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| ==1. QUIZ==
| | Please note the different location for this week’s practical class: Wallace Wurth Teaching Lab 120. |
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| [[File:Chicken.jpeg|180px]]
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| ==2. Gastrulation, early somitogenesis and neurulation lab - early chicken eggs ==
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| The chicken (Gallus gallus) embryo is an excellent model for the study of early vertebrate embryogenesis and later organogenesis. The embryo is encased within a hardened eggshell which provides a natural incubator or culture dish. Through a hole in the eggshell, the embryo can be visualized and easily manipulated with microsurgical tools or gene constructs, then allowed to continue development in ovo to determine the consequence of the experimental manipulation.
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| Fertilized chicken eggs are readily available anywhere in the world and the equipment needed is minimal – a humidified incubator (39oC, no CO2 required), a dissecting microscope, microsurgical tools that can be prepared in the lab or purchased, and either a hand-held mouth pipette or a manufactured micromanipulator and picospritzer.
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| Fertilized eggs can be held at between 13-16oC for up to 1 week prior to incubation. They are incubated at 38ºC-39ºC to the desired stage in a humidified incubator with the eggs placed on their side (horizontal). For long-term post-operative survival, it is best that the eggs be left in the incubator until experimental manipulation. However, eggs can be removed from the incubator and held at room temperature to slow development.
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| For staging of chicken embryos: https://embryology.med.unsw.edu.au/embryology/index.php/Hamburger_Hamilton_Stages
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| '''SAFE WORKING PROCEDURES AND ANIMAL ETHICS'''
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| '''Risks Associated with Practical'''
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| Eggs have the potential to be contaminated with the bacteria Salmonella. Wear gloves throughout and students should always wash their hands before leaving the lab.
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| Dissection implements are sharp, so students should take care not to cut themselves or other students.
| | PRACTICAL CLASS PROGRAM: |
| | | * Weekly Quiz + revision (15 minutes) |
| Students should wear a lab coat, gloves and enclosed shoes to protect themselves from egg splatter.
| | * Practical class activities (90 minutes) |
| | | * Practical Class Revision (15 minutes) |
| '''Animal Ethics Compliance'''
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| The procedures used in this practicum are in compliance with the UNSW Animal Care and Ethics Committee and the National Health and Medical Research Council ‘Australian code of practice (8th edition, 2013).
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| '''EXPERIMENTAL PROCEDURES'''
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| '''Opening of the egg'''
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| 1. Eggs were incubated for several days at 38.5°C to generate embryos that undergo organogenesis.
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| 2. Each student pair should have one egg and an an egg holder, a pair of blunt forceps and a pair of scissors, 3 petridishes, PBS, disposable pipet, syringe and 23G needle, Indian ink solution, dissection microscope.
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| 3. Put the egg into the holder with the blunt end up (pointed end down).
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| 4. Use the pointy end of the scissors to tap a hole in the top of the egg into the air chamber, be very careful not to push the scissors too far into the egg.
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| 5. Use the forceps to pick bits of shell out. Do not remove egg shell beyond the air chamber. You will see the air chamber and the vitelline membrane.
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| 6. Carefully remove the vitelline membrane at the top.
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| 7. If you do not see the embryo (a ring of blood vessels should be visible) then gently swirl the egg so that it floats up to the top of the yolk. If this doesn’t work you will need to take another egg if available.
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| 8. Use the forceps to break off pieces of shell down to the yolk so that the embryo (visible as a ring of blood vessels) is exposed and the rim of the shell is just above the surface of the egg white or albumen.
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| 9. You may be able to see the heart beating without magnification. If not, then put the egg under the dissection microscope.
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| '''Early stage somitogenesis embryos'''
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| 1. Draw Indian Ink solution (if available) up into a 1 ml syringe fitted with a 23G syringe needle.
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| 2. Open an egg as described above. Once the shell has been removed down to the level of the yolk and the vitelline membrane has been removed, slide the syringe needle under the
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| 3. embryo. It is easiest to insert the syringe needle vertically at the edge of the egg initially and then rotate the needle until it is almost horizontal using the edge of the egg shell as a support. The tip of the needle should end up just below the embryo in the centre of the egg.
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| 4. Slowly inject the Indian ink solution to reveal the embryo and view under a dissecting microscope.
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| 5. Identify the HH stage and the diverse embryonic structures. See last page or link below.
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| https://embryology.med.unsw.edu.au/embryology/index.php/Hamburger_Hamilton_Stages
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| 6. Draw your embryo, indicate the Hamburger & Hamilton (HH) developmental stages using the guide on course manual pages 22-24, and annotate the embryonic structures that you have identified.
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| 7. Hand your drawing in at the end of the prac.
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| '''Observation of developmental stages'''
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| Move around the class to study the embryonic chicken stages of your colleagues. Identify the following structures:
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| - Amniotic sac
| | PRACTICAL CLASS ACTIVITIES |
| - Hensen’s node
| | # Fertile egg dissections |
| - Neural groove and folds
| | # Whole body regeneration in planarians |
| - Head ectoderm
| | # (There will be back up activities available if required) |
| - Somites
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| - Brain vesicles
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| - Cardiogenic mesoderm
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| - Heart and vasculature
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| | LEARNING OBJECTIVES: |
| | * Understand the process of gastrulation, and being able to identify the defining structures in the chicken embryo. |
| | * Understand early neurulation, mesoderm and heart development, and being able to identify the defining structures in the chicken embryo. |
| | * Understand the regeneration process of planarians following amputation and being able to identify blastema formation, antero-posterior (A/P) and dorsal-ventral (D/V) axes conservation, eye regrowth, re-pigmentation of the regenerated tissues, and worm behavior. |
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| '''Tidying up after the prac'''
| | PRACTICAL CLASS NOTES: |
| - Please discard egg waste and disposable consumables into the yellow bins (not the paper bins).
| | The practical class notes are available on Moodle |
| - Dispose of Syringes and needles in sharps bins
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| - Wash forceps and scissors with water, and wipe with ethanol, place in provided containers on the sink.
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| - Clean surfaces (microscope stages, benches) with water and ethanol, and wipe dry.
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