This document summarizes a high school student's internship researching zebrafish stem cells. The student focused on locating stem cells in the zebrafish telencephalon and brainstem through immunohistochemistry techniques. Results showed nestin-positive stem cells in the telencephalon, confirming previous findings. Additionally, an experiment conducted by the student found nestin-positive cells in the brainstem, supporting the hypothesis that stem cells there may promote central nervous system regeneration in zebrafish. Through this internship, the student gained skills in zebrafish care, dissection, sectioning, immunostaining, microscopy, and independent experimentation.

1. Zebrafish Stem Cell Project
Christina Chyr
High School Research Internship done at St. Thomas University
Mentors: Dr. Jeffrey Plunkett, Dr. Alexis Tapanes-Castillo
08/2012 – 05/2013
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Figure 1. Zebrafish (Danio rerio)

2. Table of Contents:
1. Introduction: 3
2. Methods: 4-8
a. General Fish Care 4
b. Dissection Process 5
c. Brain Sectioning Process 6
d. Immunostaining 7-8
e. Microscopy 9
3. Results: 10-17
a. Stem Cell Project: Telencephalon 10-12
b. Stem Cell Project: Brainstem 13-17
4. Discussion: 18
5. References: 19
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3. 1. Introduction
The capability of nervous systems to regenerate is exhibited in certain species. Mammals can only
regenerate their peripheral nervous system (PNS), but not their central nervous system (CNS). CSPGs
(chondroitin sulfate proteglycans), which are found in almost every animal, inhibit neuroregeneration. They
are a family of proteins that have a protein core with side chains of glycosaminoglycan (sugar) molecules.
However, certain species, such as zebrafish (Danio rerio), are able to overcome this barrier.
To explore the biology of CNS regeneration, adult zebrafish were used. Throughout my internship at
St. Thomas University, I focused on stem cells. Using molecular and microscopy techniques, I was able to
study these cells in zebrafish. I also assisted with general fish care, referred to as animal husbandry, which is
essential for research. There is still much to be learned about why zebrafish have the capability of CNS
regeneration, but mammals do not. I hypothesize that the large number of stem cells located in the adult brain
is a major reason why these fish could undergo CNS regeneration. The results of this and further study could
contribute to our knowledge of the nervous system and could potentially be applied to humans in the future.
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4. 2. Methods
a. General Fish Care:
In order to provide the zebrafish with a good quality of life, certain procedures were taken while
housing, breeding, and dissecting fish. The fish were cared for following the procedures approved and
regularly monitored by the St Thomas University Institutional Animal Care and Use Committee, the United
States Department of Defense, and a local veterinarian. At specific and exclusive time intervals, the zebrafish
were monitored and fed; and their tanks washed.
• IACUC Training:
o Prior to using laboratory animals for research, all participants must go through training
procedures enforced by the Institutional Animal Care and Use Committee (IACUC).
o To prove my qualifications, I had to read and comprehend passages regarding the different
protocols of handling laboratory animals. Then, I also had to pass a final exam.
• Protocol for Cleaning Tanks:
o Change water regularly with conditioned aquarium water.
 Aquarium water recipe:
• Add a half cap of Prime Water conditioner first.
• Then, fill water convoy with half distilled water and half tap water.
o Use clean paper towels to wipe down algae on the inside of any tank the fish will live in.
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5. b. Dissection Process:
To obtain brains used for research purposes, fish are anesthetized and then decapitated. Heads were placed in
a chemical fixative (4% paraformaldehyde) in order to preserve cells. After a two-day incubation in fixative,
heads were then rinsed three times in phosphate buffered saline (PBS) and dissected.
• Formula for one liter of PBS:
o 8 g of NaCl
o 0.2 g KCl
o 1.44 g Na2HPO4
o 0.24 g KH2PO4
o 800 mL of distilled water
 pH to 7.4, then add remaining distilled water to a liter.
1. Fill Petri dish with enough PBS to just immerse the brain.
2. Using a pair of forceps in each hand, hold the head of the fish keeping the underside against the
bottom of the Petri dish.
3. With the dominant hand, slide forceps under the roof of fish’s jaw and cut apart the roof of the skull.
4. Once the superior part of the brain is exposed, discard the sides of the skull by peeling and cutting
with forceps.
5. Using the same forceps, scoop the brain out without damaging the brain’s tissues.
Following the described procedures, I was able to dissect my own zebrafish brains for my experiment. As
mentioned, dissection is a very tedious process that requires great caution, attentiveness, patience, and
practice. Because of the fish’s small body size and slippery exterior, keeping a tight hold on the fish and using
a microscope to get a closer view of cuts made on the fish were difficult to adapt to. However, using
extremely steady hands, I was able to successfully dissect out a whole fish brain.
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6. c. Brain Sectioning Process:
During this procedure, dissected brains are cut into thin 14 µm layers, called sections.
1. Place dissected fish brains in a 30% sucrose/PBS solution overnight.
2. Set cryostat rotary micotome to proper conditions and -14 °C.
3. Prepare chuck with an initial layer of OCT embedding media and let it almost freeze.
4. Place brain onto a chuck in the proper position according to the sections.
• For example, to cut coronal sections, place the brainstem away from the blade and anterior part
of the brain facing the blade.
5. Add an additional layer of OCT to embed the specimen and make the frozen medium uniform.
Avoid bubbles.
6. Cut specimen block into a rectangular shape keeping in mind not to damage the brain.
7. Place chuck with block on the holder.
8. Arrange blade until it touches the chuck.
9. Section through the block until you reach your area of interest in the brain.
10. Prepare and properly label slides to place sections with the proper labels.
11. Using a thin paintbrush, brush brain sections of interest as they are being sliced onto a microscopy
slide. The section will melt easily onto the slide upon touch.
12. Continue until the entire brain or desired area has been sectioned.
To create a perfectly aligned brain in the chuck is not an easy task. It requires practice and quick and
steady hands. Once the positioning is obtained, learning to place the sections onto the slide while being
mindful of the blade and space available on the slide was not difficult- though the first few slides were not
successful.
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7. d. Immunostaining:
To be able to observe cells under the microscope, sections must be washed and have both a primary and
secondary antibody added to. Primary antibodies will attach to specific molecules in cells and secondary
antibodies carry a fluorescent chemical tag that binds to its specific primary antibody. A control slide, in
which the secondary antibody is added to the specimen, but the primary is omitted, was also used to prove
that the fluorescence seen on the slide is due to the primary antibody binding to its specific protein and not
simply a result of background staining caused by the secondary antibody binding non-specifically to tissue.
1. Pre-blocking:
a. Rinse slides carrying brain sections three times in PBS.
2. Blocking:
a. Prepare blocking solution: (10 mL)
i. Recipe:
1. 1.5% Normal Goat Serum 150 μL
2. 0.3% Triton X 30 μL
3. PBS 9.82 mL
ii. Mix for 30 to 60 seconds ad keep in ice. (Only good for 24 hours.)
b. Prepare a humidified slide chamber:
i. Place paper towel in plastic container.
ii. Place white rack on top of paper towel.
iii. Wet paper towel lightly with distilled water to retain humidity within the
container.
iv. Place slides on rack and keep them separated from each other.
c. Add 500 μL of blocking solution to slide. (This should be enough to fill up the whole slide
thinly.)
d. Close container and do not disturb for an hour at room temperature.
3. Primary Antibody and Control:
a. Cut strips of parafilm to the same size as the slides.
b. Dilute primary antibody with blocking solution. (see Table 1 for concentrations.)
c. Choose which slides will be control group and which will be experimental group.
i. For primary antibody slides:
1. Remove blocking solution from slide that will have the primary antibody
added by tilting.
2. Add 250 μL of primary antibody.
3. Cover slide with parafilm.
a. Do not move parafilm once it is added!
ii. For control slides:
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8. 1. Keep blocking solution on and cover with parafilm.
a. Do not move parafilm once it is added!
d. Incubate at room temperature for four hours.
e. Transfer to 4 ﾟ C and incubate for two nights.
4. Secondary Antibody:
a. Use forceps to remove parafilm from all slides.
b. Remove blocking solution and primary antibody by tilting.
c. Put slides into coplin jars filled with PBS, one by one.
d. Let the slides sit for 10 minutes.
e. Take out slides, pour out PBS, and refill.
f. Repeat two more times for 10 minutes. That is a total of three 10 minute washings.
g. Dilute secondary antibody with blocking solution as specified in Table 1 and keep in ice.
h. Put slides back on rack in the humidified chamber.
i. Add 250 μL of secondary antibody and cover with parafilm one by one.
i. Keep slides in the dark to prevent the fluorescent secondary antibody from
photobleaching.
j. Incubate for an hour at room temperature.
k. Wash slides with PBS again five times for ten minutes each.
5. Mounting:
a. Remove one slide from coplin jar and clean its bottom using Kim wipes.
i. Leave residual PBS over sections.
b. Add 30 μL of DAPI containing vectorshield fluorescent mounting media and cover it with
cover glass.
i. Make sure there are no bubbles.
ii. DO NOT MOVE.
c. Store slides in 4 ﾟ C.
Table 1. Antibody Concentration
Antibody: Concentrations:
Nestin-rabbit 1/250
Tubulin-mouse 1/200
Goat-anti-rabbit-Alexa 488 1/250
Goat-anti-mouse-Alexa 594 1/250
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9. e. Microscopy
In order to visualize slides, I used the Axioskop2 plus microscope through 10x to 40x objective lenses. To
obtain image micrographs, I used Axiovision, an imaging software. I took pictures on different fluorescent
channels and set scale based on objective lens used. Finally, I processed images with Adobe Photoshop, a
photo editor software.
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10. Figure 2. Micrographs
Micrograph of an adult telencephalon. The same section was
imaged using two fluorescents.
Top Panel: DAPI (blue) labeling nuclei
Middle Panel: Anti-nestin antibody (green) labeling stem cells
Bottom Panel: Merged Image showing DAPI and nestin labeling
of the same section.
3. Results Section
a. Stem Cell Project: Telencephalon:
The purpose of this project is to work towards
understanding the role of stem cells in zebrafish CNS
regeneration. My main focus was to locate stem cells.
Stem cells are cells that can replicate indefinitely and
differentiate into multiple specialized cell types. In the
case of damage to the nervous system, the stem cell can
differentiate into a functioning neuron.
Stem cells express nestin protein. To locate
nestin, I used a primary antibody protein called anti-
nestin, which is made in rabbit, to recognize nestin
protein. This was followed by binding a secondary
antibody protein called anti-rabbit Alexa 488, which
glows green, to the primary antibody. The result was
green labeled cells.
I also labeled cells with a dye called DAPI,
which is fluorescent under UV light to observe the cell
nuclei present. Using a laser fluorescence microscope,
images called micrographs were taken of brain regions
with nestin cells.
Figure 2 is a micrograph taken from a coronal
section of an adult zebrafish brain, specifically around
cross section 92, the telencephalon (figure 3). Two
different fluorescent channels: UV (top panel) and
green (middle panel) were imaged and merged in the
bottom panel. In the top panel of Figure 2, the DAPI
dye was able to indicate the nuclei present. The second
panel indicated present nestin. In the bottom panel, cells
labeled with both nestin and DAPI are observed.
The micrograph in Figure 4 depicts another
brain section of the telencephalon with nestin positive
cells. This corresponds to a section near cross section
92 (Figure 3).
In Figures 2 and 4, green represents nestin
labeled cells, which we hypothesize are stem cells. Data
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11. from numerous labs indicates a stem cell population in the telencephalon. My results support these findings.
Furthermore, they served as a positive control that shows my capability to perform nestin
immunohistochemistry.
Figure 3. Cross section 92 diagram found in
Neuroanatomy of the Zebrafish Brain: A Topological Atlas
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12. Figure 4. Micrographs
Merged micrograph of an adult telencephalon.
DAPI (blue) labels nuclei and anti-nestin antibody (green) labels stem cells.
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Figure 4. Micrograph (DAPI and nestin)

13. b. Stem Cell Project: Experiment
The Plunkett lab is specifically interested in spinal cord regeneration. Cells found in the caudal-most
(back) region of the brain, called the brainstem, project axons into the spinal cord. Based on the knowledge
from several lab’s published data that show nestin expressions in cells found in the telencephalon, I
conducted an experiment to answer the question “Are there nestin positive cells in the brainstem?”. I
hypothesized that the answer would be yes and that these stem cells promote CNS regeneration. Referencing
to the first experiment, I reproduced my own through the use of cross sections of the brainstem. To test this
hypothesis, I wanted to reproduce the experiment identifying nestin-positive cells in the telencephalon, but
now in the brainstem. I also wanted to perform the entire histological procedure on my own.
I began this experiment by independently dissecting and sectioning a brain. I performed
immunostaining on my best sections using an anti-tubulin antibody. I used this antibody as a practice
experiment before using the nestin antibody again. Tubulin labels all cells and helps one visualize the
morphology of the entire section. If sample sections are deemed to be of high quality, then adjacent sample
sections can be utilized for the nestin stains.
The general vicinity of my sections was located to be approximately around cross section 260 in the
Atlas (see Figure 5). For my primary antibody, I used mouse anti-tubulin. For my secondary antibody, the
one that expresses the color, I used goat-anti-mouse-Alexa (GAM) 594, which is fluorescent red. I prepared
the control and experimental slides. By doing so, I was able to compare what is truly expressed by the
primary antibody and what was background staining through the four different channels: DAPI (blue), “none”
(green), 594 (red), and merged. The control group should not have any fluorescent signaling and what is seen
is considered false or background staining. Within the experimental group, what is seen in the “none” (green)
and red channel is also considered false signaling.
When the control and experimented sections were compared, there was a notable difference. The
only similarity between all three slides (one control and two experimental) was in the channels “none” and
“DAPI”. Anything shown in the “none” channel, but also seen in the red channel, indicated background
staining and not real primary antibody labeling. It is normal to see some background labels in experiments,
but without a control there will be nothing to compare the experimented slides to.
Figure 6 shows the control micrographs which contained no primary antibody. Notice there is a lack
of bright red stains in the box labeled “594”, which is also called the red channel. This means there is no
tubulin immunoreactivity. Since no green secondary antibody was used, any signal seen in the green and red
channels indicated background staining.
In Figures 7 and 8, however, there were bright and noticeable red stains that surrounded the red
channel (594). The vibrant red color shows where the primary antibody successfully binded to tubulin.
However, these two sections were not perfectly stained, because only the more external tissue seemed to stain
and not the center cells in the brainstem of the zebrafish. The red labels shown in Figures 7 and 8 were
definitely not effects of background staining, but true primary antibody binding to molecules in the cells.
This experiment confirmed that I could perform a histology experiment on my own.
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14. Figure 5. Cross section 260 diagram found in
Neuroanatomy of the Zebrafish Brain: A Topological Atlas
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15. Figure 6. Control micrographs
Top Left: DAPI (blue) labeling nuclei
Top Right: None (green)
Bottom Left: Absence of Anti-nestin antibody (red) labeling stem cells
Bottom Right: Merged image showing DAPI, none, and tubulin (which was absent)
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16. Figure 7. Micrographs
Top Left: DAPI (blue) labeling nuclei
Top Right: None (green)
Bottom Left: Anti-nestin antibody (red) labeling stem cells
Bottom Right: Merged image showing DAPI, none, and tubulin
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17. Figure 8. Micrographs
Top Left: DAPI (blue) labeling nuclei
Top Right: None (green)
Bottom Left: Anti-nestin antibody (red) labeling stem cells
Bottom Right: Merged image showing DAPI, none, and tubulin
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18. 4. Discussion
Throughout my duration at St. Thomas University, I learned to perform histological experiments. I
conducted an experiment that labeled nestin-positive cells in the telencephalon. I also performed an entire
experiment on my own labeling tubulin-positive cells in the brainstem. I did not have sufficient time,
however, to perform my final experiment which was to stain brainstem sections with the anti-nestin antibody.
Nevertheless, university students in the lab have identified nestin-positive cells in the brainstem. This finding
is novel. It has never before been documented in the field.
In addition to viewing cells in vivo, I also participated in the dissociation and culture of brain cells in
a Petri dish. This technique can be used to study stem cells under controlled conditions.
While the knowledge is currently limited on the brain and its activities, it is quite possible for the
future studies to answer the mystery of why zebrafish, unlike mammals, can regenerate their central nervous
system. This can lead to therapies that can help human patients with neurological trauma.
From basic laboratory safeties to conducting immunohistochemistry experiments, my knowledge
and interest in the science has grown. Since entering the Plunkett lab in August, I have learned many skills
that will undoubtedly assist in my future goals of becoming a doctor. While my specific field of interest is
undecided, this internship has also allowed me to explore the depths of neuroscience. The choice to conduct
research on the brain is like wading into the unknown fathoms of the ocean. However, it brings great rewards,
such as knowledge and advancement, to the world.
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19. 5. References
Mario F. Wullimann, Barbara Rupp, Heinrich Reichert. Neuroanatomy of the Zebrafish Brain: A Topological
Atlas. Boston: Deutsche Bibliothek, 1996. Print.
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