Name: methods for the study of the zebrafish maxillary barbel
Uploaded: 2009-11-23T16:00:00
Duration: 9 min 30 s
Description: Watch this Scientific Journal Video about Methods for the Study of the Zebrafish Maxillary Barbel at JoVE.com

These methods were developed by E.E.L. during a research leave in the lab of J.T. Support was provided by the DePaul University Research Council and an NIH/NIDCR R01 grant to J.T. (DE016678). We gratefully acknowledge the efforts of Caroline Hunter, animal care technician in the CMRC zebrafish core facility and Paulina Pawluczuk, our undergraduate laboratory assistant.

Zebrafish husbandry
Zebrafish are housed and reared according to standard methods 1. Past the late larval period, the developmental stage of a zebrafish can be measured by its standard length (SL), or the straight line distance in millimeters from the anteriormost point of the upper lip to the base of the caudal fin (posteriormost hypural plate)2. At approximately one month post fertilization (10-12 mm SL), the nasal and maxillary barbels appear as transparent epithelial buds near the olfactory pits and on the posterior corners of the maxillae, respectively. As the zebrafish matures, the barbels extend into narrow, whisker-like appendages. Because the maxillary barbel is larger (2-3 mm) and easier to manipulate, our protocol applies only to this particular appendage; similar techniques, however, could be adapted to the smaller nasal barbel.
Barbel Clipping
<ol>
 <li>Anaesthetize adult zebrafish at the desired stage (e.g., 1.5-2.5 cm SL, ~3-6 months post-fertilization) by immersion in 0.015% MS-222 (ethyl 3-aminobenzoate methanesulfonate) buffered to pH 7.0 in system water. Observe until swim movements stop and gill ventilation becomes slow and regular. Depending on fish size, complete anesthesia takes approximately 2-5 minutes.</li>
 <li>Using an aquarium fishnet, transfer 1-3 fish at a time to a folded piece of wet paper towel in a Petri dish. Using a blunt spatula, orient the fish so they are parallel to each other and left lateral side up. Fold part of the wet paper towel over the caudal part of the fish, covering the entire body up to the operculum (gill covers) to keep the skin and gills moist during the procedure.</li>
 <li>View the Petri plate under the stereomicroscope, using low-angle incident light to illuminate the head region. Focus on the base of the left maxillary barbel, which protrudes from the posterior ventral corner of the maxilla (Fig. 1). <img src="/files/ftp_upload/1558/1558fig1.jpg" alt="Figure 1" /> Figure 1.</li>
 <li>Tightly grasp the shaft of the barbel slightly distal to the edge of the maxilla. Elevate the barbel shaft and insert the jaws of a fine-tipped spring scissors just proximal to the forceps. The scissors can be touched to the forceps for stability. Slide the jaws of the scissors along the shaft of the barbel until the cutting edge approaches the edge of the maxilla. Close the scissors to cut through the barbel shaft. The amputated barbel, still gripped in the forceps, can be removed for fixation or further observation. </li>
 <li>Immediately transfer the fish to a small tank of clean system water (~500 mL) containing one drop of methylene blue to control superficial fungal infection. Allow the fish to recover overnight. The next morning, return the fish to the rearing system. Barbel regeneration is maximal after 2 weeks.</li>
</ol>
Agar Embedding of Matched Barbel Pairs
<ol>
 <li>Collect fish by appropriate euthanasia technique and fix in a 50 mL tube of 4% paraformaldehyde-phosphate buffered saline (PBS) with agitation at 4&deg;C overnight. The fixative volume should be at least ten times the volume of the tissue. After fixation, dispose of paraformaldehyde waste in an approved container and rinse the tissue thoroughly in several changes of PBS. </li>
 <li>Prepare in a 250 mL glass flask 150 mL of 2% agarose (Tm ~37&deg;C) in distilled water. Heat with a microwave or hot plate, agitating periodically until completely dissolved. Equilibrate the molten solution in a 50-60&deg;C water bath.</li>
 <li>Assemble a standard gel electrophoresis rig by placing a small gasketed gel mold (10 x 10 cm; ~100 mL gel volume) firmly against the sides of the buffer chamber. Add 2-4 small-toothed sample combs (4 x 1.5 mm). </li>
 <li>On a level surface, pour the melted agarose into the mold to just cover the base of the combs, forming very shallow wells. Immediately put the leftover agarose back into the warm water bath; this will be used later to embed the tissue (Steps 9-12). Put a long glass Pasteur pipette into the flask to keep the pipette to the same temperature as the agarose. </li>
 <li>Allow the gel to harden for 20-30 minutes. Remove the combs, then remove the gel mold containing the hardened gel from the buffer chamber. Firmly tape the open sides of the gel mold with laboratory tape so the gel does not slide out. This tape should extend several millimeters over the surface of the agarose so that additional agarose can be added later (Step 13). </li>
 <li>Position the gel mold on the stage of a stereomicroscope. Magnify an empty well and bring the edges into focus. </li>
 <li>Onto the surface of the agarose adjacent to the well, pipette a drop of water or buffer containing the fixed barbel specimens (e.g., regenerate and control). Using bent insect pins, drag the moist specimens into the well and push them to the bottom. Orient the barbel shafts parallel to each other and aligned against opposite long edges of the well. Surface tension will help hold the tissue in position.</li>
 <li>When ready to embed, use a fine pipette tip or the corner of a laboratory tissue to remove excess fluid from the well. Work quickly so as not to let the tissue dry out. </li>
 <li>Using the warmed glass Pasteur pipette, gently pipette fresh molten agarose in and around the specimens to seal the barbel tissue in place. Do not overfill. Before the agarose hardens, make last-minute adjustments with the insect pins. </li>
 <li>Place a numbered paper specimen label next to the well and secure it with a small drop of agarose.</li>
 <li>Repeat steps 6 through 10 for each well of specimens. </li>
 <li>If desired for image calibration, insert a piece of paper with a printed micrometer scale (e.g., <a href="http://incompetech.com/graphpaper/multiwidth/">http://incompetech.com/graphpaper/multiwidth/</a>) into the bottom of an empty well. Flatten the paper to the bottom of the well and cover it with warm agarose. </li>
 <li>After all of the specimens are embedded, the surface of the gel will be irregular, with hardened drops of agarose. Place the gel on a flat surface and gently pour in additional melted agarose until the upper surface is uniform. Use the smallest amount of agarose necessary to obtain a smooth surface; too much agarose will obscure transmitted-light photography. Allow this top layer to harden.</li>
 <li>The gel can now be photographed on the stage of a stereomicroscope to record gross morphology for each barbel pair. Image calibration is obtained by photographing the embedded micrometer scale.</li>
 <li>The entire gel can be stored wrapped in wet paper towels and sealed in a plastic bag at 4&deg; for several weeks. </li>
 <li>For further analysis, agar blocks containing matched pair of barbels can be cut out of the gel with a scalpel or razor blade and processed for paraffin histology or cryosectioning by standard methods3,4. Alternatively, individual barbels can be dissected out of the agarose with fine needles, rinsed in water, and processed for other downstream applications (e.g., whole-mount immunohistochemistry or confocal microscopy).</li>
</ol>

<ol>
	<li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+1+General+Methods+for+Zebrafish+Care.">Chapter 1 General Methods for Zebrafish Care.</a> The zebrafish book. A guide for the laboratory use of zebrafish (Danio rerio). , (2000).</li><li>Howe, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Standard+length:+Not+quite+so+standard.">Standard length: Not quite so standard.</a> Fisheries Research (Amsterdam). 56, 1-7 (2002).</li><li>Fischer, A. H., Jacobson, K. A., Rose, J., Zeller, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Paraffin+Embedding+Tissue+Samples+for+Sectioning.">Paraffin Embedding Tissue Samples for Sectioning.</a> Cold Spring Harbor Protocols. 6, (2008).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ch+8.3+Agar+Embedding+for+Cryostat+Sectioning+of+Embryos+or+Larvae.">Ch 8.3 Agar Embedding for Cryostat Sectioning of Embryos or Larvae.</a> The zebrafish book. A guide for the laboratory use of zebrafish (Danio rerio). , (2000).</li><li>Hansen, A., Reutter, K., Zeiske, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Taste+bud+development+in+the+zebrafish,+Danio+rerio.">Taste bud development in the zebrafish, Danio rerio.</a> Dev Dyn. 223, 483-496 (2002).</li></ol>

The zebrafish maxillary barbel is an underutilized tissue system for studying the growth, maintenance, and regeneration of several cell types in zebrafish. Although the barbel appendage has no human analog, the cell types it contains are highly conserved, making it possible to study skin, glands, melanocytes, circulatory vessels and nerves in an optically clear and anatomically simple cylindrical structure. Similar to the well-studied caudal fin, barbel tissue can be induced to regenerate by amputation. Using the border of the maxilla as an anatomical landmark, the amputation plane can be placed precisely, facilitating the measurement of barbel regrowth. For an experienced operator, each surgery takes only a few seconds. Recovery is rapid, and we have so far detected no short or long-term effects on fish behavior. Zebrafish with one maxillary barbel swim, eat and breed as effectively as non-surgical controls, and have comparable survival to the time of tissue collection, up to 6 months after surgery. The physiological impact of this surgery predicted to be minimal because 1) the extraoral taste buds carried on the barbel are also found on many other parts of the fish epithelium, including the lips, cheeks and head5, and 2) differentiating taste buds appear on the regenerating barbel within 72 hours (LeClair et al., unpublished data.) This makes the maxillary barbel a minimally invasive system for studying wound healing, revascularization, and reinnervation within the context of an adult vertebrate. 
After surgical induction of regeneration, barbels can be collected at intervals for morphometric measurement and/or microscopic analysis of fixed tissue. Conveniently, the maxillary barbel is approximately the length (2-3 mm) and diameter (100-200 mm) of a zebrafish embryo, facilitating the application of many standard protocols, including paraffin histology, cryosectioning, whole-mount immunohistochemistry, and in situ hybridization. Taken together, these features make the maxillary barbel a highly feasible in vivo model for studying tissue repair and regeneration.

Microscope Equipment
A stereo dissecting microscope with transmitted and incident fiber-optic illumination is required for barbel clipping and agar embedding.
Barbel Clipping
<ul>
<li>Fish system water</li>
<li>2 zebrafish crossing tanks </li>
<li>MS-222 (ethyl 3-aminobenzoate methanesulfonate; Sigma Chemical #E10521-10G) </li>
<li>sterile 120-mm Petri plate</li>
<li>wet paper towels</li>
<li>blunt metal spatula </li>
<li>Dumont #55 Bio Inox Forceps (Fine Science Tools #11255-20)</li>
<li>Mini-Vannas spring scissors (Fine Science Tools #15000-00)</li>
<li>methylene 	blue antifungal agent (Drs. Foster &amp; Smith #CD-905781)</li>
<li>small 	aquarium fishnet</li>
</ul>
Tissue collection and agar embedding
<ul>
<li>4% paraformaldehyde in 1x phosphate-buffered saline (PF-PBS), aliquoted into 50 mL conical tubes and stored frozen at -20&deg;C until use</li>
<li>1x phosphate-buffered saline (pH 7.27-6)</li>
<li>250 mL glass flask</li>
<li>2% agarose in distilled water</li>
<li>standard small agarose gel electrophoresis rig (gel size = ~10 cm square)</li>
<li>small-toothed electrophoresis sample comb (comb size 4 x 1.5 mm)</li>
<li>warm water bath (50-60&deg;C)</li>
<li>laboratory tape</li>
<li>black enameled insect pins, size 000 (Fine Science Tools #26000-25)</li>
<li>1-2 pin holders (Fine Science tools #26016-12)</li>
<li>9-inch glass Pasteur pipette</li>
<li>pipette bulb or pump</li>
<li>laboratory tissue</li>
<li>small paper labels</li>
<li>(optional) printed calibration scale</li>
<li>wet paper towels</li>
<li>zip-closure plastic bag</li>
<li>scalpel/razor blade (to cut out agar blocks)</li>
</ul>

methods for the study of the zebrafish maxillary barbel

Barbels are skin sensory appendages found in fishes, reptiles and amphibians. The zebrafish, Danio rerio, develops two pairs of barbels- a short nasal pair and a longer maxillary pair. Barbel tissue contains cells of ectodermal, mesodermal and neural crest origin, including skin cells, glands, taste buds, melanocytes, circulatory vessels and sensory nerves. Unlike most adult tissue, the maxillary barbel is optically clear, allowing us to visualize the development and maintenance of these tissue types throughout the life cycle. 
This video shows early development of the maxillary barbel (beginning approximately one month post-fertilization) and demonstrates a surgical protocol to induce regeneration in the adult appendage (&gt;3 months post-fertilization). Briefly, the left maxillary barbel of an anesthetized fish is elevated with sterile forceps just distal to the caudal edge of the maxilla. A fine, sterile spring scissors is positioned against the forceps to cut the barbel shaft at this level, establishing an anatomical landmark for the amputation plane. Regenerative growth can be measured with respect to this plane, and in comparison to the contralateral barbel. Barbel tissue regenerates rapidly, reaching maximal regrowth within 2 weeks of injury.
Techniques for analyzing the regenerated barbel include dissecting and embedding matched pairs of barbels (regenerate and control) in the wells of a standard DNA electrophoresis gel. Embedded specimens are conveniently photographed under a stereomicroscope for gross morphology and morphometry, and can be stored for weeks prior to downstream applications such as paraffin histology, cryosectioning, and/or whole mount immunohistochemistry. These methods establish the maxillary barbel as a novel in vivo tissue system for studying the regenerative capacity of multiple cell types within the genetic context of zebrafish.

Watch this Scientific Journal Video about Methods for the Study of the Zebrafish Maxillary Barbel at JoVE.com

Methods for the Study of the Zebrafish Maxillary Barbel

The zebrafish maxillary barbel is an integumentary sense organ containing ectodermal, mesodermal and neural crest derivatives. Importantly, the adult barbel can regenerate after proximal amputation. This video introduces maxillary barbel development and demonstrates a surgical protocol to induce regeneration, followed by collection, embedding and downstream imaging of barbel specimens.

Barbels are skin sensory appendages found in fishes, reptiles and amphibians.  The zebrafish, Danio rerio, develops two pairs of barbels- a short nasal ...

methods-for-the-study-of-the-zebrafish-maxillary-barbel

Research

JoVE Journal

Biology

Testing Visual Sensitivity to the Speed and Direction of Motion in Lizards

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of the visual system provide more empirical evidence for functional applications. Investigation into the sensory system provides information about the sensory capacity, learning and memory ability, and establishes known baseline behaviour in which to gauge deviations (Burghardt, 1977). However, unlike mammalian or avian systems, testing for learning and memory in a reptile species is difficult. Furthermore, using an operant paradigm as a psychophysical measure of sensory ability is likewise as difficult. Historically, reptilian species have responded poorly to conditioning trials because of issues related to motivation, physiology, metabolism, and basic biological characteristics. Here, I demonstrate an operant paradigm used a novel model lizard species, the Jacky dragon (Amphibolurus muricatus) and describe how to test peripheral sensitivity to salient speed and motion characteristics. This method uses an innovative approach to assessing learning and sensory capacity in lizards. I employ the use of random-dot kinematograms (RDKs) to measure sensitivity to speed, and manipulate the level of signal strength by changing the proportion of dots moving in a coherent direction. RDKs do not represent a biologically meaningful stimulus, engages the visual system, and is a classic psychophysical tool used to measure sensitivity in humans and other animals. Here, RDKs are displayed to lizards using three video playback systems. Lizards are to select the direction (left or right) in which they perceive dots to be moving. Selection of the appropriate direction is reinforced by biologically important prey stimuli, simulated by computer-animated invertebrates.

Testing visual sensitivity in lizards using an operant conditioning paradigm that employs video playback of random-dot kinematograms and computer-generated invertebrates.

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of ...

Computer-Generated Animal Model Stimuli

Communication between animals is diverse and complex. Animals may communicate using auditory, seismic, chemosensory, electrical, or visual signals. In particular, understanding the constraints on visual signal design for communication has been of great interest. Traditional methods for investigating animal interactions have used basic observational techniques, staged encounters, or physical manipulation of morphology. Less intrusive methods have tried to simulate conspecifics using crude playback tools, such as mirrors, still images, or models. As technology has become more advanced, video playback has emerged as another tool in which to examine visual communication (Rosenthal, 2000). However, to move one step further, the application of computer-animation now allows researchers to specifically isolate critical components necessary to elicit social responses from conspecifics, and manipulate these features to control interactions. Here, I provide detail on how to create an animation using the Jacky dragon as a model, but this process may be adaptable for other species. In building the animation, I elected to use Lightwave  3D to alter object morphology, add texture, install bones, and provide comparable weight shading that prevents exaggerated movement. The animation is then matched to select motor patterns to replicate critical movement features. Finally, the sequence must rendered into an individual clip for presentation. Although there are other adaptable techniques, this particular method had been demonstrated to be effective in eliciting both conspicuous and social responses in staged interactions.

Computer-generated stimuli using the Jacky dragon as a model.

Communication between animals is diverse and complex. Animals may communicate using auditory, seismic, chemosensory, electrical, or visual signals. In ...

Using the optokinetic response to study visual function of zebrafish

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the visual functions of larval zebrafish1-5. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adult zabrafish. Here, we introduce how to measure the OKR of adult zebrafish with our simple custom-built apparatus using a new protocol which is established in our lab. Both our apparatus and step-by-step procedure of OKR in adult zebrafish are illustrated in this video. In addition, the measurements of the larval OKR, as well as the optomotor response (OMR) test of adult zebrafish, are also demonstrated in this video. This OKR assay of adult zebrafish in our experiment may last for up to 4 hours. Such OKR test applied in adult fish will benefit to visual function investigation more efficiently when the adult fish vision system is manipulated.
Su-Qi Zou and Wu Yin contributed equally to this paper.

Optokinetic response has been widely used to assess the visual functions of larval zebrafish. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adults1-5. Here, we introduce how to measure the OKR of adult zebrafish using a new protocol which is established in our lab.

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the ...

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, ...

Laser Ablation of the Zebrafish Pronephros to Study Renal Epithelial Regeneration

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in renal failure1. AKI can be caused by a number of factors including ischemia, drug-based toxicity, or obstructive injury1. This results in an inability to maintain fluid and electrolyte homeostasis. While AKI has been observed for decades, effective clinical therapies have yet to be developed. Intriguingly, some patients with AKI recover renal functions over time, a mysterious phenomenon that has been only rudimentally characterized1,2. Research using mammalian models of AKI has shown that ischemic or nephrotoxin-injured kidneys experience epithelial cell death in nephron tubules1,2, the functional units of the kidney that are made up of a series of specialized regions (segments) of epithelial cell types3. Within nephrons, epithelial cell death is highest in proximal tubule cells. There is evidence that suggests cell destruction is followed by dedifferentiation, proliferation, and migration of surrounding epithelial cells, which can regenerate the nephron entirely1,2. However, there are many unanswered questions about the mechanisms of renal epithelial regeneration, ranging from the signals that modulate these events to reasons for the wide variation of abilities among humans to regenerate injured kidneys.
The larval zebrafish provides an excellent model to study kidney epithelial regeneration as its pronephric kidney is comprised of nephrons that are conserved with higher vertebrates including mammals4,5. The nephrons of zebrafish larvae can be visualized with fluorescence techniques because of the relative transparency of the young zebrafish6. This provides a unique opportunity to image cell and molecular changes in real-time, in contrast to mammalian models where nephrons are inaccessible because the kidneys are structurally complex systems internalized within the animal. Recent studies have employed the aminoglycoside gentamicin as a toxic causative agent for study of AKI and subsequent renal failure: gentamicin and other antibiotics have been shown to cause AKI in humans, and researchers have formulated methods to use this agent to trigger kidney damage in zebrafish7,8. However, the effects of aminoglycoside toxicity in zebrafish larvae are catastrophic and lethal, which presents a difficulty when studying epithelial regeneration and function over time. Our method presents the use of targeted cell ablation as a novel tool for the study of epithelial injury in zebrafish. Laser ablation gives researchers the ability to induce cell death in a limited population of cells. Varying areas of cells can be targeted based on morphological location, function, or even expression of a particular cellular phenotype. Thus, laser ablation will increase the specificity of what researchers can study, and can be a powerful new approach to shed light on the mechanisms of renal epithelial regeneration. This protocol can be broadly applied to target cell populations in other organs in the zebrafish embryo to study injury and regeneration in any number of contexts of interest.

Acute kidney injury (AKI) in humans is a common clinical problem caused by damage to the epithelial cells that comprise kidney nephrons, and AKI is associated with high mortality rates of 50-70%1. Following epithelial cell destruction, nephrons have a limited ability to regenerate, though the mechanisms and limitations that guide this phenomenon remain poorly understood. In this video article, we describe our technique for targeted laser ablation of kidney nephron cells in the zebrafish embryo kidney, or pronephros. Our new method can be used to complement nephrotoxicity-induced models of AKI and gain a high-resolution understanding of the cell and molecular alterations that are associated with epithelial regeneration in the kidney nephron.

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in ...

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 ...

Do-It-Yourself Device for Recovery of Cryopreserved Samples Accidentally Dropped into Cryogenic Storage Tanks

Liquid nitrogen is colorless, odorless, extremely cold (-196 &deg;C) liquid kept under pressure. It is commonly used as a cryogenic fluid for long term storage of biological materials such as blood, cells and tissues 1,2. The cryogenic nature of liquid nitrogen, while ideal for sample preservation, can cause rapid freezing of live tissues on contact - known as 'cryogenic burn'2, which may lead to severe frostbite in persons closely involved in storage and retrieval of samples from Dewars. Additionally, as liquid nitrogen evaporates it reduces the oxygen concentration in the air and might cause asphyxia, especially in confined spaces2.
In laboratories, biological samples are often stored in cryovials or cryoboxes stacked in stainless steel racks within the Dewar tanks1. These storage racks are provided with a long shaft to prevent boxes from slipping out from the racks and into the bottom of Dewars during routine handling. All too often, however, boxes or vials with precious samples slip out and sink to the bottom of liquid nitrogen filled tank. In such cases, samples could be tediously retrieved after transferring the liquid nitrogen into a spare container or discarding it. The boxes and vials can then be relatively safely recovered from emptied Dewar. However, the cryogenic nature of liquid nitrogen and its expansion rate makes sunken sample retrieval hazardous. It is commonly recommended by Safety Offices that sample retrieval be never carried out by a single person. Another alternative is to use commercially available cool grabbers or tongs to pull out the vials3. However, limited visibility within the dark liquid filled Dewars poses a major limitation in their use.
In this article, we describe the construction of a Cryotolerant DIY retrieval device, which makes sample retrieval from Dewar containing cryogenic fluids both safe and easy.

Here we present a low cost, durable cryotolerant device for sample retrieval from Dewar tanks filled with liquid nitrogen. The ease of construction and modular design of the device makes the process of sample retrieval from cryogenic tanks safe and easy.

Liquid nitrogen is colorless, odorless, extremely cold (-196 &deg;C) liquid kept under pressure. It is commonly used as a cryogenic fluid for long term ...

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 ...

Methods for the Extraction of Endosymbionts from the Whitefly Bemisia tabaci

Bacterial symbionts form an intimate relationship with their hosts and confer advantages to the hosts in most cases. Genomic information is critical to study the functions and evolution of bacterial symbionts in their host. As most symbionts cannot be cultured in vitro, methods to isolate an adequate quantity of bacteria for genome sequencing are very important. In the whitefly Bemisia tabaci, a number of endosymbionts have been identified and are predicted to be of importance in the development and reproduction of the pests through multiple approaches. However, the mechanism underpinning the associations remains largely unknown. The obstacle partially comes from the fact that the endosymbionts in whitefly, mostly restrained in bacteriocytes, are hard to separate from the host cells. Here we report a step-by-step protocol for the identification, extraction and purification of endosymbionts from the whitefly B. tabaci mainly by dissection and filtration. Endosymbiont samples prepared by this method, although still a mixture of different endosymbiont species, are suitable for subsequent genome sequencing and analysis of the possible roles of endosymbionts in B. tabaci. This method may also be used to isolate endosymbionts from other insects.

Methods for the Extraction of Endosymbionts from the Whitefly Bemisia tabaci

Here, we present a protocol to isolate endosymbionts from the whitefly Bemisia tabaci through dissection and filtration. After amplification, the DNA samples are suitable for subsequent sequencing and study of the mutualism between endosymbionts and whitefly.

Bacterial symbionts form an intimate relationship with their hosts and confer advantages to the hosts in most cases. Genomic information is critical to ...