Research funding was received through the National Institute of Neurological Disorders and Stroke &#160;(NS049177 and NS087834). The authors wish&#160;to acknowledge the intellectual contributions of the late Dr. Paul Bass and his colleagues to the original design of the strain gages; and Carol Tollefsrud for the fabrication and marketing of the strain gages until the cessation of production in 2010 as well as for her insightful correspondence.

All procedures followed National Institutes of Health guidelines and were approved by the Institutional Animal Care and Use Committee at the Penn State Hershey College of Medicine. Rats were housed using common vivarium practices. Note: This protocol uses male Wistar rats &#8805;8 weeks of age and initially weighing 175 - 200 g.
1. Procedures for Fabrication of Strain Gage
<ol>
	<li>Most tooling and components remain available from the original or successor companies and are summarized in Table 1.</li>
	<li>Preparation and Bonding of two single element strain gages
	<ol>
		<li>Handle the strain gage elements (EA-06-031-350) carefully with clean Dumont #5 forceps. To limit unwanted movement of elements, use a small, clean, self-adhesive piece of paper with the adhesive side facing up to secure elements to the work surface without the risk of contamination or excessive adhesion.</li>
		<li>Bond two single strain gage elements back-to-back, to form a dual element. Clean the back of each element film with Isopropyl alcohol and allow drying by evaporation (drying with gauze often introduces fiber contaminants that are difficult to remove). Under stereomicroscope guidance (1 - 3x), and using a clean artist brush (10-0 camel hair), apply a thin film of epoxy-phenolic adhesive to the back of one element and immediately place the opposing back of the second element in contact and align the foil grids&#160;(Figure 1A).</li>
		<li>Place the bonded elements in a 50 - 60 &#176;C oven O/N to fully cure the epoxy. 
		NOTE: Do not clamp the bonded elements since excess epoxy may seep onto the elements and pressure may cause misalignment of the grids. The two-part epoxy has a usable refrigerated-life of only 6 weeks after mixing. Bond and cure a sufficient supply of elements at one time and store them in a clean, dust free, environment for later use.</li>
	</ol>
	</li>
	<li>Sizing and wiring dual element strain gages
	<ol>
		<li>Trim bonded dual elements to a final size of 3 x 3 mm with a #11 scalpel or single edge razor blade. Delay trimming the topmost portion of the dual elements at this time in order to have an area for safely handling the element&#160;(Figure 1A).</li>
		<li>Each element requires a four-conductor wire fabricated from three-conductor, bondable, Teflon insulated wire (P/N 336-FTE). Disassemble one 30 cm braided strand of three-conductor wire into three constituent wires.</li>
		<li>To make a four wire cable, pair one of the resulting single wires with the like-colored wire contained within a second 30 cm length of three conductor wire. In the following steps, these matching colored wires will be joined at the terminal end to form a common wire for the final strain gage (Figure 1B).</li>
		<li>Remove approximately 1mm of Teflon insulation from both ends of each wire with thermal wire strippers. Using activated rosin soldering flux and low temperature solder (melting point 183 &#186;C) tin the wire ends with a soldering pencil.</li>
		<li>For the next stage, a microsoldering tip is needed to form a more discrete solder joint to prevent heat damage to the film layer of the element (Figure 1C). To fabricate a smaller microsoldering tip, wrap a small piece of copper wire (~0.25 mm diameter) once around the standard soldering tip, ensuring that the copper wire extends beyond the length of the standard soldering tip.</li>
		<li>Flux just the solder pads on one side of the bonded dual element with a clean 10-0 brush and solder one single lead and one of the paired common leads to the solder pad (Figure 1D). Residual flux can be removed afterward with a clean brush dipped in resin solvent.</li>
		<li>Repeat the process on the opposite side, ensuring that the remaining common wire lead is soldered to the pad opposite the original common lead.</li>
	</ol>
	</li>
	<li>Testing and epoxying dual element strain gages
	<ol>
		<li>Solder gold socket connectors (E363/0) to the free ends of the wire leads. At this point, connect the strain gage to a recording amplifier (described below) to test the integrity of the dual element assembly.</li>
		<li>Measure the resistance with a good quality volt-ohm meter. Elements register a resistance of approximately 350 &#937;. Resolder inadequate connections at this point with fresh solder.</li>
		<li>If the solder connections and the dual element assembly are deemed satisfactory, trim off any remaining element film.</li>
		<li>Insulate the solder joints on the element solder pads with a thin layer of two-part silicone-rubber epoxy resin (P/N E211). For best results, partly cure the resin for 20 - 30 min prior to application (Figure 1E).</li>
	</ol>
	</li>
	<li>Encapsulating dual element strain gages in silicone
	<ol>
		<li>Cut three pieces of 0.5 mm thick silicone sheet (P/N 20-20) to 15 mm 2 and clean the silicone with distilled water. Cut one piece of silicone sheet into a U-shape in order to accommodate the final dual element assembly without deforming the encapsulating silicone (Figure 1F).</li>
		<li>Coat the inner surfaces of the notch-free silicone sheets with clear silicone adhesive.</li>
		<li>Sandwich the dual element assembly within the notch and the aligned outer sheets, then gently press out any excess silicone, as well as air bubbles, from the center outward. Carefully clamp the encapsulated assembly between two blocks of metal bar stock for 24 hr to ensure uniform thickness and that no deformations occur.</li>
		<li>Allow the excess silicone to remain along the boundaries of the assembly and cure. This excess will be removed when the sheet silicone is trimmed to the desired final dimensions (commonly 6 mm x 8 mm; Figure 1G).</li>
	</ol>
	</li>
	<li>Completion of wire connector and calibration
	<ol>
		<li>Reinforce the solder joint of the gold socket connectors on the individual terminal wire leads with 3 mm (1/8 inch) shrink tubing and align within a plastic electrode pedestal (MS363, Figure 1H). Secure the electrode pedestal and wires with 0.125 and 0.25 inch diameter shrink tubing to prevent disconnection during the experiment (Figure 1I).</li>
		<li>Strain gage signals are processed through a high gain bridge amplifier (P/N AMP-01-SG). Connect the strain gage to the amplifier using a cable with a mated plug (363 - SL/6) to match the electrode pedestal. The threaded cap provides additional security to maintain uninterrupted signals during the experiment.</li>
		<li>Adjust the Bridge, Balance and Gain settings on the amplifier to a dedicated strain gage per manufacturer instructions. Affix the end of the strain gage where the wires exit horizontally to a rigid clamp and calibrate by placing a 1 g static load on the opposite end as originally described by Pascaud and colleagues 2, 16, 17.</li>
	</ol>
	</li>
</ol>
2. Surgical Procedures for Acute Implantation of Strain Gage
<ol>
	<li>Animal Care and Preparation:
	<ol>
		<li>Food deprive experimental animals the night before surgical implantation (water may be provided ad libitum).</li>
		<li>Deeply anesthetize the animal. Thiobutabarbital (100 &#8211; 150 mg/kg; i.p. for rats) is preferred for terminal (i.e., non-survival) strain gage implantation and experimentation due to sustained anesthetic effect and minimal alteration of gastric reflexes in the rat 10. Test for absence of paw pinch reflex to determine depth of anesthesia.</li>
		<li>Prepare the rat for aseptic surgery as dictated by the experimental design and approved IACUC guidelines including sterilizing surgical tools, shaving incision sites, applying vet eye ointment and disinfecting all surgical areas.&#160;NOTE:&#160;If the strain gage is being adapted for chronic use, sterilize the the strain gage only with ethylene oxide (gas) sterilization. The use of heat or chemical sterilization techniques may damage the strain gage.</li>
	</ol>
	</li>
	<li>Tracheal intubation for terminal experiments:
	<ol>
		<li>For long duration, terminal, experiments intubate the rat with a tracheal tube to maintain an open airway. Make a 1 - 2 cm midline incision on the ventral side of the neck from the inferior border of the mandible to the sternal notch.</li>
		<li>Separate the underlying strap muscles using blunt dissection at the midline to expose the trachea. Isolate the trachea from the underlying esophagus and place a loop of 3-0 ethilon suture between the trachea and esophagus to form a ligature.</li>
		<li>Open the trachea anteriorly by making a small incison&#160;in the membrane between two of the cartilaginous rings of the trachea just distal to the thyroid gland. Insert a small piece of polyethylene tubing (P/N PE-270), 5mm in length (and beveled at one end) into the trachea and secure it into place with the ligature.</li>
		<li>Put the strap muscles back in place and suture the overlying skin with 3-0 ethilon.</li>
	</ol>
	</li>
	<li>Strain gage instrumentation to gastrointestinal surface:
	<ol>
		<li>Thread the four corners of the strain gage with 4 - 5 cm lengths of 4-0, or smaller, sterile silk suture using a #14 taper point 3/8 circle needle prior to surgery. Silk suture provides a high level of flexibility and is less likely to damage the silicone encapsulating the strain gage element. 
		NOTE: silk thread is acceptable for non-survival surgeries and for internal applications, where the wicking of bacteria across an epithelial barrier is not a risk. In applications requiring survival surgery, a prolene suture is necessary in order to reduce the risk of infection inherent in the braided cloth fibers of the silk suture.</li>
		<li>Perform a laparotomy by incising the abdominal skin along the midline. Section the rectus abdominus musculature along the connecting linea alba (avascular) to prevent bleeding. Then make a very superficial midline incision in the parietal peritoneum to avoid lacerating underlying abdominal viscera.</li>
		<li>Exteriorize the stomach with the aid of saline soaked cotton tipped applicators. Keep the stomach in position by carefully placing it on a saline soaked gauze pad at the caudal end of the abdominal incision.</li>
		<li>Align the grid of the encapsulated strain gage in parallel with the circular smooth muscle fibers. Using the previously threaded sutures (step 2.3.1), attach the corners of the gage to the ventral serosal surface of the gastric corpus using a #14 taper point 3/8 circle needle. In order to minimize tissue damage and potential bleeding, do not use cutting-edged needles and do not perforate any superficial blood vessels on the surface of the stomach.</li>
		<li>Begin the suture pattern of the gage along the greater curvature of the stomach near the fundus/corpus boundary and proceed next along the fundus/corpus boundary toward the lesser curvature. The serosa underlying the strain gage should neither be slack nor overly stretched in order to obtain the best results.</li>
		<li>Carefully return the stomach to its anatomical position using saline-soaked cotton tipped applicators.</li>
		<li>In an acute model, exteriorize the strain gage leads at the caudal end of the midline incision before closure of the abdominal incision. Secure the free wires to the animal (e.g., hind foot) in order to provide strain relief during manipulation of the animal or terminal wire connector. Close the rectus abdominus muscles and the abdominal skin separately with 3-0 nylon suture. In a chronic model, secure the leads subcutaneously along the dorsal side of the rat and exteriorize them above the skull 18.</li>
		<li>After surgical instrumentation, place animals in a stereotaxic frame to support the head and elevate the upper torso. The latter step helps to reduce respiration artifact during recording. Monitor rectal temperature and maintain at 37&#177;1 &#176;C using a feedback-controlled heating pad.</li>
		<li>At the conclusion of terminal experiments utilizing thiobutabarbital anesthesia, the animal must be euthanized in a manner consistent with American Veterinary Medical Association (AVMA) Guidelines on Euthanasia.</li>
	</ol>
	</li>
	<li>Gastric Motility recordings:
	<ol>
		<li>Amplify the strain gage signal with any commercially available DC bridge amplifier.</li>
		<li>Record the DC output signals on a computer using the chart recorder function of any commercially available data acquisition system. 
		NOTE: A hardcopy of the amplifier output can be generated through a polygraph chart recorder.</li>
	</ol>
	</li>
</ol>
3. Representative Measurement of Gastric Contractions Following Brainstem Stimulation
<ol>
	<li>Exposure of brainstem and fourth ventricle
	<ol>
		<li>After surgical instrumentation, and placement of the Thiobutabarbital-anesthetized animal in a stereotaxic frame, make a 1.5 - 2 cm midline skin incision from the occipital bone toward the base of the neck.</li>
		<li>Separate the connective tissue joining the bilateral muscle bellies of the underlying neck muscles along the midline (muscles from superficial to deep are levator auris&#160;longus cranial portion, levator auris&#160;longus caudal portion, and platysma cranial portion).</li>
		<li>Detach the levator auris&#160;longus from the occipital bone once the midline is clearly defined and exposed.</li>
		<li>Carefully expose the caudal region of the skull by using blunt dissection to detach platysma muscle from the underlying dura mater.</li>
		<li>Use a new 25 G&#160;needle to carefully detach the dura mater along the foramen magnum extending bilaterally to the occipital condyles.</li>
		<li>Use #5 Dumont forceps to remove the pia and arachnoid meninges overlying the fourth ventricle and expose the brainstem.</li>
	</ol>
	</li>
	<li>Administering fourth ventricle thyrotropin releasing hormone or intravenous sodium nitroprusside
	<ol>
		<li>Weigh and dissolve thyrotropin releasing hormone (TRH) in sterile saline to reach a final concentration of 50 &#181;M TRH.</li>
		<li>Weigh and dissolve sodium nitroprusside (SNP) in sterile saline to reach a final concentration of 150 &#181;M SNP.</li>
		<li>Using a 10 &#181;l syringe, administer 2 &#181;l of TRH (final dose equals 100 pmol) to the dorsal surface of the brainstem fourth ventricle to facilitate recording of gastric contractions.</li>
		<li>Using a sterile syringe and 27 G&#160;needle, administer 150 &#181;mol/kg of SNP through the tail vein to facilitate recording of gastric relaxation.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests. The suppliers listed in this manuscript are provided for reference only.

The procedures presented here allow individual laboratories to fabricate sensitive miniature strain gages for biological applications including, but not limited to, gastrointestinal motility in small laboratory animals. Since the commercial manufacture of these strain gages has ceased, laboratories investigating gastrointestinal function are limited to other techniques which may not permit the full range of experimental applications that are available. This report provides an updated and more detailed description of prev...

Experimental studies that record in vivo gastrointestinal (GI) motility across a number of experimental conditions remain a powerful tool for understanding the underlying normal and pathophysiological processes necessary for nutrient homeostasis. Traditionally, numerous experimental methodologies, some with similarities to those found in clinical practice 1, have been employed to directly quantify changes in GI contraction rate 2-5, intraluminal pressure 6, 7, or the GI transit of non-absorbable markers 8, 9 or stable isotopes 10-12. Each of these techniques has unique advantages and disadvantages, which have been addressed previously in the literature. For example, the utility of balloon manometry to quantify pressure changes has been questioned due to the inherent compliance of the balloon material while gastrointestinal recovery of nonabsorbable markers requires euthanizing the experimental animal for a single data point. Recently, the application and validation of a miniaturized arterial pressure catheter has been reported that offers a non-surgical method for monitoring gastric contractility in rats and mice 3. While an orogastrically placed pressure transducer effectively eliminates confounding variables on gastrointestinal function by avoiding invasive surgical procedures, such an approach is only suitable for anesthetized preparations. Furthermore, the lack of visual guidance does not permit consistent placement of the transducer within specific regions of the stomach. As such, this application is restricted to the stomach or colon since visualization, coupled with the relatively stiff transducer wire, within the duodenum or ileum is not an option.
Similarly, the biomagnetic alternate current biosusceptometry (ACB) technique has been validated for GI contraction analysis 4. While the ACB technique provides a noninvasive approach for measuring gastrointestinal contractions, ACB suffers from a similar limitation in that the use of ingested magnetic detection media does not permit precise recording of specific regions of the GI tract. This limitation can be overcome through the surgical implantation of magnetic markers. Nonetheless, the ACB technique necessitates that the animal be anesthetized for data collection.
Ultrasonomicrometry has been employed in some GI studies 13, 14 in order to take advantage of the small size, spatial, and temporal advantages of piezoelectric crystal transmitter/receivers. Waves of gastric smooth muscle contraction are not a high-frequency event and occur at a rate of approximately 3 - 5 cycles/min. Therefore, the temporal advantages of sonomicrometry may be unnecessary to justify the cost. Furthermore, while linear motion is accurately measured with sonomicrometry, limitations have been presented regarding accurate gastrointestinal data interpretation that may result from implanting an insufficient number of crystals 14.
Based upon the original designs of Bass and colleagues 2, 15 this visualized protocol more fully documents the step-by-step fabrication and experimental application of miniature, dual element strain gages that possess high sensitivity and flexibility for recording smooth muscle contractions along the entire GI tract. The dimensions of the strain gage elements are suitable for any rodent application since sensitivity and size of the finished strain gage are most dependent upon the silicone sheets encapsulating the elements. These strain gages are readily adapted for acute and chronic application in anesthetized and freely behaving laboratory animal models thereby providing a single technique for quantifying smooth muscle contractions.

<table><tbody><tr><td>Strain gage element</td><td>Micro-Measurements (Vishay Product Group)</td><td>EA-06-031-350</td><td>&amp;nbsp;Linear pattern, foil, stress analysis strain gage (2 required)&lt;br/&gt; &lt;a href=&quot;http://www.vishaypg.com/micro-measurements/&quot;&gt;www.vishaypg.com/micro-measurements/&lt;/a&gt;&lt;br/&gt; or&lt;br/&gt; &lt;a href=&quot;http://www.vishaypg.com/docs/11070/031ce.pdf&quot;&gt;http://www.vishaypg.com/docs/11070/031ce.pdf&lt;/a&gt;</td></tr><tr><td>Epoxy-phenolic adhesive</td><td>M-bond 610</td><td>General purpose adhesive for bonding strain gage elements</td><td>&lt;a href=&quot;http://www.vishaypg.com/docs/11024/wirecable.pdf&quot;&gt;http://www.vishaypg.com/docs/11024/wirecable.pdf&lt;/a&gt;</td></tr><tr><td>3 conductor insulated wire</td><td>336-FTE</td><td>Fine gage, flexible general purpose wire</td><td>&lt;a href=&quot;http://www.vishaypg.com/docs/11024/wirecable.pdf&quot;&gt;http://www.vishaypg.com/docs/11024/wirecable.pdf&lt;/a&gt;</td></tr><tr><td>Flux and rosin solvent kit</td><td>FAR-2 M-Flux AR kit</td><td>Liquid solder flux</td><td>&lt;a href=&quot;http://www.vishaypg.com/docs/11023/soldacce.pdf&quot;&gt;http://www.vishaypg.com/docs/11023/soldacce.pdf&lt;/a&gt;</td></tr><tr><td>Solder</td><td>361A-20R-25</td><td>Optimized and recommended for strain gage applications</td><td>&lt;a href=&quot;http://www.vishaypg.com/docs/11023/soldacce.pdf&quot;&gt;http://www.vishaypg.com/docs/11023/soldacce.pdf&lt;/a&gt;</td></tr><tr><td>Gold socket connector</td><td>PlasticsOne</td><td>E363/0</td><td>Socket contact for electrode pedestal&lt;br/&gt; &lt;a href=&quot;http://www.plastics1.com/PCR/Catalog/Item.php?item=407&quot;&gt;http://www.plastics1.com/PCR/Catalog/Item.php?item=407&lt;/a&gt;</td></tr><tr><td>Electrode pedestal</td><td>MS363</td><td>Secure platform for wire contacts</td><td>&lt;a href=&quot;http://www.plastics1.com/PCR/Catalog/Item.php?item=499&quot;&gt;http://www.plastics1.com/PCR/Catalog/Item.php?item=499&lt;/a&gt;</td></tr><tr><td>6-wire cable</td><td>363 PLUG W/VINYL SL/6</td><td>Pre-fabricated vinyl-coated cable (in customized lengths) with plug adaptor to match electrode pedestal and tinned solder lugs on terminal end</td><td></td></tr><tr><td>Silicone rubber casting compound</td><td>EIS electrical products</td><td>Elan Tron E211</td><td>Potting medium for gage/wire solder joints&lt;br/&gt; &lt;a href=&quot;http://www.eis-inc.com/&quot;&gt;http://www.eis-inc.com&lt;/a&gt;</td></tr><tr><td>HOTweezers</td><td>Meisei Corporation</td><td>Model 4B</td><td>Wire insulation strippers&lt;br/&gt; &lt;a href=&quot;http://www.impexron.us/&quot;&gt;http://www.impexron.us&lt;/a&gt;</td></tr><tr><td>Soldering station</td><td>Weller (Apex Tool Group)</td><td>WES 51</td><td>High quality soldering equipment&lt;br/&gt; &lt;a href=&quot;http://www.apexhandtools.com/weller/index.cfm&quot;&gt;http://www.apexhandtools.com/weller/index.cfm&lt;/a&gt;</td></tr><tr><td>Available through http://www.eis-inc.com or http://www.amazon.com</td><td></td><td></td><td></td></tr><tr><td>Silicone sheet</td><td>Trelleborg Sealing Solutions Northborough-Life Sciences</td><td>Pharmelast 20-20</td><td>Encapsulating strain gage elements&lt;br/&gt; 10 B Forbes Road Northborough, MA 01532 (800) 634-2000</td></tr><tr><td>Amplifier</td><td>Experimetria Ltd</td><td>AMP-01-SG</td><td>&lt;br/&gt; &lt;a href=&quot;http://experimetria.com/Biological_amplifiers.php&quot;&gt;http://experimetria.com/Biological_amplifiers.php&lt;/a&gt;</td></tr></tbody></table>

fabrication and implantation of miniature dual-element strain gages for measuring in vivo gastrointestinal contractions in rodents.

Gastrointestinal dysfunction remains a major cause of morbidity and mortality. Indeed, gastrointestinal (GI) motility in health and disease remains an area of productive research with over 1,400 published animal studies in just the last 5 years. Numerous techniques have been developed for quantifying smooth muscle activity of the stomach, small intestine, and colon. In vitro and ex vivo techniques offer powerful tools for mechanistic studies of GI function, but outside the context of the integrated systems inherent to an intact organism. Typically, measuring in vivo smooth muscle contractions of the stomach has involved an anesthetized preparation coupled with the introduction of a surgically placed pressure sensor, a static pressure load such as a mildly inflated balloon or by distending the stomach with fluid under barostatically-controlled feedback. Yet many of these approaches present unique disadvantages regarding both the interpretation of results as well as applicability for in vivo use in conscious experimental animal models. The use of dual element strain gages that have been affixed to the serosal surface of the GI tract has offered numerous experimental advantages, which may continue to outweigh the disadvantages. Since these gages are not commercially available, this video presentation provides a detailed, step-by-step guide to the fabrication of the current design of these gages. The strain gage described in this protocol is a design for recording gastric motility in rats. This design has been modified for recording smooth muscle activity along the entire GI tract and requires only subtle variation in the overall fabrication. Representative data from the entire GI tract are included as well as discussion of analysis methods, data interpretation and presentation.

Representative data from a Thiobutabarbital-anesthetized rat are shown in Figure 2. The top trace represents the gastric corpus contractions from the rat during the brainstem administration of thyrotropin releasing hormone (TRH, 100 pmol), a known motility-enhancing peptide 3, 19. It shows baseline contractions prior to the increase in phasic gastric smooth muscle activity. Note: Analysis of these peaks in gastric contractions follow the original formula devised by Ormsby and Bass 20</sup...

Watch this Scientific Journal Video about Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents. at JoVE.com

Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents.

The in vivo measurement of smooth muscle contractions along the gastrointestinal tract of laboratory animals remains a powerful, though underutilized, technique. Flexible, dual element strain gages are not commercially available and require fabrication. This protocol describes the construction of reliable, inexpensive strain gages for acute or chronic implantation in rodents.

Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents.

Gastrointestinal dysfunction remains a major cause of morbidity and mortality. Indeed, gastrointestinal (GI) motility in health and disease remains an ...

fabrication-implantation-miniature-dual-element-strain-gages-for

Research

JoVE Journal

Bioengineering

10.3K Views.  Penn State University College of Medicine. The in vivo measurement of smooth muscle contractions along the gastrointestinal tract of laboratory animals remains a powerful, though underutilized, technique. Flexible, dual element strain gages are not commercially available and require fabrication. This protocol describes the construction of reliable, inexpensive strain gages for acute or chronic implantation in rodents.

Video: Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents.

An in vivo Rodent Model of Contraction-induced Injury and Non-invasive Monitoring of Recovery

Muscle strains are one of the most common complaints treated by physicians. A muscle injury is typically diagnosed from the patient history and physical exam alone, however the clinical presentation can vary greatly depending on the extent of injury, the patient's pain tolerance, etc. In patients with muscle injury or muscle disease, assessment of muscle damage is typically limited to clinical signs, such as tenderness, strength, range of motion, and more recently, imaging studies. Biological markers, such as serum creatine kinase levels, are typically elevated with muscle injury, but their levels do not always correlate with the loss of force production. This is even true of histological findings from animals, which provide a "direct measure" of damage, but do not account for all the loss of function. Some have argued that the most comprehensive measure of the overall health of the muscle in contractile force. Because muscle injury is a random event that occurs under a variety of biomechanical conditions, it is difficult to study. Here, we describe an in vivo animal model to measure torque and to produce a reliable muscle injury. We also describe our model for measurement of force from an isolated muscle in situ. Furthermore, we describe our small animal MRI procedure.

An in vivo Rodent Model of Contraction-induced Injury and Non-invasive Monitoring of Recovery

An in vivo animal model of injury is described. The method takes advantage of the subcutaneous position of the fibular nerve. Velocity, timing of muscle activation, and arc of motion are all pre-determined and synchronized using commercial software. Post injury changes are monitored in vivo using MR imaging/spectroscopy.

Muscle strains are one of the most  common complaints treated by physicians. A muscle injury is typically diagnosed from the patient history and  physical ...

Medicine

Physiological Experimentation with the Crayfish Hindgut: A Student Laboratory Exercise

The purpose of the report is to describe dissection techniques for preparing the crayfish hindgut and to demonstrate how to make physiological recordings with a force transducer to monitor the strength of contraction. In addition, we demonstrate how to visually monitor peristaltic activity, which can be used as a bioassay for various peptides, biogenic amines and neurotransmitters. This preparation is amenable to student laboratories in physiology and for demonstrating pharmacological concepts to students. This preparation has been in use for over 100 years, and it still offers much as a model for investigating the generation and regulation of peristaltic rhythms and for describing the mechanisms underlying their modulation. The pharmacological assays and receptor sub-typing that were started over 50 years ago on the hindgut still contribute to research today. This robust preparation is well suited to training students in physiology and pharmacology.

In this report we demonstrate techniques that can be used to investigate the biology of the crayfish hindgut. We show how to dissect a crayfish abdomen and study the associated anatomy, physiology and modulation of activity. The peristaltic activity and strength of contractions are measured using a force transducer.

The purpose of the report is to describe dissection techniques for preparing the crayfish hindgut and to demonstrate how to make physiological recordings ...

Neuroscience

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. Despite the discovery of key regulators of uterine contractility, this process is still not fully understood. Transgenic mice provide an ideal model in which to study parturition. Previously, the only method to study uterine contractility in the mouse was ex vivo isometric tension recordings, which are suboptimal for several reasons. The uterus must be removed from its physiological environment, a limited time course of investigation is possible, and the mice must be sacrificed. The recent development of radiometric telemetry has allowed for longitudinal, real-time measurements of in vivo intrauterine pressure in mice. Here, the implantation of an intrauterine telemeter to measure pressure changes in the mouse uterus from mid-pregnancy until delivery is described. By comparing differences in pressures between wild type and transgenic mice, the physiological impact of a gene of interest can be elucidated. This technique should expedite the development of therapeutics used to treat myometrial disorders during pregnancy, including preterm labor.

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

This manuscript provides a protocol for implanting telemeters in the mouse for the purpose of measuring intrauterine pressures during pregnancy.

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. ...

Wireless Intramuscular EMG System for Locomotion Studies in Stroke Model Rats

A Real-Time Wearable Electromyography Measurement System for Small Animals

The intramuscular electromyography (EMG) measurement method for experimental animals has been implemented in various ways. Among these methods, tethering cables to external measurement devices can restrict the movement of experimental animals, while implantable devices may cause unwanted side effects due to the constant presence of a device with considerable size and weight. To address these issues, we propose a low-cost, wireless, detachable EMG measurement system and experimental procedure. This article focuses on the surgical installation of intramuscular wire electrodes with small connectors and the development of the wireless system. Notably, in this system, only the wire electrodes are inserted into the animal's body. Using this system, EMG measurements can be easily performed by attaching the circuit system to a connector installed on the animal's back, with real-time monitoring achievable on a laptop. The proposed method is explained in a detailed, step-by-step manner, followed by a demonstration involving the insertion of intramuscular electrodes into the hindlimbs of a rat. A treadmill experiment is conducted for a locomotion study, and the resulting electrophysiological signals are subsequently obtained and analyzed.

A protocol is presented for the installation and preparation of intramuscular electrodes, along with an animal-mountable, miniaturized measurement device for electromyography (EMG) analysis in locomotion studies. This system enables the wireless transmission of real-time EMG data for investigating locomotion recovery in stroke model rats.

The intramuscular electromyography (EMG) measurement method for experimental animals has been implemented in various ways. Among these methods, tethering ...

Engineering

Gastrointestinal Motility Monitor (GIMM)

The Gastrointestinal Motility Monitor (GIMM; Catamount Research and Development; St. Albans, VT) is an in vitro system that monitors propulsive motility in isolated segments of guinea pig distal colon. The complete system consists of a computer, video camera, illuminated organ bath, peristaltic and heated water bath circulating pumps, and custom GIMM software to record and analyze data. Compared with traditional methods of monitoring colonic peristalsis, the GIMM system allows for continuous, quantitative evaluation of motility. The guinea pig distal colon is bathed in warmed, oxygenated Krebs solution, and fecal pellets inserted in the oral end are propelled along the segment of colon at a rate of about 2 mm/sec. Movies of the fecal pellet proceeding along the segment are captured, and the GIMM software can be used track the progress of the fecal pellet. Rates of propulsive motility can be obtained for the entire segment or for any particular region of interest. In addition to analysis of bolus-induced motility patterns, spatiotemporal maps can be constructed from captured video segments to assess spontaneous motor activity patterns. Applications of this system include pharmacological evaluation of the effects of receptor agonists and antagonists on propulsive motility, as well as assessment of changes that result from pathophysiological conditions, such as inflammation or stress. The guinea pig distal colon propulsive motility assay, using the GIMM system, is straightforward and simple to learn, and it provides a reliable and reproducible method of assessing propulsive motility.

Gastrointestinal Motility Monitor GIMM

Evaluation of colonic motility in the guinea pig distal colon with the Gastrointestinal Motility Monitor (GIMM) is a straightforward and simple to learn approach to quantitatively evaluate propulsive motility in the gastrointestinal tract.

The Gastrointestinal Motility Monitor (GIMM; Catamount Research and Development; St. Albans, VT) is an in vitro system that monitors propulsive motility ...

Biology

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

One central goal of mechanobiology is to understand the reciprocal effect of mechanical stress on proteins and cells. Despite its importance, the influence of mechanical stress on cellular function is still poorly understood. In part, this knowledge gap exists because few tools enable simultaneous deformation of tissue and cells, imaging of cellular activity in live animals, and efficient restriction of motility in otherwise highly mobile model organisms, such as the nematode Caenorhabditis elegans. The small size of C. elegans makes them an excellent match to microfluidics-based research devices, and solutions for immobilization have been presented using microfluidic devices. Although these devices allow for high-resolution imaging, the animal is fully encased in polydimethylsiloxane (PDMS) and glass, limiting physical access for delivery of mechanical force or electrophysiological recordings. Recently, we created a device that integrates pneumatic actuators with a trapping design that is compatible with high-resolution fluorescence microscopy. The actuation channel is separated from the worm-trapping channel by a thin PDMS diaphragm. This diaphragm is deflected into the side of a worm by applying pressure from an external source. The device can target individual mechanosensitive neurons. The activation of these neurons is imaged at high-resolution with genetically-encoded calcium indicators. This article presents the general method using C. elegans strains expressing calcium-sensitive activity indicator (GCaMP6s) in their touch receptor neurons (TRNs). The method, however, is not limited to TRNs nor to calcium sensors as a probe, but can be expanded to other mechanically-sensitive cells or sensors.

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

New tools for mechanobiology research are needed to understand how mechanical stress activates biochemical pathways and elicits biological responses. Here, we showcase a new method for selective mechanical stimulation of immobilized animals with a microfluidic trap allowing high-resolution imaging of cellular responses.

One central goal of mechanobiology is to understand the reciprocal effect of mechanical stress on proteins and cells. Despite its importance, the ...

Studying Murine Small Bowel Mechanosensing of Luminal Particulates

Gastrointestinal (GI) motility is critical for normal digestion and absorption. In the small bowel, which absorbs nutrients, motility optimizes digestion and absorption. For this reason, some of the motility patterns in the small bowel include segmentation for mixing of luminal contents and peristalsis for their propulsion. Physical properties of luminal contents modulate the patterns of small bowel motility. The mechanical stimulation of GI mechanosensory circuits by transiting luminal contents and underlying gut motility initiate and modulate complex GI motor patterns. Yet, the mechanosensory mechanisms that drive this process remain poorly understood. This is primarily due to a lack of tools to dissect how the small bowel handles materials of different physical properties. To study how the small bowel handles particulates of varying sizes, we have modified an established in vivo method to determine small bowel transit. We gavage live mice with fluorescent liquid or tiny fluorescent beads. After 30 minutes, we dissect out the bowels to image the distribution of fluorescent contents across the entirety of the GI tract. In addition to high-resolution measurements of the geometric center, we use variable size binning and spectral analysis to determine how different materials affect small bowel transit. We have explored how a recently discovered "gut touch" mechanism affects small bowel motility using this approach.

To study how the small bowel handles particulates of varying sizes, we have modified an established in vivo method to determine small bowel transit.

Gastrointestinal (GI) motility is critical for normal digestion and absorption. In the small bowel, which absorbs nutrients, motility optimizes digestion ...

Studying Gut-Brain Interaction via Transcranial Calcium Imaging in Mice

Real-time Analysis of Gut-brain Neural Communication: Cortex wide Calcium Dynamics in Response to Intestinal Glucose Stimulation

Communication between the gastrointestinal tract and the brain after nutrient absorption plays an essential role in food preference, metabolism, and feeding behaviors. Particularly concerning specific nutrients, many studies have elucidated that the assimilation of glucose within gut epithelial cells instigates the activation of many signaling molecules. Hormones such as glucagon-like peptide-1 are renowned as quintessential signaling mediators. Since hormones predominantly influence the brain through circulatory pathways, they slowly modulate brain activity.
However, recent studies have shown two expeditious gut-brain pathways facilitated by the autonomic nervous system. One operates via the spinal afferent neural pathway, while the vagus nerve mediates the other. Consequently, brain responses following glucose assimilation in the gastrointestinal tract are complicated. Moreover, as intestinal stimulation finally induces diverse cortical activities, including sensory, nociceptive, reward, and motor responses, it is necessary to employ methodologies that facilitate the visualization of localized brain circuits and pan-cortical activities to comprehend gut-brain neural transmission fully. Some studies have indicated precipitous alterations in calcium ion (Ca2+) concentrations within the hypothalamus and ventral tegmental area independently through different pathways after intestinal stimulation. However, whether there are changes in cerebral cortex activity has not been known.
To observe cerebral cortex activity after intragastric glucose injection, we developed an imaging technique for real-time visualization of cortex wide Ca2+ dynamics through a fully intact skull, using transgenic mice expressing genetically encoded Ca2+ indicators. This study presents a comprehensive protocol for a technique designed to monitor intestinal stimulation-induced transcranial cortex wide Ca2+ imaging following intragastric glucose injection via an implanted catheter. The preliminary data suggest that administering glucose solution into the gut activates the frontal cortex, which remains unresponsive to water administration.

Gut-brain communication, facilitated by the vagus nerve, is crucial for communication between the gastrointestinal endocrine system and the brain. However, whether intragastric glucose injection can change cortical activity is still not understood. Here, we offer a comprehensive protocol to observe changes in cortical activity after glucose injection into the duodenum.

Communication between the gastrointestinal tract and the brain after nutrient absorption plays an essential role in food preference, metabolism, and ...

Advanced Imaging of Dorsal Root Ganglion Neurons for Sensory Modality Research

In Vivo Calcium Imaging of Dorsal Root Ganglia Neurons' Response to Somatic and Visceral Stimuli

A technique is described for surgically exposing the dorsal root ganglion (DRG) of the lumbar-6 in a live, anesthetized laboratory mouse, along with the protocol for in vivo calcium imaging of the exposed DRG in response to various visceral and somatic stimuli. Pirt-GCaMP6s mice or C57BL6 mice intrathecally injected with AAV viruses packaged with GCaMP6s were utilized to capture Ca2+ transients. The amplitude of these transients indicates sensitivity to specific sensory modalities. Afferent fibers originate from internal organs, with primary neuronal cell bodies in spinal or vagal ganglia. Studies on visceral nociception and acupuncture analgesia can potentially be conducted on primary sensory neurons using advanced imaging technologies like in vivo calcium imaging, allowing for the recording of neuronal activity ensembles in the intact animal during stimulation or intervention. The responses of DRG neuron ensembles to somatic and visceral stimuli applied to their corresponding receptive fields were recorded. This technique illustrates how neuronal populations react to various types of somatic and visceral stimuli. It is possible to comprehensively compare neuronal ensemble responses to different stimuli, which is a particularly valuable approach in research on visceral pain and segmental mechanisms of somatic stimulation, such as acupuncture.

The present protocol outlines in vivo calcium imaging for measuring the responses of ensembles of lumbar-6 DRG neurons to somatic and visceral stimuli. Thorough comparisons can be made among neurons responding to different stimuli. This protocol is valuable for investigating mechanisms of visceral pain and somatic stimulation, such as acupuncture.

A technique is described for surgically exposing the dorsal root ganglion (DRG) of the lumbar-6 in a live, anesthetized laboratory mouse, along with the ...

Fabrication and Surgical Implantation of Electromyographic Electrodes in Mice

Recording Forelimb Muscle Activity in Head&#45;Fixed Mice with Chronically Implanted EMG Electrodes

Powerful genetic and molecular tools available in mouse systems neuroscience research have enabled researchers to interrogate motor system function with unprecedented precision in head-fixed mice performing a variety of tasks. The small size of the mouse makes the measurement of motor output difficult, as the traditional method of electromyographic (EMG) recording of muscle activity was designed for larger animals like cats and primates. Pending commercially available EMG electrodes for mice, the current gold-standard method for recording muscle activity in mice is to make electrode sets in-house. This article describes a refinement of established procedures for hand fabrication of an electrode set, implantation of electrodes in the same surgery as headplate implantation, fixation of a connector on the headplate, and post-operative recovery care. Following recovery, millisecond-resolution EMG recordings can be obtained during head-fixed behavior for several weeks without noticeable changes in signal quality. These recordings enable precise measurement of forelimb muscle activity alongside in vivo neural recording and/or perturbation to probe mechanisms of motor control in mice.

Author Spotlight: Investigating Mouse Motor Cortex Interactions from Muscle Activity to Neural Dynamics

This protocol describes the hand fabrication and surgical implantation of electromyographic (EMG) electrodes in the forelimb muscles of mice to record muscle activity during head-fixed behavior experiments.

Powerful genetic and molecular tools available in mouse systems neuroscience research have enabled researchers to interrogate motor system function with ...

Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents.

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Przeglądaj więcej filmów

An in vivo Rodent Model of Contraction-induced Injury and Non-invasive Monitoring of Recovery

Physiological Experimentation with the Crayfish Hindgut: A Student Laboratory Exercise

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

A Real-Time Wearable Electromyography Measurement System for Small Animals

Gastrointestinal Motility Monitor (GIMM)

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

Studying Murine Small Bowel Mechanosensing of Luminal Particulates

Real-time Analysis of Gut-brain Neural Communication: Cortex wide Calcium Dynamics in Response to Intestinal Glucose Stimulation

In Vivo Calcium Imaging of Dorsal Root Ganglia Neurons' Response to Somatic and Visceral Stimuli

Recording Forelimb Muscle Activity in Head-Fixed Mice with Chronically Implanted EMG Electrodes