Biology 306
DEVELOPMENTAL
BIOLOGY
LABORATORY MANUAL
FALL 2000
This laboratory manual was written and compiled
by Dr. Frances E. Weaver
Note: In this second edition of the Biology 306 manual, exercises came from a
variety of sources and therefore are in a variety of formats.
Instructions for Laboratory Assignments
Laboratory Notebooks: Each student will keep a detailed, dated laboratory notebook of all
activities in the lab, which will include, but are not limited to, all methodologies and
observations (many of which will take place outside of the scheduled lab period).
Accurate drawings, raw and analyzed data, answers to questions, discussions and suggestions for
future experiments or improvements to the labs must also be included.
Lab notebooks will be collected and graded twice during the "standard" laboratories, and
returned to each student the following week (or sooner as necessary). I expect your lab note book
to improve each time it is collected. A GOOD LAB NOTEBOOK IS PRECISE, DETAILED
AND LEGIBLE!!!!!
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GENES AND DEVELOPMENT MINI PROJECT
Literature review and project report:
General: This paper and the subsequent presentation are, because of budget constraints,
somewhat of a hodgepodge of lab report and research proposal. The literature review should
refer only to the "unknown" gene you have been assigned, although you are going to work with
knowns in the laboratory.
Reports must be typewritten, including tables, table and figure titles and figure legends.
Graphs may be hand drawn but must be done neatly. Begin each section with the title of the
section i.e. Results:
Reports must follow the guidelines below. Points will be taken off if the report does not
adhere to these standards. Plagiarism, collaborative reports (unless authorized) and other
forms of academic fraud are not permitted and will be grounds for FAILURE in the
course. All sources must be referenced.
Your paper must read like a scientific paper published in a referred journal, by this time
you should be very familiar with such articles.
Title:
Will include the title of the study and the your name. Titles must be descriptive of the
work. Titles such as Lab Report or Gene Report are not acceptable.
Literature review: The literature review should refer only to the "unknown" gene you have
been assigned, although you are going to work with knowns in the laboratory. Here we propose
that the gene you have been assigned exists and functions in the embryonic development of the
horseshoe crab.
Clearly you must learn something about how horseshoe crabs develop. I will provide the
initial references for you, as some of these are difficult to obtain quickly, but you should do some
research on your own. Relate horseshoe crabs to other species, why is this approach valid?
Which species are the best sources of information that might relate closely to the way horseshoe
crabs do things? What is there in the scientific literature about the function of the gene you have
been assigned in these other species? Using the articles you are able to obtain and the textbook
write a brief, review of the role of this gene in embryonic development. Limit this section to
about five pages.
Your final literature review must include at least five research (that is, experimental
articles) papers and at least one literature review article relating directly to the gene you
have been assigned, as well as additional references about horseshoe crabs in particular or
developmental pathways in general (as in what you glean from the text book). I will mark
the paper down if you don't follow these rules.
The preliminary review will likely be sketchy, as you may not have or yet understand the
articles you need. The second draft of the literature review will a complete draft and must
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include either more information or more depth than the first draft. The final draft, obviously,
include either more information or more depth than the second draft. I will mark the paper
down if all you do from one draft to another is correct typos.
You must include a bibliography in any draft you turn in. Please remember that a
responsible scientist works from full text articles, not abstracts* and that plagiarism is
unacceptable. If you are not sure how properly reference your sources, please see me.
Methods and Results (the lab report part)
Clearly write out in your own words what we did in the laboratory using the primers to
known genes. Write this section in the past tense. Use a paragraph format, and remember that
someone must be able to repeat your work from this section. Illustrating your methods section
with tables or flow charts often helps to clarify what was done, but flow charts do not substitute
for the text of the methods section. This section should include a straightforward, factual
presentation of the outcome of laboratory efforts. Do include gel pictures as figures if we are
able to get some!
Discussion of the laboratory work : Discuss the relative success or failure of our laboratory
work with the known genes. What could you do to improve our success? If we have been
sucessful, in what direction might we know go?
Proposed future experiments with your gene: Now use what we did in the laboratory to design
an approach that you would use for a new set of primers to your assigned gene. Then, assume
that you will be successful in obtaining a cDNA to your gene, and propose additional studies to
ascertain the function of this gene in development. This requires a good understanding of the
experiments you have read about and some creativity on your part. Explain exactly what you
propose to do, giving hypotheses and methodology. Then dissect possible outcomes, for
example, you propose to study gene expression using in situ hybridization, what do you conclude
if signal is detected in the limb buds, or in the gut or not at all? Note: transgenic horseshoe crabs
are not possible, so limit your experiments to what can be done directly with embryos.
Bibiliography or References
You must list any sources you used in preparing this paper, including your lab. manual or
handout, and textbook. References must be properly cited in the text, as well. DO NOT USE
FOOTNOTES. Consult a style manual (reference section of the library) for appropriate usage of
citations and how to prepare a reference list. MLA format is best, although many scientific
journals have a preferred format.
This is a citation in the body of the text:
Our results disagree with those obtained by Morris and Weaver (1996) by nearly ten
percent.
This is the (fictional) reference list that follows:
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Dewy, T. H. and Howe, D. N. (1956) Mechanisms of olfaction in insects. J.
Entomological Soc. 23:1-4
Morris, R. W. and Weaver, F. E. (1996) Positive taxis of Sarcophaga larvae in response
to fat substitutes. J. Entomological Soc. 86:1-4
More help:
I keep copies of several source books on scientific writing in my office, don't hesitate to
borrow one. Feel free to consult me, multiple times if necessary, before you hand anything in for
a grade.
If your report is all marked up for grammatical, spelling or composition errors, you may
wish to contact the Writing Center (X 4332)
*Why? Because the abstract is the authors' assessment of what was done and what it
means. A sensible scientist examines the data first and then decides if the authors are justified in
making these claims.
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Biology 306 Laboratory IV:
Journal Club
Practicing scientists frequently share newly published findings with their immediate colleagues
in a forum called a journal club. We will conduct one journal club this semester.
The procedure for preparation to give a journal club talk is:
1. Choose one article on the gene you have been assigned topic for presentation. It must be a
research paper, not a review article, news and views article or something from the popular press.
Choose one that will help you do the best job of explaining the function of the gene you have
been assigned.
2. Obtain a full text version of the article and look it over carefully. Please give me a copy at
least three days before the presentation. If you are able to understand most of the article using
only your textbook and your personal store of knowledge as references, its presentable, if you'll
need more than one other background source, it is not.
4. Review your choice with me.
5. Prepare a 15 minute presentation on the research described in the article. WE GOTTA HAVE
VISUALS!! Flow charts of procedures really help.
Journal club presentations should consist of the following:
Background: in which you place this article in context. Assume we know very little
about the system(s) that you are about to describe.
Methods and Results: Present the experiments that were conducted in the study one at a
time* by answering these questions for each:
What was the experiment designed to do? (you can say this experiment attempts to
show….)
How was it done? Your audience will appreciate diagrams of complex procedures.
What were the data (show and explain tables or figures)?
What new questions arose from these results?
Discussion: Did the experiments conducted support or refute the hypothesis(es) of the study?
Why or why not? Where there any flaws in the study? What new questions do you have?
*Not all experiments of a lengthy paper can be covered in 15 minutes, if it seems that you must
cut something please consult me before you prepare the talk.
Remember to: Organize your information into a clear and concise sequence and make smooth
transitions from section to section. PRACTICE YOUR TALK BEFORE A LIVE AUDIENCE
(preferably of humans), practice more than once, practice more than twice, practice more than
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three times (you get the idea). Speak clearly and look at your audience (even if they are not
looking at you). POINT THINGS OUT ON YOUR VISUALS.
No one is expecting you to have this stuff memorized, but the only way to avoid the
pitfall of reading the paper with your head down is to practice! Nervousness is normal! Take a
deep breath, consult your notes, and resume! Finish your talk with the words: “ Thank you”.
Your audience will appreciate it!
7
GENES AND DEVELOPMENT MINI PROJECT
Power point or web based presentation
Prepare a visually intense presentation that will introduce us to the gene you are working
on, how it operates in other species, what it might do in horseshoe crabs and how you propose to
test that (those) hypothesis. Assume we do not know much about this gene or the species that
have been studied, but don’t bore us with a laundry list of details that we can't grasp in 10 or 15
minutes; hit us with the essential information.
The presentation should consist of an introduction, a review of the role of this gene in
embryonic development in other species, a proposed role for this in horseshoe crab development,
how you might detect this gene in the crabs and what experiments you propose to do if it was
indeed found.
Use figures scanned in from any source, I'll help, to illuminate the role(s) of this gene in
development. Make extensive use of bulleted lists, diagrams and flow charts to illustrate your
points, which must be presented in a logical fashion. Please don't waste time making fancy
transitions from slide to slide, concentrate on the information you are presenting.
Review my advice on journal club presentations before standing up with your
presentation.
Peer and instructor review of presentations
Each student and the instructor will review in written form each presentation according to
a rubric that we will agree on prior to the date of the presentations. It is my feeling that you will
best be able to perform to and evaluate with guidelines that you have participated in developing.
8
Biology 306 Laboratory I
Differentiation
In this laboratory you will examine an organism that is capable of regeneration, and a
single celled organism, Dictyostelium, that is capable of assembly and differentiation. We will
initiate the observations in this lab, but you must return to your cultures several times in the next
week to complete the exercise.
Cactus spine needles:
A cactus spine needle is useful for bisecting such organisms as Hydra, Lumbriculus or
Planaria, and for microsurgery on chick embryos.
With a razorblade slit and slightly sharpen the end of an applicator stick.
Obtain a cactus spine; insert the blunt end into the slit and seal with nail polish. Make several
needles using different spines and different angles of insertion. Store your needles in the
instrument cups provided.
Regeneration in Lumbriculus
Lumbriculus variegatus, the California blackworm or mud worm is a segmented,
hermaphroditic annelid that prefers extremely wet to aquatic environments. The body is
composed of about 120 to 250 segments. Both sexual reproduction, which involves embryonic
development from a cocoon and asexual reproduction by self-fragmentation occur. It is possible
to culture these organisms in the laboratory under very simple conditions by taking advantage of
their ability to reproduce asexually.
Capture and handle worms and worm fragments while they are in spring water using
transfer pipettes. Do not use forceps or needles, as the worms will self fragment (automize).
When a worm is separated into fragments, the pieces regenerate over a period of about two
weeks. A regeneration blastema, a region of dedifferentiated cells will form at the cut surfaces.
The regenerated segments are smaller and lighter in color than the original segments, and the
original segments may have undergone a developmental reorganization (or re-specification, if
you will) to perform new functions.
Obtain a specimen of Lumbriculus in a dish of spring water, and observe its behavior
under the dissecting microscope. Locate the head and “tail” ends and observe locomotion.
You should fragment the worm using a razor blade or cactus spine needle into pieces
small enough to fit in eppendorf tubes. Don’t make the pieces too small or you will have far too
many to record over the next two weeks.
Consult your partner: How will you determine that regeneration has indeed taken place? Count
segments? Measure lengths?
Record the initial length or segment number of each fragment you create.
Fill each eppendorf tube with ~1 ml of spring water, and label the tube to indicate the specific
pieces that you will transfer into it. Transfer each segment to the tube as you cut it. Cap the
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tubes tightly and store them in the laboratory until next week’s laboratory when you will observe
and document the progress of regeneration in these segments.
Differentiation in Dictyostelium discoideum
The “cellular slime mold”, which is not a mold is an excellent system in which to study
differentiation. Single celled myxamoebae feed on bacteria, aggregate when starved, and the
multicellular aggregate called a grex or slug migrates in a coordinated fashion, seeking further
food. If none is encountered, the grex differentiates into a sporocarp to weather out the bad
times. New myxamoebae are produced when the sporocarp “germinates”. The life cycle of
Dictyostelium is diagrammed below, see Chapter 2 of your textbook for a review of some of the
interesting work that has been done of Dictyostelium.
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Culturing Dictyostelium
Dictyostelium and E. coli or Enterobacter stock cultures will be provided.
Wash hands and disinfect surfaces. Mark bottom of two lactose agar plates with name, date and
compass directions (N, S, E, W)
Using a flamed metal loop (or sterile plastic one), aseptically inoculate a dish of lactose agar
with E. coli using a North –South streak.
N
S
Using a flamed metal loop (or sterile plastic one), scrape up enough Dictyostelium
growth, about the size of a pea, for inoculation.
Inoculate the same lactose agar plate with the Dictyostelium, streaking now in the East –West
direction, and being sure to leave bacteria free areas where the Dictyostelium can migrate and
E
W
differentiate. Repeat with second plate.
Dispose of all contaminated materials in the biohazard bags provided.
Disinfect the work area, and wash your hands.
Incubate the plate at room temperature in sunlight for 7 days, checking on the culture every two
days (approx.). In the past, cultures have grown well in Room 423 on the windowsill. It is best
to leave the cultures upside down so that condensation on the lids does not spread bacteria over
the entire plate.
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Record your observations, which should be made every day, in your laboratory notebook.
References:
Cactus spine needles: England, M.A. (1981) The application of the scanning electron microscope
to the analysis of morphogenetic events. J. Micros. 123:133-146
Dictyostelium culture and life cycle: Carolina Biological Supply handout CB161389602, 1996
Dictyostelium life cycle diagram dowloaded from
http://biodidac.bio.uottawa.ca/Thumbnails/MYXO002B-GIF.htm on 8/11/00
Lumbriculus: Carolina Tips, 59 (3) 1996
Thanks to Leland Johnson for ideas presented at the Developmental Biology Laboratory
Teaching workshop, 1997
Frances E. Weaver 1/24/1999; 8/11/00
12
Biology 306 Laboratory II
Meiosis and fertilization
Chlamydomonas
Oedogonium
Sea Urchins (if available)
Gametes and gametogenesis: slide study
Ascaris
Frogs
Chicks
Mammals
Ongoing:
Differentiation in Dictyostelium
Regeneration in Lumbriculus
Isogamous fertilization in Chlamydomonas
( see Gilbert, Developmental Biology 6th ed. Fig 2.8; p. 33)
These haploid cells exist as two mating types, + and - both participate equally in
contributing cytoplasm to the zygote. Recognition is between adhesion molecules on the flagella
membranes. The plus cell extends a fertilization tube, contacting a receptor site on the minus
individual. In practice, cells tend to aggregate in clumps. Once two cells have mated, cell walls
are dissolved at the anterior poles, the cytoplasms fuse, and the nuclei unite to form a zygote
(diploid). For a brief period a quadriflagellated zygospores may be visible, these loose their
flagellae when they become zygotes. After a period of dormancy a zygote “matures” undergoing
meiosis (just like other diploid cells) and then germinates, producing 2 + and 2 - haploid
organisms. This organism thus displays both sex and reproduction.
Both sexual and asexual reproduction are
depicted here.
13
Four hours before lab the + and - strains were placed under a lighted desk lamp at a
distance of 18 to 25 cm. At t = 2hr the equal amounts of the mating solution (provided by
Carolina Biological Supply) was added to the cultures, and they were gently agitated and
replaced under the lamp. These cells should now be ready to mate.
Mix a drop of + with a drop of - strain on a slide, preferably in an observation chamber
constructed using a cover slip and Carolina observation gel. Work quickly!! Use just enough
fluid to keep the cells moist, and seal the edges of the coverslip with the gel or nail polish.
Place the slide on the compound microscope, observe the cells, and continue observations at
convenient times throughout the lab period. In a healthy culture the entire life cycle will be
complete in 3 hours.
Mating in Oedogonium
In the lab we will have a representative of these oogamous filamentous green algae which
are often found growing epiphytically on aquatic plants such as water lilies. Oedogonium may
reproduce asexually via budding of zoospores from vegetative cells. Zoospores posses about 120
flagellae!
In the sexual phase, the large egg is produced in an oogonium. Single filaments are either
"male" or "female" or both depending on the species. An oogonial pore opens on one side when
the egg is ready for fertilization. Sperm are multiply flagellated and produced in shortened boxlike structures called antheridia.
In those species that have both “male and female” on the same filament, mature eggs
produce a substance chemotaxtic for sperm (sound familiar?) this compound is one of a group of
substances called erotactins. A diagram of Oedogonium:
oogonium
antheridia
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Obtain a specimen of the Oedogonium mating culture on a microscope slide and observe
under the compound microscope. You are looking for oogonia and antheridia. Depending upon
the length of time since the initiation of mating, you may also see zoospores (use a higher power
objective and adjust the lighting). Young germlings might be found at the edge of the mating
dish. Sketch and label what you see. This organism can also be used for student projects, but be
aware that 48-72 hours must elapse between the initiation of mating and the appearance of large
quantities of oogonia and antheridia.
Fertilization and early cleavage in sea urchins:
Sea urchins are deuterostome organisms that have radial symmetry as adults, but bilateral
symmetry as larvae. The eggs are variously colored, but the developing embryos become
progressively less opaque. A jelly coat, which releases peptides that attract sperm, very useful
when the adults are meters apart in the ocean, surrounds the each egg. This egg jelly may or may
not be visible. Some echinoderm species invest their jelly with pigment granules. Fertilization is
external, and the entering sperm stimulates the exocytosis of cortical granules to the egg surface,
resulting in the elevation and eventual hardening of a fertilization envelope. The fertilization
envelope represents a slow block to polyspermy, and also serves to protect the embryo until it
reaches the blastula stage. Blastulae, which hatch from the fertilization envelope, are ciliated
and capable of swimming.
For a summary of early embryonic development in sea urchins, view the appropriate
sections of the video "A Dozen Eggs"
The gonads lie close to the under side of the aboral surface. The sexes look alike, so the
only way to tell them apart is by getting them to release their gametes. Eggs and sperm are
obtained by intra-coelomic injection of 0.5 M KCl. The female is placed over a beaker of
seawater to collect the eggs. Sperm are collected dry onto a plastic petri dish.
Observe gamete collection
Obtain a sample of eggs on a microscope slide, and observe. The dot in the eggs is the
egg nucleus or germinal vesicle.
Add a drop of "dry" sperm to the edge of the coverslip and watch the eggs carefully as
the sperm make contact. Can you see the fertilization envelope as it rises? If thing are too
murky, try diluting the sperm with seawater.
Sketch your results, refer to chapter 2 of your atlas.
We will batch fertilize some eggs and check for early cleavage throughout the period. If
we are successful, we will keep our cultures and observe them over the next week.
Gametogenesis: Study of Prepared slides
Bring Mathews and Schoenwolf Atlas of Descriptive Embryology 5th edition with you to lab
Using your compound microscope, your textbook (see Chapter 19 and study, compare and sketch
the prepared slides of:
Frog ovary and testis; see chapter 5 of your atlas.
Chick ovary and testis (lower power lenses are best for the ovary);refer to chapter 6 of
your atlas.
Small mammal ovary and testis
15
Human sperm smear (use oil immersion lens)
Human or cat ovary and testicle
One process that you are observing is meiosis. Meiosis takes place in the gonad of all these
species. Following or accompanying meiosis are the differentiation steps. The end products;
oocytes, eggs or sperm are very different from each other.
Once you have completed your sketches answer the following thought and discussion
questions for each of the species observed: How do the differences in the morphology of the
eggs and sperm arise during gametogenesis. How do these visible differences relate to the
differing functions of the gametes in fertilization and early development? Are there cellular or
acellular materials associated with the egg once it is ovulated, such as follicle cells? How do the
differences in structure of the eggs from the various species relate the environment in which the
embryos will develop?
Nuclear events of fertilization: Prepared slides of Acaris
The order and extent of the nuclear events of fertilization vary with species, but these
usually include sperm entry, decondensation of sperm DNA, swelling and union of sperm and
egg pronuclei. In most mammals meiosis II resumes and completes at the time of fertilization in
order to create the female pronucleus, so there are a number of things happening at once.
Examine prepared (use 45X and 100X oil immersion lenses) of uteri of the parasitic
roundworm Ascaris megalocephala also known as Parascaris aequorum. Sperm are amoeboid
and crawl into the uterus where there are many ova in various stages of maturation. The sperm
are darkly staining, almost triangular bodies, maturation (decondensation) and fusion of the male
and female pronuclei are easy to see, as is the beginning of mitosis. There are only 2
chromosomes per haploid cell (4 per diploid), so you should be able to spot the condensed
chromosomes. Remember that you are looking at the whole worm on most slides so your first
job is to locate the uterus.
Make detailed, labeled sketches of the stages of fertilization you are able to locate
referring to chapter 2 of your atlas. When might the DNA synthesis needed for cleavage
occur?
If we are successful with the sea urchins, you may have to complete your slide studies in
the next laboratory period.
References
Carolina Biological Supply Company "Unialgal Mating Kits" handout #cb160617810
Information on the life cycle of Oedogonium was taken from Morphology of Plants, 3 ed. by
Harold C. Bold, Harper and Row, New York, 1973.
Oedogonium life cycle diagram downloaded from the site below on 8/11/00
http://biodidac.bio.uottawa.ca/Thumbnails/filedet.htm?File_name=CHLO013B&File_type=GIF
Thanks to Leland Johnson for ideas presented at the Developmental Biology Laboratory
Teaching workshop, 1997
Frances E. Weaver 1/27/1999; 8/11/00
16
Biology 306 Laboratory III
Lab notebooks due on the Monday that follows this lab.
Early Frog Development: Exercise taken from S. L. Biroc, Developmental Biology. A
Laboratory Course with Readings, Macmillan, New York, 1986, attached.
Early development of Zebra fish
http://zfish.uoregon.edu/zf_info/zfbook/stages/stages.html
Ongoing:
Differentiation in Dictyostelium
Regeneration in Lumbriculus
Sea Urchin development (if available)
Slide study, if incomplete from last week.
Bring Mathews and Schoenwolf Atlas of Descriptive Embryology 5th ed. and your text
book with you to lab.
We must work on frog and zebra fish embryos at the same time, so be prepared to go back and
forth.
Early development of Zebra fish
Zebra fish development is considerably more rapid than frog. Depending on the time of embryo
collection this morning, our embryos may be well into cleavage by lab time.
Obtain a dish of zebra fish embryos in aged tap water, observe and sketch the resulting stages of
development approximately every half-hour. Refer to the on-line manual of developmental
stages available at http://zfish.uoregon.edu/zf_info/zfbook/stages/stages.html (which we should be able to
access during the laboratory period) and your text book. At the end of the lab period, transfer
your embryos to a clean dish of tap water and count them, discarding any obviously defective
ones. Record the number of embryos in your lab book. Return to the lab to observe them on
Friday and Monday, record the numbers and developmental stages each time. What percentage
of your embryos matured to larvae?
Early Frog Development: Exercise taken from S. L. Biroc, Developmental Biology. A
Laboratory Course with Readings, Macmillan, New York, 1986, attached.
Dictyostelium: complete observations if you have not already done so.
Lumbriculus: Record in a table (% length change/piece or % increase in segment
number) a numerical assesment of ; and comment on the extent of regeneration in
Lumbriculus. What factors such as size of piece or position of the cut influence regeneration?
Did your middle pieces show signs of developing both heads and tails? Are the regenerating
segments smaller or larger than the "old" segments?
17
Biology 306 Laboratory IV-VI
Genes and Development Mini-Project
Detailed laboratory instructions will be provided prior to the start of week IV. Project
instructions and guidelines for journal club appear in this booklet.
We will be attempting to prepare and clone cDNAs of developmental interest from RNA isolated
from horseshoe crab embryos.
Outline of Activities:
Week IV
Each student will be assigned a gene of interest and a pair of degenerate oligonucleotides to
sequences derived from homologs of that gene in vertebrate and invertebrate species.
We will go over the requirements for this research project, journal club and presentations.
We will perform RT-PCR using these oligonucleotide primers on total RNA isolated from
horseshoe crab embryos.
It will be your responsibility to research the gene you have been assigned for your paper and
presentation. We will make use of on-line data bases and search engines to locate, order and/or
print out the first few references during this period. The remainder of the literature based
research will be done on your own time.
Week V
We will determine our success, or lack of it in the RT-PCR reactions by gel electrophoresis.
If successful we will attempt to ligate our cDNAs into plasmid vectors. Otherwise, we will
"trouble shoot" the RT-PCR
Week VI
We will transform our successful ligations into E. coli and select for transformants or
We determine our success, or lack of it, in the repeated RT-PCR reactions by gel electrophoresis.
If successful we will attempt to ligate our cDNAs into plasmid vectors. Otherwise, we will
continue to "trouble shoot" the RT-PCR.
18
Biology 306 Laboratory VII
Drosophila Development and Imaginal Disk Dissection
See exercise attached from Mary S. Tyler, Developmental Biology, A Guide for Experimental
Study Sinauer Associates, Sunderland MA 1994
Eversion of Imaginal Disks
Ongoing:
Genes and development mini project, if needed.
Premature Eversion of Imaginal Disks
In Drosophila, the mouthparts, antennae, eyes, legs, wings, halteres and genitalia of the
adult develop during pupation from imaginal disks. These disks represent tissue set aside from
the larval structures. The original disk invaginates from the body wall (see figure 17.1 from
Cruz, below) and becomes, effectively, a collapsed pouch. The pouch everts at pupation, in a
motion that is rather like the extension of a telescoping structure such as a pointer or a car
antenna (see figure 19.16 from Gilbert, below). Molecular analysis has shown that the anterior
posterior axis as well as the specific regions of each structure is determined prior to eversion and
that natural eversion is primarily the result of shape changes of the disk cells. Although disk
extension during pupation is normally under hormonal control (ecdysone), we can prematurely
evert disks by treating them with trypsin.
Obtain several third instar larvae and place them in a drop of Insect Ringers in a small petri dish.
Anesthetize the larvae by placing the dish on ice for a few minutes.
Place an anesthetized larva on a microscope slide
Using the naked eye or the dissecting microscope, firmly grasp a larva by its anterior end with
one pair of fine forceps and about 1/3 of the way down with another pair.
Pull the larva apart, the digestive tube should be expelled from the preparation.
Most of the disks are associated with the esophagus and the anterior end of the digestive tract.
They are clearer than fat bodies or digestive tract and recognizable by the concentric marks of
the imaginal tissue.
Put a little more Insect Ringers on the slide and dissect the disks away from the rest of the larva
using catcus spine needles. Take care not to let the disks dry out.
19
Using the compound microscope, sketch and label the disks using the figures below and in the
previous exercise as guides.
Push your dissected disks to one side of the slide, and remove any other debris.
Remove as much Ringer's as possible, and cover the disks with 0.1% Trypsin in Insect Ringers,
pH 7.0.
Set the slide aside and make observations at half-hour intervals. Eversion make take up to an
hour (or more) to complete.
When eversion appears to be completed, sketch the results of the trypsin treatment, being sure to
use the same magnification as the disks sketched before eversion. Did eversion occur?
Speculate as to why trypsinization leads to eversion.
References
Yolana P. Cruz Laboratory Exercises in Developmental Biology Academic Press, San Diego
1993
Scott F. Gilbert Developmental Biology 5th edition, Sinauer Associates, Sunderland MA, 1997
Mary S. Tyler, Developmental Biology, A Guide for Experimental Study Sinauer Associates,
Sunderland MA 1994
20
Observation of Chick Development: Exercise taken from Johnson and Volpe, Patterns
and Experiments in Developmental Biology; Wm. C. Brown, Dubuque IA 1973, attached
Explantation of chick embryos and cardia bifida Exercise from S. L. Biroc,
Developmental Biology. A Laboratory Course with Readings, Macmillan, New York, 1986,
attached.
Bring Mathews and Schoenwolf Atlas of Descriptive Embryology 5th edition with you to
lab.
A word of caution: Eggs may b e contaminated with Salmonella bacteria. Typically the
contamination is on the outside of the shell, but occasionally bacteria are found internally. We
are handling a great number of raw eggs, please exercise great caution. Wash the eggshells
gently and be certain to wash your hands thoroughly before leaving the lab.
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Biology 306 Laboratory X
Explantation of chick embryos, staining for cartilage
Bring Mathews and Schoenwolf Atlas of Descriptive Embryology 5th edition with you to lab
Other references on chick development will be available in the laboratory.
Ongoing:
Repeat of cardia bifida
In the development of the appendages cartilaginous skeletal elements precede the formation of
bone proper. This process is known as endochondral ossification. Here we will explant and stain
chick embryos at relatively early stages for the formation of cartilage. Our expectation is that the
developing spinal vertebrae and the bones of the developing appendages will be intensely
stained. In earlier embryos we may see more diffuse staining, if so we will attempt to correlate
these patches with our understanding of chondrogenesis. The procedure is simple, but does
require that you return to the lab to change solutions.
The fixative and staining solutions are either caustic or toxic or both. Take note of the
position of the eye wash station. Wear a lab coat and take care not to spill or spatter your
fixative and staining solution. Report any spills to the lab instructor.
Read the procedure carefully. Some steps must be done 1 and 2 days after lab. Devise a plan
for sharing the labor before starting the explantations.
Chicken eggs that have been incubated between 3 and 10 days will be available.
Crack incubated eggs into saline and explant embryos from at least two different stages into a jar
of 5% TCA (trichloroacetic acid). Note the developmental stage of each embryo you explant,
and in particular the status of the limb buds, by referring to the available atlases and sketching
the embryos in your notebook. The acid treatment will fix the tissues in place.
Fix the embryos for 3hr or longer in the 5% TCA. During this time, explant 24hr embryos and
repeat cardia bifida experiment if last week's was unsuccessful.
Use just enough of the solutions to cover the embryos.
Pour the TCA off into a waste bottle and wash the embryos once in distilled water (use forceps to
transfer embryos or hold them in the jar).
Cover the embryos with Alcian Green stain (0.1% in acid alcohol) Acid alcohol is 1% HCl in
70% ethanol. Leave several hours or over night. Stain will bind tightly to cartilage and loosely
to other tissues so the embryos will be very green after staining.
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Pour off the stain and transfer the embryos to acid alcohol (destain), incubate overnight.
Replace the acid alcohol with absolute ethanol, two changes of one hour each. This is a
dehydration step to remove excess water.
Transfer the embryos to methyl salicylate to clear, clearing may take several hours.
Caution - methyl salicylate is toxic! Handle with care, and clean up any spills.
Observe the stained embryos in methyl salicylate under the dissecting microscope. Sketch and
label what you see, paying particular attention to the limb buds. Use your textbook or a
comparative vertebrate anatomy book as a reference for the bones of the chick wing and leg.
Reference: Tickle, C. (1993) Chick Limb Buds in Essential Developmental Biology A Practical
Approach, C. D. Stern and P. W. H. Holland ed., IRL Press at Oxford University Press, Oxford
UK pp. 119-125
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Developmental Biology Laboratory XI
Effect of retinoic acid on zebrafish development
http://zfish.uoregon.edu/zf_info/zfbook/stages/stages.html
Obtain several zebrafish embryos and sort them by developmental stage.
Be sure to have both control and experimental embryos free of debris
Design an experiment to test the effects of retinoic acid on embryonic development.
The RA concentration can vary from 0.1 M to 1 M. Depending on the number of embryos we
get, you may be able to use only one or several concentrations.
We will have:
aged tap water
small glass petri dishes
transfer pipets
A stock solution of retinoic acid in aged tap water, the concentration of which will dictate how
much or how many dilute solutions we can make.
microscopes
Your experiment could include prolonged exposure or be a pulse chase (washout) experiment.
You must plan at least one time outside of this period to view the embryos (larvae), which can
include Friday at 10 a.m. (our class time).
Make groups a manageable size (5-10 if we have that many) and be sure to reserve some
untreated embryos as a control.
Keep accurate logs of your treatment (s) and their results in your lab book.
Ongoing: Observation of chick embryos stained for cartilage
Caution - methyl salicylate is toxic! Wear gloves and tip the embryos out (or transfer them with
forceps) into a glass petri dish under the hood. Pour excess methyl salicylate back into your
staining bottle, leaving just enough to cover the embryo. Handle stained embryos with care, and
clean up any spills.
Observe the stained embryos in methyl salicylate under the dissecting microscope. Sketch and
label what you see, paying particular attention to the limb buds. Use your textbook or a
comparative vertebrate anatomy book as a reference for the bones of the chick wing and leg.
Biology 306 Laboratory
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Developmental Biology Laboratory XII
Comparative study of stages of mammalian embryonic development.
See typewritten exercise by Dr. Weaver, attached.
Bring Mathews and Schoenwolf Atlas of Descriptive Embryology 5th edition with you to lab
Other references on rat development will be available in the laboratory.
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Biology 306 Laboratory XIV
Presentations and Peer Reviews
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