This work was supported by the National Institutes of Health R01 HL141452 to TPN. ADInstruments kindly provided generous funding to defray the cost of open access publishing but had no role in either experimental design, data acquisition, data analysis of this study or any access to the manuscript prior to publication.

All experiments in this study were conducted in accordance with the US National Institutes of Health Guide for the Care and Use of Laboratory Animals. All animal protocols in this study were approved by the UCLA Institutional Animal Care and Use Committee.
1. Preparation of the experimental set up
<ol>
	<li>Maintain zebrafish in flow-through aquarium systems on a 14 h light, 10 h dark photoperiod at 28 &#176;C &#177; 0.5 &#176;C. Feed with flake food daily and live brine shrimp (Artemia nauplii) twice daily. Zebrafish in this study were maintained and fed by the UCLA Zebrafish Core.</li>
	<li>On the day of the experiment, transport zebrafish from the aquarium to the laboratory.</li>
	<li>Set up the in vivo ECG recording system by connecting the essential pieces of equipment and inserting the three color-coded stainless-steel electrodes into the three color-matched access portals of the amplifier (Figure 3). Start the system at the start of an ECG recording and/or analysis session.</li>
	<li>Procure&#160;necessary tools, such as a timer/stopwatch, a wet sponge with a slit to hold the fish, forceps, scissors, Pasteur pipettes, and culture dishes (100 mm x 20 mm).</li>
</ol>
2. Anesthesia induction
<ol>
	<li>Prepare immersion anesthesia for pain control and fish immobilization to avoid motion artifacts during ECG data acquisition. Most laboratories use immersion tricaine (ethyl 3-aminobenzoate methanesulfonate, MS-222).
	<ol>
		<li>To make the tricaine 0.4% stock solution, combine the following items in a screw-capped dark glass bottle: 400 mg of tricaine powder, 98 mL of double distilled water, and 2 mL of 1 M Tris (pH 9). Adjust to pH 7.0 using 1 N NaOH or 1 N HCl as needed8.</li>
		<li>To make the tricaine final immersion solution, determine the minimum concentration that is appropriate for the zebrafish age9, size, metabolic state, strain, disease model, scientific objectives, and procedural duration.</li>
		<li>Perform a tricaine concentration-response study, titrating up or down from the recommended concentration of 168 mg/L (or 0.0168%)9 if necessary, to attain&#160;level 4 of anesthesia within 3 min with the fewest possible cardiorespiratory toxicities. For example, in this study, the immersion of wild-type AB zebrafish of 12-18 months of age into a 0.02-0.04% tricaine solution will induce level 4 of anesthesia within 3 min. 
		NOTE: At level 4 of anesthesia, equilibrium and muscle tone are completely lost and opercular movement rate is reduced8.</li>
		<li>If necessary, consult the veterinarian in the Institutional Animal Care and Use Committee (IACUC) for additional guidance on the appropriateness of the selection of anesthetic(s) and route of administration.</li>
	</ol>
	</li>
	<li>Immerse an adult zebrafish into a dish containing tricaine solution of the lowest predetermined and IACUC-approved concentration (e.g., 0.02-0.04% in this study) to induce level 4 of anesthesia within 3 min (Figure 2).
	<ol>
		<li>For survival ECG protocol, keep the ECG recording session as brief as possible (under 10 min). For brief ECG recording sessions lasting less than 15 min, anesthesia maintenance is not necessary.</li>
		<li>For long ECG recording sessions lasting hours, use a long-acting intramuscular paralytic and an oral perfusion system to provide ample hydration and oxygenation6.</li>
	</ol>
	</li>
</ol>
3. ECG lead placement
<ol>
	<li>Once the zebrafish maintains level 4 of anesthesia for 3 s, use a pair of blunt forceps to transfer the fish immediately onto the damp sponge slit with its ventral surface uppermost for placement of ECG lead electrodes (Figure 4).</li>
	<li>Gently insert the three ECG lead electrodes into the fish musculature to approximately 1 mm in depth to establish a bipolar lead in the frontal plane that parallels the left caudal-right cranial orientation of the cardiac main axis.</li>
	<li>Position the positive (red) electrode in the ventral midline at the level of the bulbus arteriosus, i.e., at 1-2 mm above an imaginary line connecting the two lower edges of the operculums (Figure 4A).</li>
	<li>Position the negative (black) electrode caudally and 0.5-1.0 mm left laterally to the positive electrode, at a distance greater than the maximal apicobasal length of the adult zebrafish ventricle (Figure 4A).</li>
	<li>Position the reference (green) electrode caudally, near the anal region. 
	NOTE: Since the cardiac main axis varies somewhat from fish to fish, to maximize the R and T wave amplitudes, adjust the lead positions by making only small, systematic changes through trial and error. For example, change one electrode (positive or negative), instead of both electrodes, at a time and make gradual changes in one specified direction before changing to another direction instead of making erratic changes in random directions.</li>
</ol>
4. ECG recording
<ol>
	<li>Open the ECG data acquisition program. Select a desired setting from the drop-down menus for range, low pass, and high pass. For example, the following setting in the in vivo ECG recording system used in this experiment yields consistent, satisfactory signal-to-noise ratio for a normal adult zebrafish: range &#8220;2 mV&#8221;, low pass &#8220;120 Hz&#8221;, and high pass &#8220;0.03 s&#8221;.</li>
	<li>Press Start to start continuous gap-free ECG recording at a sampling rate of 1 kHz.</li>
	<li>To optimize lead positioning for maximal signal-to-noise ratio, press Stop to stop ECG recording and review the ECG trace soon after the very first recording attempt for each heart.&#160; To diagnose that an adult zebrafish ECG is normal, confirm that&#160;all of the following four validating criteria are satisfied (Figure 1):
	<ol>
		<li>Criterion 1: Ensure that all ECG waveforms (P, QRS, and T) are distinct and readily visible.</li>
		<li>Criterion 2: Ensure that the P wave is positive.</li>
		<li>Criterion 3: Ensure that the net QRS complex is positive (i.e. the R wave amplitude is larger than the sum of Q and S wave amplitudes).</li>
		<li>Criterion 4: Ensure that the T wave is positive.</li>
	</ol>
	</li>
	<li>If a normal ECG is expected, reposition the electrodes (try the negative electrode first) if necessary, until all four validating criteria are satisfied.</li>
	<li>If a normal T wave is expected, but the T wave is too small, reposition the electrodes to maximize the T wave amplitude.</li>
	<li>Resume ECG recording after optimizing lead positioning. Save the ECG sweeps for subsequent analysis.</li>
</ol>
5. Recovery from anesthesia
<ol>
	<li>At the end of the ECG recording session, carefully remove the electrodes without injuring&#160;the fish. Transfer the fish&#160;to fresh, oxygenated fish water free of tricaine.</li>
	<li>To facilitate recovery from anesthesia, squirt water over the gills vigorously with a Pasteur pipette until the fish resumes regular gill movement or swimming.</li>
	<li>Monitor the fish for full recovery from anesthesia (typically 1-2 min), as indicated by the fish ability to swim upright for at least 5 s.</li>
</ol>
6. ECG interpretation
<ol>
	<li>Define the analysis settings.
	<ol>
		<li>Know the software interface (Table of Materials) by reading the operating manual of the ECG data analysis software. 
		NOTE: Although the directions below are specific to the commercial software used in our laboratory, the basic tasks to accomplish are essentially the same in any software package for ECG analysis.</li>
		<li>Open the ECG data analysis program. From the File menu, select Open to open the ECG file of interest and display the full ECG trace. Use the mouse to drag out a section of interest in the ECG trace to analyze.</li>
		<li>From the ECG Analysis menu, select ECG Settings to open a dialogue box to pre-define various parameter settings for software automatic analysis (Figure 5A).</li>
	</ol>
	</li>
	<li>Analyze the heart rhythm and rate. 
	NOTE: Heart rate depends on several factors, including zebrafish age and strain, anesthesia agents (e.g., tricaine, isoflurane, etc.) and concentration, anesthesia usage (single agent5,7 vs. combined agents5) and exposure time5. For example, in this study the heart rate of 12-18 month-old wild-type AB zebrafish after 3-5 min of immersion in 0.02-0.04% tricaine solution was 116 &#177; 17 beats per minute (n = 9), consistent with literature reports of heart rate for this age group and anesthetic5,7.
	<ol>
		<li>Determine whether the heart rhythm is sinus or not, regular or irregular. 
		NOTE: The presence (or absence) of sinus rhythm is based on the presence (or absence) of an upright P wave preceding each QRS by a normal PR interval (e.g., 60-65 ms for Liu et al.&#8217;s 10-12 month-old7 and 12-18 month-old wild-type AB zebrafish in this study). The atrial and ventricular rhythm regularity (or irregularity) is based on the regularity (or irregularity) of successive PP or RR intervals, respectively.</li>
		<li>To determine the heart rate, ensure that the software correctly identifies all the P and R waves. Based on these automatic identifications (or manual corrections) of the P and R waves, the software automatically measures all the PP and RR intervals in the ECG selection, calculates the interval averages to generate the atrial and ventricular rate. 
		NOTE: The atrial rate is the average PP interval whereas the ventricular rate is the average RR interval. To determine the heart rate, correct identification of the P and R waves is critical.</li>
		<li>Correct any auto-identification mistakes by moving the misplaced cursors to the appropriate P and R waves (Figure 5B). 
		NOTE: If the heart is in sinus rhythm, the atrial rate and ventricular rate are the same because of the one-to-one correspondence between the sinus P waves and the QRS complexes. However, in the case of atrioventricular dissociation (e.g., in ventricular tachycardia or third-degree atrioventricular block), this one-to-one correspondence between the P waves and QRS complexes is lost; therefore, there are two heart rates because the atrial rate is different from the ventricular rate.</li>
		<li>Determine the heart rate based on at least five consecutive complete cardiac cycles if the heart rhythm is regular, or a strip of at least six seconds if the heart rhythm is irregular.</li>
	</ol>
	</li>
	<li>Calculate&#160;intervals and wave durations.
	<ol>
		<li>Go to ECG Analysis &#62; Averaging View to concatenate n (e.g., 5) consecutive cardiac cycles into a single average signal (Figure 5C). 
		NOTE: If the ECG waveforms of an individual cardiac cycle diverge substantially from the average signal, study that cardiac cycle separately without concatenation.</li>
		<li>Ensure that the software correctly identifies the start and end of the P wave, QRS complex, and T wave displayed in the Averaging View window (Figure 5C). Based on these automatic identifications (or manual corrections) of these waves and intervals, the software automatically measures the durations as defined conventionally. 
		NOTE: The PR interval extends from the start of the P wave to the start of the QRS complex (or the RS complex if the Q wave is not visible). The QRS duration extends from the start of the Q wave (or the R wave if the Q wave is not visible) to the end of the S wave (i.e., the J point; Figure 1). The QT interval extends from the start of the Q wave (or the R wave if the Q wave is not visible) to the end of the T wave. Therefore, to calculate intervals and durations, correct identification of the start and end of the P wave, QRS complex, and R wave is critical.</li>
		<li>Correct any auto-identification mistakes by moving the misplaced cursors to the appropriate positions.</li>
		<li>Select the negative peak of the S wave as the end of the QRS complex7 because the zebrafish J point that signals the end of the S wave can be particularly difficult to identify accurately. This will cause a slight underestimate of the true QRS duration. 
		NOTE: The ECG analysis software automatically corrects the QT interval to the ventricular rate (or RR interval) to generate the corrected QT interval QTc using the method pre-selected by the user in step 6.1.3, for example, Bazett (Figure 5A). The Bazett&#8217;s formula (1920) QTc = QT / &#8730;RR is the most popular and the first of several methods proposed to correct the human QT interval for heart rate. Because the accuracy of the Bazett&#8217;s formula has been questioned, refer to other methods proposed for humans10,11 and zebrafish6 (Figure 5D).</li>
	</ol>
	</li>
	<li>Interpret ECG abnormalities by recognizing exceptions for the four validating criteria in step 4.3.
	<ol>
		<li>Recognize exceptions for criterion 1. In the absence of any P waves (which indicates the absence of sinus rhythm), rely on the RR intervals and QRS duration to diagnose the heart rhythm. For example, if the RR intervals are irregularly irregular, diagnose atrial fibrillation; if the RR intervals are regular and the QRS is normally narrow, diagnose junctional escape rhythm; on the other hand, if the RR intervals are regular and the QRS is abnormally prolonged, diagnose ventricular escape rhythm.</li>
		<li>Recognize exceptions for criterion 2. When the P wave is negative (or inverted), diagnose retrograde atrial activation from an ectopic pacemaker (such as an atrial site downstream of the sinus node, the atrioventricular node, or the ventricle).</li>
		<li>Recognize exceptions for criterion 3. When tall and narrow Q waves present with negative P and negative T waves, diagnose lead reversal due to an erroneous switch of the positive and negative electrode positions because those tall and narrow Q waves were true R waves mistakenly inverted (Figure 6D). In contrast, when broad Q waves present with positive P waves following significant cardiac injury, diagnose myocardial infarction because those broad Q waves are true pathologic Q waves.</li>
		<li>Recognize exceptions for criterion 4. When the T wave is inverted, inspect ventricular activation to identify whether the ventricular repolarization abnormality is primary or secondary. Rely on the clinical scenario to narrow down the correct diagnosis from a differential list of primary ventricular repolarization abnormality (from drug effects or myocardial ischemia; Figure 6C) vs. secondary ventricular repolarization abnormality (due to aberrant ventricular activation from pre-excitation, ventricular ectopy, or ventricular pacing).</li>
	</ol>
	</li>
	<li>Export ECG findings.
	<ol>
		<li>Select Table View to review all ECG measurements. Select the measurements of interest to copy and paste into the desired document (e.g., Excel spreadsheet).</li>
		<li>To export an ECG trace, highlight a section of interest in the ECG sweep using the magnifier icon. Copy and paste into the desired document (e.g., Word or PowerPoint).</li>
	</ol>
	</li>
</ol>

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Open Access for this video-article is sponsored by AD Instruments.

When recording in vivo ECG for adult zebrafish by means of a single lead as we demonstrated in this study, there are a number of caveats concerning the quality and validity of the ECG recording results. First, in choosing the appropriate anesthetics and determining the minimal needed anesthesia concentration, depth, and duration, balance the anesthetic cardiotoxicities against the critical need to suppress motion artifacts and the a priori determination for a survival vs. terminal experimental design. Capitalizing on the synergistic potency of a combination of multiple anesthetics from different drug classes5,14 and paralytics1,6 to lower the dose of individual agents5 or administering a low maintenance dose following a higher induction dose are typical strategies. However, despite its well-known potential cardiorespiratory toxicities, including death8, tricaine is still the most widely used, the best available, and the only anesthetic approved by the US Food and Drug Administration (FDA)&#160; for zebrafish anesthesia. Tricaine has been popularly used in ECG recording of adult zebrafish either as a single agent or in combination with other anesthetics or paralytics.
Second, lead placement accuracy can be ensured at least for healthy normal zebrafish using our four validating criteria for a normal adult zebrafish ECG. Of the four validating criteria that we propose here, the last two criteria together confirm the fundamental concordance between the polarity of the R wave and that of the T wave in a normal ECG5,7,15. This R and T wave concordance is a fortuitous, yet critical, similarity between zebrafish and human16,17 normal ECG that contributes to the clinical relevance of the zebrafish heart model as a surrogate for human cardiac electrophysiology. However, several benign or malignant conditions may invalidate any of the four validating criteria. For example, the R and T wave concordance is lost in myocardial ischemia7,15. This loss of R and T wave concordance in myocardial ischemia is another striking resemblance between zebrafish and human ECG that contributes to the clinical relevance of the zebrafish myocardial infarction model.
Lastly, we recommend a standard practice in ECG analysis. With the advent of technology, ECG analysis software can generate automatic ECG interpretation. However, we strongly recommend that trained humans should always re-interpret and verify all ECGs based on the respective clinical scenario leading to ECG recording. Routine over-reliance solely on automatic interpretation by an ECG analysis software is inadvisable, particularly in the presence of common normal ECG variants, cardiac pathologies, or suboptimal lead placement.
This study focuses on the minimally invasive method for brief ECG recording sessions. However, should the need arise for terminal prolonged ECG recording sessions lasting hours, modifications are necessary to provide adequate oxygenation, hydration, and anesthesia by continuous perfusion6.
Additionally, enhance the signal-to-noise ratio by one of at least three ways. Choosing a more powerful amplifier is often a costly, if not impractical, option. Opening the pericardial sac to reduce the volume conductor is a reasonable, although invasive, approach that has been adopted7. Strategic lead placement to align the lead axis in a direction parallel to the main cardiac axis (Figure 4B) will maximize the ECG voltage signals but may require trial and error, especially in the absence of pericardiotomy.
The in vivo ECG interrogation method for adult zebrafish that we presented here offers four main advantages. First, our minimally invasive approach requires only electrode insertion, but no fish scale removal or thoracotomy-pericardiotomy. Therefore, by minimizing pain for the fish, our approach enables repeated ECG interrogations in longitudinal survival studies. Second, when anesthetics adequately suppress fish motion, the in vivo ECG recording system in our study consistently yields a satisfactory signal-to-noise ratio with noise-free raw signals. Third, the four-criterion quality validation that we propose here ensures data accuracy and robustness early in the ECG data acquisition and minimizes operator-dependent variations. Lastly, in particular, our last validating criterion (the normal T wave is upright) encapsulates the concordance of the R wave and T wave, an important human-like feature of zebrafish normal ECG (Figure 1).
However, there still exist four major limitations to current in vivo ECG methodology for adult zebrafish by our group and others.
First, the lack of subject cooperation necessitates the need for anesthesia with its limiting cardiorespiratory toxicity consequences. For in vivo ECG interrogation, whereas human patients never need sedation, zebrafish always require anesthetics or paralytics, all of which cause variable cardiorespiratory toxicities.
Second, the need to secure the attached ECG leads slightly elevates the invasiveness of an otherwise non-invasive procedure. Whereas lead placement in body-surface ECG recording of humans is entirely non-invasive because electrodes adhere to the human epidermis, lead placement for in vivo ECG recording of zebrafish is more invasive because, at the minimum, steel electrodes must puncture the fish skin for secure insertion into the fish musculature.
The last two limitations stem from the anatomical constraints of the zebrafish chest and heart. Third, the minuscule size of the adult zebrafish heart necessitates a drastic reduction in the number of ECG leads. While humans readily accommodate twelve leads in a standard ECG recording, adult zebrafish can typically accommodate only a single unipolar or bipolar lead. The ramification of a single ECG lead is the challenge to optimize concurrently the amplitudes of all three P, R, and T waves. Hence, the importance of optimal and accurate lead placement in zebrafish ECG interrogation cannot be overstated. In zebrafish, the T wave presents a unique detection challenge because it is often the smallest of these three waves. Therefore, the zebrafish T wave amplitude should receive optimization priority over the typically larger P and R waves.
Fourth, determining the zebrafish main cardiac axis to maximize the R wave amplitude can be challenging. The reason is that the zebrafish heart has more freedom of motion within its loose pericardial sac compared to the human heart within its form-fitting glove-like pericardium.
Overall, these limitations will stimulate future method innovation. With the advent of 3D printing and deformable electronics18, there is hope for direct lead implantation one day in awake, alert, swimming zebrafish using a &#8216;cardiac sock&#8217; of wireless electrode sensors.

The zebrafish (Danio rerio) heart is located anteroventrally to the thoracic cavity between the operculum and the pectoral girdles. The heart is enclosed rather loosely within a silver-colored pericardial sac. Anatomically, the zebrafish heart is different from the four-chambered human and other mammalian hearts because of its diminutive scale (100-fold smaller than the human heart) and its two-chambered structure consisting of only one atrium and one ventricle. Nonetheless, the electrocardiogram (ECG) waveforms and the duration of the QT interval of both species are remarkably similar (Figure 1). Accordingly, zebrafish has emerged as a popular model for studying human inherited arrhythmias1,2,3 and for high-throughput drug screening of potential human cardiotoxicities4,5, such as QT prolongation.
In the routine evaluation of human cardiac diseases, the body-surface ECG has become the most extensively used first-line non-invasive diagnostic tool since its invention by Einthoven in 1903. In contrast, since the first adaptation of the body-surface ECG recording method for adult zebrafish in 20066 and several modifications thereafter7, this technique has remained largely inaccessible to many researchers in the field despite the popularity of this animal model. For other researchers who performed in vivo ECG interrogation for adult zebrafish, wide variations among operators led to inconsistency in ECG findings from different studies. Common reasons include cumbersome and expensive specialized devices and software, low signal-to-noise ratio, and confusion regarding the electrode placement, all further aggravated by an incomplete understanding of the adult zebrafish ECG features and underlying tissue mechanisms. Given that in vivo ECG is the only diagnostic tool to electrically phenotype live zebrafish, there is a clear need for a standardized method to improve sensitivity and specificity, reproducibility and accessibility.
Here, we present a practical, reliable, and validated approach to record and interpret zebrafish in vivo electrocardiograms (Figure 2). Using a single bipolar lead in the frontal plane, we investigated the changes in ECG waveforms and interval durations of live anesthetized healthy wild-type AB adult zebrafish.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Culture dishes</td>
			<td>Fisher Scientific</td>
			<td>FB087571</td>
			<td>100 mm x 20 mm</td>
		</tr>
		<tr>
			<td>Dumont Forceps</td>
			<td>Fine Sciense Tools</td>
			<td>11253-20</td>
			<td>0.1 x 0.06 mm</td>
		</tr>
		<tr>
			<td>FE136 Animal Bio Amp&#160;</td>
			<td>AD Instruments</td>
			<td>FE231</td>
			<td></td>
		</tr>
		<tr>
			<td>Iris Forceps</td>
			<td>Fine Sciense Tools</td>
			<td>11064-07</td>
			<td>0.6 x 0.5 mm</td>
		</tr>
		<tr>
			<td>LabChart 8 Pro</td>
			<td>AD Instruments</td>
			<td></td>
			<td>Software with ECG Module</td>
		</tr>
		<tr>
			<td>Needle electrodes for Animal Bio Amp&#160;</td>
			<td>AD Instruments</td>
			<td>MLA1213</td>
			<td>29 gauge</td>
		</tr>
		<tr>
			<td>Plastic Disposable Transfer Pipets</td>
			<td>Fisher Scientific</td>
			<td>13-669-12</td>
			<td>6 in., 1.2 mL</td>
		</tr>
		<tr>
			<td>PowerLab 4/35</td>
			<td>AD Instruments</td>
			<td>4//35</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>Fine Sciense Tools</td>
			<td>15000-08</td>
			<td>2.5 mm, 0.075 mm</td>
		</tr>
		<tr>
			<td>Tricaine (Ethyl 3-aminobenzoate methanesulfonate)</td>
			<td>Sigma</td>
			<td>E10521-10G</td>
			<td>MS-222</td>
		</tr>
	</tbody>
</table>

in vivo surface electrocardiography for adult zebrafish

The electrocardiogram waveforms of adult zebrafish and those of humans are remarkably similar. These electrocardiogram similarities enhance the value of zebrafish not only as a research model for human cardiac electrophysiology and myopathies but also as a surrogate model in high throughput pharmaceutical screening for potential cardiotoxicities to humans, such as QT prolongation. As such, in vivo electrocardiography for adult zebrafish is an electrical phenotyping tool that is necessary, if not indispensable, for cross-sectional or longitudinal in vivo electrophysiological characterizations. However, too often, the lack of a reliable, practical, and cost-effective recording method remains a major challenge preventing this in vivo diagnostic tool from becoming more readily accessible. Here, we describe a practical, straightforward approach to in vivo electrocardiography for adult zebrafish using a low-maintenance, cost-effective, and comprehensive system that yields consistent, reliable recordings. We illustrate our protocol using healthy adult male zebrafish of 12-18 months of age. We also introduce a rapid real-time interpretation strategy for quality validation to ensure data accuracy and robustness early in the electrocardiogram recording process.

Figure 1 illustrates the clinical relevance of the method presented here. In vivo surface electrocardiography for adult zebrafish is an essential electrical phenotyping tool because of the remarkable similarities between the zebrafish and human ECG despite their vast anatomical differences. The zebrafish heart has only one atrium and one ventricle in contrast to the human heart with two atria and two ventricles (top row; right and left, respectively). However, despite its apparent anatomical simplicity, the zebrafish heart shares several ECG features with the human heart (bottom row; right and left, respectively) Therefore, the zebrafish heart has emerged as a surrogate model for human cardiac electrophysiology5,12,13. Figure 1 illustrates a small but distinct Q wave from a live, healthy 14-month-old zebrafish. However, in zebrafish ECG, lead positioning is not commonly optimized to demonstrate the Q wave. Therefore, the Q wave is commonly invisible, and an RS complex is more commonly seen than the full QRS complex in zebrafish ECG.
Figure 2 summarizes the four essential action steps to conduct minimally invasive in vivo electrocardiography for adult zebrafish. Following anesthesia induction (step 1) and electrode placement (step 2), we recorded baseline ECG signals (step 3) from healthy wild-type AB zebrafish of 12 to 18 months of age (n = 9). Our electrode insertion technique was only minimally invasive because we did not need to peel fish scales or perform pericardiotomy. Following data acquisition, we manually reviewed and verified each ECG recording (step 4) to avoid potential misinterpretation by software automatic analysis.
Figure 3 shows the three indispensable components of a typical ECG data acquisition and processing system: a high-performance data acquisition hardware, a high-gain differential amplifier, and a computer uploaded with software for ECG data acquisition and analysis. In our laboratory, we adapted an existing commercial in vivo ECG recording system originally designed for small mammalian models (such as mice, rats, and rabbits) to accommodate the adult zebrafish model.
Figure 4 demonstrates that proper lead placement requires aligning the lead with the presumed cardiac main axis. In zebrafish in vivo ECG recording, because only one single lead is used, proper lead positioning to maximize concurrently both R and T wave amplitudes is critical. To maximize R and T wave amplitudes, we aligned the positive and negative lead electrodes with the cardiac main axis, presumably in the left caudal to right cranial orientation. Following thoracotomy and pericardiotomy to open the pericardial sac and expose the heart, the cardiac main axis becomes apparent (Figure 4B white dashed line). In fact, pericardiotomy to expose the heart is a commonly used strategy to increase the signal-to-noise ratio7 at the cost of converting the ECG recording from a minimally invasive into a highly invasive procedure.
Figure 5 illustrates critical steps in ECG analysis. First, we pre-defined the various parameter settings for software automatic analysis using the ECG Settings dialogue box (Figure 5A). Because we repurpose an existing ECG recording equipment designed for mammalian models to accommodate adult zebrafish, the Detection and Analysis setting for zebrafish is not available. We selected the Human Preset instead, given the remarkable similarity of zebrafish ECG to human ECG (Figure 5A). Second, we manually verified the software automatic ECG identification (in black) of the R wave peaks and correct (in red) any R wave auto-identification mistakes prior to commanding the software to recalculate the average ventricular rate. For example, in Figure 5B, a large P wave in relation to the R wave fooled the software into misidentifying the R waves, leading to the subsequent automatic miscalculation of the RR interval or ventricular rate. Therefore, human verification and appropriate corrections as needed are critical in ECG analysis. Third, we quickly assessed rhythm regularity and calculated the average duration of waves and intervals using the Averaging View (Figure 5C) to concatenate several consecutive cardiac cycles (green) into one single average signal (black). Here in Figure 5C, the negligible deviation between each of the nine cardiac cycles and the average signal argues for the excellent rhythm regularity of this zebrafish heart. Lastly, we enabled the software to automatically correct the QT interval for heart rate using Bazett, one of the seven different methods available (Figure 5D).
Figure 6A-C demonstrates how the depth of electrode placement affects the amplitudes of the ECG signals. When we incorrectly inserted the electrodes too superficially in the dermis (Figure 6A), the lead was &#8220;indirect&#8221;-like (more than two cardiac diameters from the heart, similar to the indirect standard human ECG limb leads I, II, and III) and the voltage signals were small. When we appropriately inserted the electrodes 1 mm deeper into the pectoralis musculature (Figure 6B), the lead became &#8220;semidirect&#8221; (in close proximity but not in direct contact with the heart) and the voltage signals increased. The ECG waveforms became readily visible. However, when we incorrectly inserted the electrodes even deeper into the ventricle (Figure 6C), the lead became &#8220;direct&#8221; (in direct contact with the heart) and the voltage signals increased further. The R wave amplitude in Figure 6C increased by eight-fold compared to Figure 6A and by four-fold compared to Figure 6B. However, the ECG trace in Figure 6C revealed new signs of injury to the ventricular myocardium, such as new ST depression and new T wave inversion.
Figure 6D demonstrates how the unusual inversions of all ECG waveforms (P, Q, R, S, and T) should signal a lead reversal mistake, in which the positive and negative electrodes switched place. Note that by definition Q and S are always negative whereas R is always positive.
Figure 6E-F shows how inappropriate anesthesia depth can impair the quality of in vivo ECG recording. In Figure 6E, inadequate anesthesia (0.017% tricaine) led to failure to immobilize the zebrafish completely. The resultant motion artifacts lowered the signal-to-noise ratio by both contaminating the signal (asterisk) and increasing the noise (arrows). In contrast, in Figure 6F, overdosed anesthesia (0.08% tricaine) induced severe sinus bradyarrhythmia as well as changes of the ST segment and T wave.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60011/60011fig1.jpg" /> 
Figure 1: Contrasting anatomy and ECG of human and zebrafish hearts. In contrast to the human heart with two atria and two ventricles, the zebrafish heart has only one atrium and one ventricle (top row). Abbreviations: RA, right atrium; LA, left atrium; RV, right ventricle; LV: left ventricle. The zebrafish heart shares several common ECG features with the human heart (bottom row). <a href="https://www.jove.com/files/ftp_upload/60011/60011fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60011/60011fig2.jpg" /> 
Figure 2: Minimally invasive in vivo ECG recording protocol. A schematic flow chart illustrates four critical action steps in conducting an in vivo ECG interrogation: induce anesthesia, place ECG lead electrodes, record ECG, and analyze the ECG recordings. <a href="https://www.jove.com/files/ftp_upload/60011/60011fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60011/60011fig3.jpg" /> 
Figure 3: ECG data acquisition and processing system. The three key components of an integrated in vivo ECG recording system include a hardware to acquire data, an amplifier, and computer software for data acquisition and analysis. The amplifier comes with three ready-to-use 29-gauge stainless steel microelectrodes. <a href="https://www.jove.com/files/ftp_upload/60011/60011fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60011/60011fig4.jpg" /> 
Figure 4: ECG lead placement. Three 29-gauge color-coded stainless steel electrodes are inserted securely into the fish musculature to approximately 1 mm in depth. Placement of the negative (black) electrode and the positive (red) electrode establishes a bipolar lead in the frontal plane, along a left caudal to right cranial orientation. Abbreviation: ref, reference electrode <a href="https://www.jove.com/files/ftp_upload/60011/60011fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60011/60011fig5.jpg" /> 
Figure 5: Critical steps in ECG analysis. (A) Pre-define the various parameter settings for software automatic analysis. (B) Manually correct (red) two automatic misidentifications by the software (black) of the P and R waves to rectify software miscalculation of the atrial and ventricular rate. (C) Concatenate nine consecutive cardiac cycles (green) into a single average signal (black) to quickly assess rhythm regularities/irregularities and calculate average durations of waves and intervals. (D) Correct the QT interval for heart rate using one of the various methods, such as Bazett. <a href="https://www.jove.com/files/ftp_upload/60011/60011fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60011/60011fig6.jpg" /> 
Figure 6: Effects of lead placement and anesthesia depth on ECG signals. Two most critical steps that determine the success of in vivo ECG recording are lead placement (A-D) and anesthesia depth (E-F). <a href="https://www.jove.com/files/ftp_upload/60011/60011fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Vivo Surface Electrocardiography for Adult Zebrafish at JoVE.com

In Vivo Surface Electrocardiography for Adult Zebrafish

Here, we present a reliable, minimally invasive, and cost-effective method to record and interpret electrocardiograms in live anesthetized adult zebrafish.

The electrocardiogram waveforms of adult zebrafish and those of humans are remarkably similar. These electrocardiogram similarities enhance the value of ...

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13.9K Views.  David Geffen School of Medicine at UCLA. Here, we present a reliable, minimally invasive, and cost-effective method to record and interpret electrocardiograms in live anesthetized adult zebrafish.

Video: In Vivo Surface Electrocardiography for Adult Zebrafish

Transplantation of Whole Kidney Marrow in Adult Zebrafish

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an organism. The functional definition of a HSC is a transplanted cell that has the ability to reconstitute all the blood lineages of an irradiated recipient long term. This designation was established by decades of seminal work in mammalian systems. Using hematopoietic cell transplantation (HCT) and reverse genetic manipulations in the mouse, the underlying regulatory factors of HSC biology are beginning to be unveiled, but are still largely under-explored. Recently, the zebrafish has emerged as a powerful genetic model to study vertebrate hematopoiesis. Establishing HCT in zebrafish will allow scientists to utilize the large-scale genetic and chemical screening methodologies available in zebrafish to reveal novel mechanisms underlying HSC regulation. In this article, we demonstrate a method to perform HCT in adult zebrafish. We show the dissection and preparation of zebrafish whole kidney marrow, the site of adult hematopoiesis in the zebrafish, and the introduction of these donor cells into the circulation of irradiated recipient fish via intracardiac injection. Additionally, we describe the post-transplant care of fish in an "ICU" to increase their long-term health. In general, gentle care of the fish before, during, and after the transplant is critical to increase the number of fish that will survive more than one month following the procedure, which is essential for assessment of long term (&lt;3 month) engraftment. The experimental data used to establish this protocol will be published elsewhere. The establishment of this protocol will allow for the merger of large-scale zebrafish genetics and transplant biology.

In this article, we demonstrate a method to perform HCT in adult zebrafish.

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an ...

Retro-orbital Injection in Adult Zebrafish

Drug treatment of whole animals is an essential tool in any model system for pharmacological and chemical genetic studies. Intravenous (IV) injection is often the most effective and noninvasive form of delivery of an agent of interest. In the zebrafish (Danio rerio), IV injection of drugs has long been a challenge because of the small vessel diameter. This has also proved a significant hurdle for the injection of cells during hematopoeitic stem cell transplantation. Historically, injections into the bloodstream were done directly through the heart. However, this intra-cardiac procedure has a very high mortality rate as the heart is often punctured during injection leaving the fish prone to infection, massive blood loss or fatal organ damage. Drawing on our experience with the mouse, we have developed a new injection procedure in the zebrafish in which the injection site is behind the eye and into the retro-orbital venous sinus. This retro-orbital (RO) injection technique has been successfully employed in both the injection of drugs in the adult fish as well as transplantation of whole kidney marrow cells. RO injection has a much lower mortality rate than traditional intra-cardiac injection. Fish that are injected retro-orbitally tend to bleed less following injection and are at a much lower risk of injury to a major organ like the heart. Further, when performed properly, injected cells and/or drugs quickly enter the bloodstream allowing compounds to exert their effect on the whole fish and kidney cells to easily home to their niche. Thus, this new injection technique minimizes mortality while allowing efficient delivery of material into the bloodstream of adult fish. Here we exemplify this technique by retro-orbital injection of Tg(globin:GFP) cells into adult casper fish as well as injection of a red fluorescent dye (dextran, Texas Red ) into adult casper fish. We then visualize successful injections by whole animal fluorescence microscopy.

Here we show how to do retro-orbital injection in adult zebrafish.

Drug treatment of whole animals is an essential tool in any model system for pharmacological and chemical genetic studies. Intravenous (IV) injection is ...

Intraperitoneal Injection into Adult Zebrafish

A convenient method for chemically treating zebrafish is to introduce the reagent into the tank water, where it will be taken up by the fish. However, this method makes it difficult to know how much reagent is absorbed or taken up per fish. Some experimental questions, particularly those related to metabolic studies, may be better addressed by delivering a defined quantity to each fish, based on weight. Here we present a method for intraperitoneal (IP) injection into adult zebrafish. Injection is into the abdominal cavity, posterior to the pelvic girdle. This procedure is adapted from veterinary methods used for larger fish. It is safe, as we have observed zero mortality. Additionally, we have seen bleeding at the injection site in only 5 out of 127 injections, and in each of those cases the bleeding was brief, lasting several seconds, and the quantity of blood lost was small. Success with this procedure requires gentle handling of the fish through several steps including fasting, weighing, anesthetizing, injection, and recovery. Precautions are required to minimize stress throughout the procedure. Our precautions include using a small injection volume and a 35G needle. We use Cortland salt solution as the vehicle, which is osmotically balanced for freshwater fish. Aeration of the gills is maintained during the injection procedure by first bringing the fish into a surgical plane of anesthesia, which allows slow operculum movements, and second, by holding the fish in a trough within a water-saturated sponge during the injection itself. We demonstrate the utility of IP injection by injecting glucose and monitoring the rise in blood glucose level and its subsequent return to normal. As stress is known to increase blood glucose in teleost fish, we compare blood glucose levels in vehicle-injected and non-injected adults and show that the procedure does not cause a significant rise in blood glucose.

We demonstrate intraperitoneal injection into adult zebrafish. We use a 10 &mu;l NanoFil microsyringe controlled by a Micro4 controller and UltraMicroPump III. This demonstration includes the use of cold water as an anesthetic.

A convenient method for chemically treating zebrafish is to introduce the reagent into the tank water, where it will be taken up by the fish. However, ...

Transplantation of Cells Directly into the Kidney of Adult Zebrafish

Regenerative medicine based on the transplantation of stem or progenitor cells into damaged tissues has the potential to treat a wide range of chronic diseases1. However, most organs are not easily accessible, necessitating the need to develop surgical methods to gain access to these structures. In this video article, we describe a method for transplanting cells directly into the kidney of adult zebrafish, a popular model to study regeneration and disease2. Recipient fish are pre-conditioned by irradiation to suppress the immune rejection of the injected cells3. We demonstrate how the head kidney can be exposed by a lateral incision in the flank of the fish, followed by the injection of cells directly in to the organ. Using fluorescently labeled whole kidney marrow cells comprising a mixed population of renal and hematopoietic precursors, we show that nephron progenitors can engraft and differentiate into new renal tissue - the gold standard of any cell-based regenerative therapy. This technique can be adapted to deliver purified stem or progenitor cells and/or small molecules to the kidney as well as other internal organs and further enhances the zebrafish as a versatile model to study regenerative medicine.

Cell transplantation is an essential technique for studying tissue regeneration and for developing cell-based therapies of disease. We demonstrate here a microsurgical technique that permits the transplantation of genetically labeled cells directly into the kidney of adult zebrafish fish.

Regenerative medicine based on the transplantation of stem or progenitor cells into damaged tissues has the potential to treat a wide range of chronic ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...

Blood Collection for Biochemical Analysis in Adult Zebrafish

The zebrafish has been used as an animal model for studies of several human diseases. It can serve as a powerful preclinical platform for studies of molecular events and therapeutic strategies as well as for evaluating the physiological mechanisms of some pathologies1. 
There are relatively few publications related to adult zebrafish physiology of organs and systems2, which may lead researchers to infer that the basic techniques needed to allow the exploration of zebrafish systems are lacking3. Hematologic biochemical values of zebrafish were first reported in 2003 by Murtha and colleagues4 who employed a blood collection technique first described by Jagadeeswaran and colleagues in 1999. Briefly, blood was collected via a micropipette tip through a lateral incision, approximately 0.3 cm in length, in the region of the dorsal aorta5. Because of the minute dimensions involved, this is a high-precision technique requiring a highly skilled practitioner. The same technique was used by the same group in another publication in that same year6. In 2010, Eames and colleagues assessed whole blood glucose levels in zebrafish7. They gained access to the blood by performing decapitations with scissors and then inserting a heparinized microcapillary collection tube into the pectoral articulation. They mention difficulties with hemolysis that were solved with an appropriate storage temperature based on the work Kilpatrick et al.8. When attempting to use Jagadeeswaran's technique in our laboratory, we found that it was difficult to make the incision in precisely the right place as not to allow a significant amount of blood to be lost before collection could be started. 
Recently, Gupta et al.9 described how to dissect adult zebrafish organs, Kinkle et al.10 described how to perform intraperitoneal injections, and Pugach et al.11 described how to perform retro-orbital injections. However, more work is needed to more fully explore basic techniques for research in zebrafish.
The small size of zebrafish presents challenges for researchers using it as an experimental model. Furthermore, given this smallness of scale, it is important that simple techniques are developed to enable researchers to explore the advantages of the zebrafish model.

This paper presents a technique for collection of blood from the dorsal aorta of Zebrafish. It also provides instructions for obtaining serum for use in biochemical analyses, such as tests for determining cholesterol and triglyceride levels.

The zebrafish has been used as an animal model for studies of several human diseases. It can serve as a powerful preclinical platform for studies of ...

Gavaging Adult Zebrafish

The zebrafish has become an important in vivo model in biomedical research. Effective methods must be developed and utilized to deliver compounds or agents in solutions for scientific research. Current methods for administering compounds orally to adult zebrafish are inaccurate due to variability in voluntary consumption by the fish. A gavage procedure was developed to deliver precise quantities of infectious agents to zebrafish for study in biomedical research. Adult zebrafish over 6 months of age were anesthetized with 150 mg/L of buffered MS-222 and gavaged with 5 &mu;l of solution using flexible catheter implantation tubing attached to a cut 22-G needle tip. The flexible tubing was lowered into the oral cavity of the zebrafish until the tip of the tubing extended past the gills (approximately 1 cm). The solution was then injected slowly into the intestinal tract. This method was effective 88% of the time, with fish recovering uneventfully. This procedure is also efficient as one person can gavage 20-30 fish in one hour. This method can be used to precisely administer agents for infectious diseases studies, or studies of other compounds in adult zebrafish.

The increasing use of zebrafish as an animal model requires the development of effective methods for the delivery of known quantities of compounds and solutions. The gavage procedure described below allows for the oral delivery of precise volumes of solution reliably, safely and efficiently to adult zebrafish.

The zebrafish has become an important in vivo model in biomedical research. Effective methods must be developed and utilized to deliver compounds or ...

An Assay for Lateral Line Regeneration in Adult Zebrafish

Due to the clinical importance of hearing and balance disorders in man, model organisms such as the zebrafish have been used to study lateral line development and regeneration. The zebrafish is particularly attractive for such studies because of its rapid development time and its high regenerative capacity. To date, zebrafish studies of lateral line regeneration have mainly utilized fish of the embryonic and larval stages because of the lower number of neuromasts at these stages. This has made quantitative analysis of lateral line regeneration/and or development easier in the earlier developmental stages. Because many zebrafish models of neurological and non-neurological diseases are studied in the adult fish and not in the embryo/larvae, we focused on developing a quantitative lateral line regenerative assay in adult zebrafish so that an assay was available that could be applied to current adult zebrafish disease models. Building on previous studies by Van Trump et al.17&#160;that described procedures for ablation of hair cells in adult Mexican blind cave fish and zebrafish (Danio rerio), our assay was designed to allow quantitative comparison between control and experimental groups. This was accomplished by developing a regenerative neuromast standard curve based on the percent of neuromast reappearance over a 24 hr time period following gentamicin-induced necrosis of hair cells in a defined region of the lateral line. The assay was also designed to allow extension of the analysis to the individual hair cell level when a higher level of resolution is required.

Because many zebrafish models of neurological and non-neurological diseases are studied in the adult fish rather than the embryo/larvae, we developed a quantitative lateral line regenerative assay that can be applied to adult zebrafish disease models. The assay involved resolution at the 1) neuromast and 2) individual hair cell levels.

Due to the clinical importance of hearing and balance disorders in man, model organisms such as the zebrafish have been used to study lateral line ...

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and purification of RNA, DNA, and proteins. All of these applications demand the rapid recovery of significant numbers of zebrafish hearts to avoid gene regulatory, metabolic, and other changes that begin after death. Adult zebrafish hearts are also required for studying heart structure for a variety of mutants and for studying heart regeneration. However, the traditional zebrafish heart dissection is slow and difficult and requires specialized tools, making large-scale dissection of adult zebrafish hearts tedious. Traditional methods also harbor the risk of damaging the heart during the dissection. Here, we describe a method for dissection of adult zebrafish hearts that is fast, reproducible, and preserves heart architecture. Furthermore, this method does not require specialized tools, is painless for the zebrafish, can be performed on fresh or fixed specimens, and can be performed on zebrafish as young as one month old. The approach described expands the use of adult zebrafish for cardiovascular research.

Use of zebrafish for cardiovascular research is expanding towards research on adult hearts. For these applications, quick and simple isolation of cardiac tissues is key to avoid post-mortem changes and to obtain an adequate number of samples. Here, we describe a fast and reproducible method for dissecting adult zebrafish hearts.

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and ...