Name: measures of heart and ventilatory rates in freely moving crayfish
Uploaded: 2009-10-15T23:30:00
Duration: 6 min 47 s
Description: Watch this Scientific Journal Video about Measures of Heart and Ventilatory Rates in Freely Moving Crayfish at JoVE.com

Introduction
Organisms must exhibit a wide range of behaviors to respond to threatening stimuli and changes in the environment. The autonomic response is well studied in vertebrates as the fight or flight response in which the sympathetic nervous system physiologically readies the internal environment (Carpenter, 1976; Nicholls et al., 2001). It is through physiological changes that an organism can regulate the stress response and alter behavior to react to an impending attack or environmental change. Evidence shows that the cardio-vascular system is tightly connected with the respiratory system thus enabling the internal environment to respond and implement a response. It is well known that the autonomic control of the respiratory and cardiovascular systems can regulate oxygen availability and nutrients to specific target tissues needed for an impending behavioral response.
In vertebrates, the autonomic and physiological response is so well evolved; it would seem likely that complex invertebrates would similarly possess a developed response system (Schapker et al., 2002; Zavarzin, 1941). Even though vertebrates and invertebrates are very different systematically; highly developed invertebrates require the rapid cardiovascular and respiratory response to respond in a fight or flight manner. For many invertebrates, a sympathetic-like physiological response can be quantified. Dating back to 1927, many of the very early studies have been conducted in invertebrates, specifically using arthropods in general (Alexandrowicz, 1932; Orlov, 1927; Zavarzin, 1941).
Crayfish are known to exhibit a wide range of rapid behaviors as well as the ability to assess and respond to environmental stimuli. Very early studies have noted a sympathetic-like response of an immediate and rapid response of defense posturing (Bethe, 1897; Huxley, 1880; Shuranova et al., 2006; Wiersma, 1961). The adult crayfish heart is neurogenic since the beat and rhythm are controlled by the central nervous system (Alexandrowicz, 1932; Yamagishi and Hirose, 1997; Yamagishi et al., 1997; Wilkens, 1999). Routinely, measurement of an animal's heart rate (HR) provides a direct measure of excitability and readiness of the internal environment. In crayfish, an increase in HR is noted during defense posturing when presented with a perceived threatening stimulus (Listerman et al., 2000). This type of nervous system response was also shown in spiny lobsters (Yazawa and Katsuyama, 2001).
The ventilatory system is also neuronally controlled with a ventilatory central pattern generator (VPG) responsible for oxygen uptake across the gills by the pumping action of two specialized appendages termed scaphognathites (Mendelson, 1971). A single scaphognathite resides in each branchial chamber at the anterior ends and draws water across the gills by a rhythmic movement (Pasztor, 1968). The VPG, like with the heart, can be modulated by many factors. Previous work in the crab, Cancer magister shows cardiac and ventilatory rhythm can be altered by central control through command neurons (Wilkens et al., 1974). In addition, activity of the VPG is known to change with changes in the internal response and with social interactions or environmental changes in crustaceans (Burmistrov and Shuranova, 1996; Cuadras, 1979, 1980; Li et al., 2000; Listerman et al., 2000; McMahon and Wilkens, 1983; Schapker et al., 2002; Shuranova et al., 2002; Wilkens, 1976). As seen in the crayfish, ventilatory activity (VR) can vary depending on the internal state and changes in VR can be recorded during unexpected external stimuli (Shuranova et al., 1993; Shuranova et al., 2002).
Due to the dependability and feasibility of HR and VR measures, the future questions to be asked are endless. As noted previously, work has been done to examine the autonomic response during social interactions and environmental disturbances. Interestingly, many areas are left to be explored such as examination of the autonomic response during natural hormonal changes and/or other physiological processes occurring in the organism. Possible future directions may also examine sympathetic-like responses during mating and olfactory chemical reception. Most importantly, physiological responses may provide greater insights into an organism response to the environment when compared to behavioral observations since animals do not always respond behaviorally when they are readying themselves internally. Thus, the physiological measurement provides a biological index for assessing an organism's internal state which may explain overall changes in an organism. 
Methods
Electrocardiograms recording (ECGs) procedures for heart and ventilation rates
Before beginning to wire a crayfish, preparation of wires is required. Cut two insulated stainless steel wires (diameter 0.005 inches and with the coating 0.008 inches; A-M Systems, Carlsburg, WA) long enough to insert into the crayfish and adequately reach an impedance detector (UFI, model 2991). Remove insulation (~ 0.5 mm) by fire on the ends of both wires which will be inserted into the carapace. Be careful not remove too much insulation as a bare wire on the outside of the animal must be recoated to prevent short circuiting impedance recordings. Then, using forceps bend the burned ends of the wires in a 90 angle for placement into the carapace. Be sure that the inserted portion of the wire is not too long because this could do damage to internal organs. For feasibility of inserting the wires into a plastic tube (which protects the wires), it is best to loosely twist the wires together. If recording heart and ventilation rates, repeat the above steps a second time and the two sets of wires can be loosely braided together before placement into the plastic tubing. Be sure to designate the wires for heart and ventilation rate so as to not confuse which wires attach to which impedance detector for recording. It is best to label the wires or cut one pair shorter than the other in the beginning.
To feasibly and safely wire the crayfish, it is best to wrap the claws and legs (leaving the back and one side exposed) in wet cloth or paper towels to eliminate injury to the person and/or crayfish (Figure 1). 
<img src="/files/ftp_upload/1594/1594fig1.jpg" style="float:left;" alt="Figure 1" />Figure 1. Wrapping of the chelipeds and legs of the crayfish. Wrapping the crayfish in a wet paper towel will prevent injury to the person and the crayfish. The wrapping will hinder the ability of the crayfish to move the chelipeds and legs.
 To begin the wiring process, it is best to begin in the gill chamber since carapace thickness varies with each crayfish and excess force can cause damage to internal organs, thus by beginning in the gill chamber, excessive pressure is less likely to kill the crayfish as compared to the thoracic chamber. The first hole is made by using a fine point scalpel and just large enough for a wire to be inserted under the cuticle in the rostral area of the gill chamber (i.e., prebranchial chamber to monitor ventilatory rate; Schapker et al., 2002). A large hole will have excessive hemolymph loss and will not seal properly and this will increase the probability of the wires not remaining in the designated hole. Once the wire is in place, drop a small amount of glue (cyanoacrylate ester) and accelerator (HobbyTown USA, Lexington, KY) onto the wire. The use of the fast drying glue reduces handling stress of the animals. Be careful to place only a small amount of accelerator onto the cuticle since this is toxic to the crayfish and will cause death. It is best to immediately wipe any excess accelerator off with paper towel. Repeat these steps for the second wire in the gill chamber. These two wires are placed to span the scaphognathite (respiratory organ) to insure a strong accurate impedance measure during each scaphognathite movement (Figure 2). To get an accurate recording, be sure that the second wire is not placed outside the gill chamber and into the thoracic chamber. 
<img src="/files/ftp_upload/1594/1594fig2.jpg" style="float:left;" alt="Figure 2" />Figure 2. Placement of stainless steel wires in the gill chamber. The white dots on the carapace represent points for the stainless steel wires to be placed under the cuticle. The wires span the scaphognathite and measure movement through dynamic impedance.
The steps for wiring heart rate in the thoracic chamber is the same as above except the wires are placed under the dorsal carapace directly over the heart and span the heart in a rostral-caudal arrangement (Figure 3). This placement will ensure a strong accurate impedance measure during each heart contraction (Listerman et al., 2001).
 <img src="/files/ftp_upload/1594/1594fig3.jpg" style="float:left;" alt="Figure 3" />Figure 3. Placement of stainless steel wires in the thoracic chamber. The white dots on the carapace represent points for the stainless steel wires to be placed under the cuticle. The wires span the heart in a rostral-caudal arrangement. This will allow for measures of heart contraction through dynamic impedance. 
 Once placement of all four wires is complete, mix an instant adhesive (Eastman, 5-min drying epoxy) and coat the holes for each location as well as the wires. Once the wires and drilled holes are adequately covered, use the remaining epoxy to hold the plastic tubing to the back of the crayfish. This will ensure that the wires will not be pulled out of the holes in future handlings. Since the epoxy takes a few minutes to dry, hold or place the crayfish in a container without water until the epoxy is no longer sticky to the touch. Once the epoxy is dry, wash the crayfish in water to remove excess chemicals and place them back into an aquatic environment. To ensure the health of the crayfish, it is best to change the water in approximately one hour to remove any chemicals not previously removed. As noted from previous investigators, we found that prolonged handling alters the physiological measurements due stress for a time duration of 24 h to 3 days (Wilkens et al., 1985); therefore, animals should be left for 2 to 3 days before any experimentation.
To record ECGs, connect the wires from the crayfish to impedance detectors. The two wires spanning the heart (measure HR) should be attached to a single impedance detector and likewise for the two wires spanning the scaphognathite (measure VR). The impedance detector measures dynamic resistance between the two stainless steel wires. These signals are recorded by connecting the detector on-line to a PowerLab via a PowerLab/4SP interface (AD Instruments). All events are measured and calibrated with the PowerLab Chart software version 5.5.6 (AD Instruments, Australia). To begin, set an acquisition rate at 4 kHz. The large deflections will represent heartbeat and ventilation rate. For ease in counting each HR/VR, a smoothing function can applied to the data set. If 100 points were smoothed (i.e. averaged) the resultant trace will depict only the heartbeats. The heart rates can be determined by direct counts of each beat over 10-s intervals and then converted to beats per min (BPM).

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<table border="1">
<tbody>
<tr>
<td>Material Name</td>
<td>Type</td>
<td>Company</td>
<td>Catalogue Number</td>
</tr>	
<tr>
<td>Scalpel (Stainless steel surgical blades No. 11 for No. 3 handle)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Stainless steel wire (diameter 0.005 inches with the coating 0.008 inches; A-M Systems, Carlsburg, WA)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Impedance detector (UFI, model 2991)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Plastic tubing (Tygon Tubing ID .0812, OD .1492, Cole-Parmer Instrument Company)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Fast-drying glue (cyanoacrylate ester, HobbyTown USA, Lexington, KY)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Accelerator (HobbyTown USA, Lexington, KY)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Adhesive (Eastman, 5-min drying epoxy)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td><a href="http://www.adinstruments.com/products/hardware/corporate/product/ML866/" target="_blank">PowerLab via a PowerLab/4SP interface (AD Instruments)</a>.</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td><a href="http://www.adinstruments.com/products/software/research/LabChart-Software/" target="_blank">PowerLab Chart software version 5.5.6 (AD Instruments, Australia)</a></td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td><a href="http://www.adinstruments.com/products/software/research/LabChart-Software/" target="_blank">Chart software version 3.5.6 (AD Instruments, Australia)</a></td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
</tbody>
		</table>

measures of heart and ventilatory rates in freely moving crayfish

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

Watch this Scientific Journal Video about Measures of Heart and Ventilatory Rates in Freely Moving Crayfish at JoVE.com

Measures of Heart and Ventilatory Rates in Freely Moving Crayfish

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

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

measures-of-heart-and-ventilatory-rates-in-freely-moving-crayfish

Research

JoVE Journal

Biology

Heterotopic Heart Transplantation in Mice

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable for studying rejection and immune response now that newer transgenic and gene knockout mice are available, and a large number of immunologic reagents have been developed. The heart transplant model is less stringent than the skin transplant models, although technically more challenging. We have developed a modified technique and have completed over 1000 successful cases of heterotopic heart transplantation in mice. When making anastomosis of the ascending aorta and abdominal aorta, two stay sutures are placed at the proximal and distal apexes of recipient abdominal aorta with the donor s ascending aorta, then using 11-0 suture for anastomosis on both side of aorta with continuing sutures. The stay sutures make the anastomosis easier and 11-0 is an ideal suture size to avoid bleeding and thrombosis.
When making anastomosis of pulmonary artery and inferior vena cava, two stay sutures are made at the proximal apex and distal apex of the recipient s inferior vena cava with the donor s pulmonary artery. The left wall of the inferior vena cava and donor s pulmonary artery is closed with continuing sutures in the inside of the inferior vena cava after, one knot with the proximal apex stay suture the right wall of the inferior vena cava and the donor s pulmonary artery are closed with continuing sutures outside the inferior vena cave with 10-0 sutures. This method is easier to perform because anastomosis is made just on the one side of the inferior vena cava and 10-0 sutures is the right size to avoid bleeding and thrombosis. In this article, we provide details of the technique to supplement the video.

The mouse heterotopic heart transplantation model has been proven by many investigators to be an important method for studying mechanisms of rejection and immune response. However, the techniques involved are still challenging. By modifying standard techniques we have had success with more than 1000 transplants.

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable ...

Implantation of Engineered Tissue in the Rat Heart

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in common laboratory rats or mice and these quite frequently serve as an intermediate step between bench-top experiments and large animal testing or human trials. Given that rodents provide an established, cost-effective, and physiologically-relevant model system in which to test novel combinations of scaffolding materials and cells, they are particularly well-suited for cardiovascular tissue engineering studies. Presently, we describe an open-heart surgical procedure to implant engineered tissue containing myogenic progenitor cells in the atrioventricular (AV) groove of a rat heart. These implants are intended to create an electrical conduit between the right atrium and right ventricle with the ultimate goal of providing an alternative treatment to conventional pacemaker implantation in pediatric patients with complete heart block[1]. The engineered tissue is implanted in the AV-groove by means of a thoracotomy. For our purposes, Lewis rats are anesthetized and invasively ventilated to maintain positive airway pressure during the sterile surgical procedure. The approach to the heart is performed by a right thoracotomy through an antero-lateral incision at the 5th intercostal space. The tissue construct is fixed in the AV groove using a single 7-0 Prolene suture and positioned between the right ventricle and atrium at the ventral portion of the heart. The epicardium is partially removed to allow direct contact between the recipient myocardial cells and those contained in the engineered tissue. Following implantation, the chest wall is closed in layers, any pneumothorax is evacuated, and the animal is extubated and treated with analgesic.

Here, we describe a cardiac surgical procedure to implant engineered tissue in the atrioventricular (AV)-groove of an adult Lewis rat.

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in ...

Recordings of Neural Circuit Activation in Freely Behaving Animals

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

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

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

Visualizing the Beating Heart in Drosophila

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in myogenic hearts. A key step in examining heart function in the fly is finding a way to access the heart in a manner that preserves its myogenic function while still allowing the beating heart organ to be observed and recorded. Two different methods for observing and recording the beating heart in both larva and adult Drosophila are described here. Our semi-intact preparation using adult flies allows clear visualization of the abdominal heart without interference from the pigmented cuticle and overlying fat bodies. To record larval heart beats it is necessary to immobilize the larva, which minimizes body wall movements thereby reducing heart movements that are not associated with myocardial contractions. Our methodologies produce stable adult and larval heart preparations that can beat for hours at rates of 1-3 Hz.

Visualizing the Beating Heart in Drosophila

Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

Historical View and Physiology Demonstration at the NMJ of the Crayfish Opener Muscle

Here we present some of the key important discoveries made with the opener neuromuscular (NMJ) preparation of crustaceans and illustrate that there is still much to learn from this model preparation. In understanding the history one can appreciate why even today this NMJ still offers a rich playground to address questions regarding pre- and post-synaptic function and plasticity. The viability and ease of access to the terminal for intracellular as well as extracellular electrophysiology and imaging are significant advantages. The mechanisms behind the modulation of vesicular kinetics and fusion within the high- and low-output terminals are begging for investigation. The preparation also offers a testable model system for computational assessments and manipulations to examine key variables in theoretical models of synaptic function, for example calcium dynamics during short-term facilitation. The synaptic complexity of active zone and statistical nature of quantal release is also an open area for future investigation both experimentally and computationally.

The opener muscle of the crayfish leg is presented for its historical importance and experimental versatility in muscle phenotype, synaptic physiology and plasticity.

Here we present some of the key important discoveries made with the opener neuromuscular (NMJ) preparation of crustaceans and illustrate that there is ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Quantification of &gamma;H2AX Foci in Response to Ionising Radiation

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA lesions with respect to survival and preservation of genomic integrity. An early response to the induction of DSBs is phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX1. Following induction of DSBs, H2AX is rapidly phosphorylated by the phosphatidyl-inosito 3-kinase (PIKK) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Typically, only a few base-pairs (bp) are implicated in a DSB, however, there is significant signal amplification, given the importance of chromatin modifications in DNA damage signalling and repair. Phosphorylation of H2AX mediated predominantly by ATM spreads to adjacent areas of chromatin, affecting approximately 0.03% of total cellular H2AX per DSB2,3. This corresponds to phosphorylation of approximately 2000 H2AX molecules spanning ~2 Mbp regions of chromatin surrounding the site of the DSB and results in the formation of discrete &gamma;H2AX foci which can be easily visualized and quantitated by immunofluorescence microscopy2. The loss of &gamma;H2AX at DSB reflects repair, however, there is some controversy as to what defines complete repair of DSBs; it has been proposed that rejoining of both strands of DNA is adequate however, it has also been suggested that re-instatement of the original chromatin state of compaction is necessary4-8. The disappearence of &gamma;H2AX involves at least in part, dephosphorylation by phosphatases, phosphatase 2A and phosphatase 4C5,6. Further, removal of &gamma;H2AX by redistribution involving histone exchange with H2A.Z has been implicated7,8. Importantly, the quantitative analysis of &gamma;H2AX foci has led to a wide range of applications in medical and nuclear research. Here, we demonstrate the most commonly used immunofluorescence method for evaluation of initial DNA damage by detection and quantitation of &gamma;H2AX foci in &gamma;-irradiated adherent human keratinocytes9.

Quantification of &amp;#947;H2AX Foci in Response to Ionising Radiation

Quantification of DNA double-strand streaks using &gamma;H2AX formation as a molecular marker has become an invaluable tool in radiation biology. Here we demonstrate the use of an immunofluorescence assay for quantification of &gamma;H2AX foci after exposure of cells to radiation.

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA ...

Evaluation of the Spatial Distribution of &gamma;H2AX following Ionizing Radiation

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone H2AX1,2. This phosphorylation of H2AX is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)3. The phosphorylated form of H2AX, referred to as &gamma;H2AX, spreads to adjacent regions of chromatin from the site of the DSB, forming discrete foci, which are easily visualized by immunofluorecence microscopy3. Analysis and quantitation of &gamma;H2AX foci has been widely used to evaluate DSB formation and repair, particularly in response to ionizing radiation and for evaluating the efficacy of various radiation modifying compounds and cytotoxic compounds4.
Given the exquisite specificity and sensitivity of this de novo marker of DSBs, it has provided new insights into the processes of DNA damage and repair in the context of chromatin. For example, in radiation biology the central paradigm is that the nuclear DNA is the critical target with respect to radiation sensitivity. Indeed, the general consensus in the field has largely been to view chromatin as a homogeneous template for DNA damage and repair. However, with the use of &gamma;H2AX as molecular marker of DSBs, a disparity in &gamma;-irradiation-induced &gamma;H2AX foci formation in euchromatin and heterochromatin has been observed5-7. Recently, we used a panel of antibodies to either mono-, di- or tri- methylated histone H3 at lysine 9 (H3K9me1, H3K9me2, H3K9me3) which are epigenetic imprints of constitutive heterochromatin and transcriptional silencing and lysine 4 (H3K4me1, H3K4me2, H3K4me3), which are tightly correlated actively transcribing euchromatic regions, to investigate the spatial distribution of &gamma;H2AX following ionizing radiation8. In accordance with the prevailing ideas regarding chromatin biology, our findings indicated a close correlation between &gamma;H2AX formation and active transcription9. Here we demonstrate our immunofluorescence method for detection and quantitation of &gamma;H2AX foci in non-adherent cells, with a particular focus on co-localization with other epigenetic markers, image analysis and 3D-modeling.

Evaluation of the Spatial Distribution of &amp;#947;H2AX following Ionizing Radiation

Microscopic analysis of &gamma;H2AX foci, which form following the phosphorylation of H2AX at Ser-139 in response to DNA double-strand breaks, has become an invaluable tool in radiation biology. Here we used an antibody to mono-methylated histone H3 at lysine 4 as an epigenetic marker of actively transcribing euchromatin, to evaluate the spatial distribution of radiation-induced &gamma;H2AX formation within the nucleus.

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

Measurement of mRNA Decay Rates in Saccharomyces cerevisiae Using rpb1-1 Strains

mRNA steady state levels vary depending on environmental conditions. Regulation of the steady state accumulation levels of an mRNA ensures that the correct amount of protein is synthesized for the cell&#8217;s specific growth conditions. One approach for measuring mRNA decay rates is inhibiting transcription and subsequently monitoring the disappearance of the already present mRNA. The rate of mRNA decay can then be quantified, and an accurate half-life can be determined utilizing several techniques. In S. cerevisiae, protocols that measure mRNA half-lives have been developed and include inhibiting transcription of mRNA using strains that harbor a temperature sensitive allele of RNA polymerase II, rpb1-1. Other techniques for measuring mRNA half-lives include inhibiting transcription with transcriptional inhibitors such as thiolutin or 1,10-phenanthroline, or alternatively, by utilizing mRNAs that are under the control of a regulatable promoter such as the galactose inducible promoter and the TET-off system. Here, we describe measurement of S. cerevisiae mRNA decay rates using the temperature sensitive allele of RNA polymerase II. This technique can be used to measure mRNA decay rates of individual mRNAs or genome-wide.

Measurement of mRNA Decay Rates in Saccharomyces cerevisiae Using rpb1-1 Strains

The steady state level of specific mRNAs is determined by the rate of synthesis and decay of the mRNA. Genome-wide mRNA degradation rates or the decay rates of specific mRNAs can be measured by determining mRNA half-lives. This protocol focuses on measurement of mRNA decay rates in Saccharomyces cerevisiae.

mRNA steady state levels vary depending on environmental conditions. Regulation of the steady state accumulation levels of an mRNA ensures that the ...