

This protocol is used to evaluate spatial and temporal gait variables of neurological/orthopedic patients and older persons by means of a recently-introduced floor-based photocell system.

Spatial and temporal characteristics of human walking are frequently evaluated to identify possible gait impairments, mainly in orthopedic and ...

Reproducible Arterial Denudation Injury by Infrarenal Abdominal Aortic Clamping in a Murine Model

Percutaneous vascular interventions uniformly result in arterial denudation injuries that subsequently lead to thrombosis and restenosis. These complications can be attributed to impairments in re-endothelialization within the wound margins. Yet, the cellular and molecular mechanisms of re-endothelialization remain to be defined. While several animal models to study re-endothelialization after arterial denudation are available, few are performed in the mouse because of surgical limitations. This undermines the opportunity to exploit transgenic mouse lines and investigate the contribution of specific genes to the process of re-endothelialization. Here, we present a step-by-step protocol for creating a highly reproducible murine model of arterial denudation injury in the infrarenal abdominal aorta using external vascular clamping. Immunocytochemical staining of injured aortas for fibrinogen and &#946;-catenin demonstrate the exposure of a pro-thrombotic surface and the border of intact endothelium, respectively. The method presented here has the advantages of speed, excellent overall survival rate, and relative technical ease, creating a uniquely practical tool for imposing arterial denudation injury in transgenic mouse models. Using this method, investigators may elucidate the mechanisms of re-endothelialization under normal or pathological conditions.

Understanding the cellular and molecular mechanisms of re-endothelialization following arterial denudation injury is of paramount importance in preventing thrombosis and restenosis of arteries. Here we describe a protocol for reproducible arterial denudation injury of the infrarenal abdominal aorta. The procedure was developed to investigate the underlying mechanisms that regulate endothelial regeneration using mouse models.

Percutaneous vascular interventions uniformly result in arterial denudation injuries that subsequently lead to thrombosis and restenosis. These ...

Repeated Measurement of Respiratory Muscle Activity and Ventilation in Mouse Models of Neuromuscular Disease

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated measurements over weeks or months of accessory respiratory muscle activity while simultaneously measuring ventilation in a non-anesthetized, freely behaving mouse. The technique includes the surgical implantation of a radio transmitter and the insertion of electrode leads into the scalene and trapezius muscles to measure the electromyogram activity of these inspiratory muscles. Ventilation is measured by whole-body plethysmography, and animal movement is assessed by video and is synchronized with electromyogram activity. Measurements of muscle activity and ventilation in a mouse model of amyotrophic lateral sclerosis are presented to show how this tool can be used to investigate how respiratory muscle activity changes over time and to assess the impact of muscle activity on ventilation. The described methods can easily be adapted to measure the activity of other muscles or to assess accessory respiratory muscle activity in additional mouse models of disease or injury.

This paper introduces a method for repeated measurements of ventilation and respiratory muscle activity in a freely behaving amyotrophic lateral sclerosis (ALS) mouse model throughout disease progression with whole-body plethysmography and electromyography via an implanted telemetry device.

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated ...