Download Fluorescent Analysis of Drosophila Amelia Dosio Studies have

Fluorescent Analysis of Drosophila
Amelia Dosio
Studies have shown that Drosophila melanogaster, the common fruit fly, is a prime
model for viewing developmental stages under florescence. Drosophila gives us a model system,
in biology, on the aspects of: formation, behavior, ageing, evolution with production of a large
number of offspring for viewing. Drosophila’s rapid development can quickly get in the way of
long periods of developmental viewing. With this, it is important to master techniques that will
freeze and enhance developmental aspects for study.
Immunofluorescence and Western Blotting are useful techniques that will aid in this
examination. For my analysis of Drosophila, the following are key points that were focused on:
the development of tools and techniques for a teaching lab setting and to observe outcomes of
existing protocol alternations. With Immunofluorescence and Western Blotting my main
objective was to provide a backbone study of antibodies to proteins important to development
and observe the antibody patterns made when placed under florescence. Special attention was
placed on the concentration and molecular weight of each antibody. It was found that a high
enough concentration was needed in order to express the antibody just enough for proper
observation.
Immunofluorescence is an efficient procedure that is inexpensive and easy to perform.
Embryos of early developmental stages are more commonly used in this technique. They provide
a series of nuclear divisions that establish patterning in the embryo that can be analyzed through
florescence. The first step in Immunofluorescence is the addition to bleach to remove the
chorion or the outer shell. The embryo has a series of shell like protection barriers that allow the
embryo to be protected during growth. These shells must be broken in order to allow the
antibodies to penetrate and bind for later detection under a florescent scope. Once the chorion
has been removed, a solution of formaldehyde is added to fix the embryos or to freeze them in
time. Once the embryo is fixed the vitelline membrane must be removed. Both methanol and
heptanes are used to separate the vitelline. This process must be done quickly since embryos can
become trapped in the membrane or not free of optical obstructions.
Embryos will need to be rehydrated since antibodies are present in aqueous forms. The
embryos will need to be in the same phase to allow proper binding of antibody. Increasing
proportions of methanol and PBS are used. Once the embryos are rehydrating stock piling can be
completed or the continuation of Immunofluorescence can be done. At this stage, the embryos
are prone to permeability or the making of holes on the surface of the embryo. A blocking
solution of BSA and PBT is used to coat the embryo and prevent non specific binding of
antibodies to occur. Primary antibody has been pre absorbed. Pre absorbing allows only the
specific antibody to be bound to the embryo since the entire nonspecific sites have been absorbed
previously by other embryos not used. Once absorption of the primary antibody is complete,
secondary antibody is added. Secondary antibody adds in the detection of primary antibody
through florophones. These molecules will be excited with ultraviolet light to observe our protein
of interest. However, they are light sensitive and bleaching can occur if left in light for a long
period of time. Once binding of primary and secondary antibodies have taken place embryos will
need to be dehydrated and prepared for mounting. A clearing solution is needed to make the
embryo have the same refractive qualities as the yolk and to improve the optics under the scope.
Western Blotting allows us to measure concentration amounts and determine the
molecular weights of certain antibodies. Embryos can be used as whole or separate extractions as
long as there is a large enough sample to obtain good results. For the analysis of interest, whole
embryos are prepared and used. Embryos are collected and dechorionated with bleach to remove
the chorion as before in the Immunofluorescence step. Laemmli Sample Buffer is used in twice
amounts as embryos collected. Laemmli buffer must be fresh since we will need the active
reducing agent of beta-mercaptoethanol or DTT to be present. Embryos are squashed and spun
down to collect all supernatant for SDS-Page gel electrophoresis run. Bio-Rad Semi-dry
electrophoretic transfer to a membrane from SDS gel is completed and later the addition of
antibodies. It must be noted that blocking also needs to take place since antibody binding is the
same as in Immunofluorescence protocol. Once antibodies are added, development with Alkaline
Phosphatase and HRP regents completes the western blot process. Both regents will process
bands at the molecular weight of target protein and band strength will show approximate
concentration and activity of protein present.
The fluorescent analysis of Drosophila has been broken into nine experiments to get a
wide range of experimental data for analysis. With the two protocols explained above, the
following is the results of each experiment in greater detail.
Experiment 1:
Primary:
 mouse anti-actin 1:250
 rat anti-alpha tublin 1:250
Secondary:
 goat anti-rat 1gG Alexatluor 488 Green 1:250
 goat anti-mouse IgG Texas Red 1:250
This experiment was to obtain a base line for antibody Immunofluorescence. It is seen in the
images below that actin is present throughout the embryo. It is noted that in many of the images,
it was found that there was a very high concentration of secondary antibody. In later
experiments, possible experimental changes will be noted to help fix this. The secondary
antibody of IgG Texas Red gives us the red color. The scope that was used is programmed to
read this antibody to detect protein of interest.
Figures 1&2: Actin present in fly embryo under florescence.
Experiment 2:
Primary:
 Monoclonal anti-beta tublin 1:20
 Rat anti alpha tublin 1:250
Secondary:


goat anti-rat 1gG Alexatluor 488 Green 1:250
goat anti-mouse IgG Texas Red 1:250
In this experiment, beta tublin was the protein of interest. Beta tublin is known to been seen to
concentrate at the center of the embryo. In this experiment there also was a high concentration of
secondary antibody that was seen. In the images below, blue is DNA stating which is seen at
very high concentration at the center of the embryo. Beta tublin was sparely seen in the images
and leaves to question of what went wrong? Some concern was that the secondary was very high
and giving us the results seen. A comparison image taken from The Interactive Fly website is
used in comparison here for the banding pattern of beta tublin.
Images 3 &4: Beta Tublin images of different embryos treated with beta tublin
Image 5: Beta Tublin expression and
banding pattern taken from
interactive fly website.
Experiment 3:
Primary:


Mouse anti-alpha tublin 1:1000
Mouse anti-beta tublin 1:1000
o 45 kDa
Secondary:

Goat anti-mouse IgG Alkaline Phosphatase
For this experiment western blotting was used to test to see if the proteins of alpha and beta
tublin were present and strong enough in the obtained embryos. It was know that beta tublin was
at a molecular weight of 45 kDa. Mutant flies and GFP flies where tested in this experiment. It
was seen that the GFP flies gave weaker band strengths at approximately 50kDa. It was
concluded that this could be a result of a weak expression in GFP flies or not enough fly embryos
to give a good signal. As an overall result, the western blot concluded that these two proteins
where a good baseline for protein analysis. The proteins were active enough to produce a strong
enough concentration band with the help of the AP regents to develop them.
Mouse anti-alpha tublin 1:1000
Mouse anti-beta tublin 1:1000
Western Blot 1: Analysis of beta and alpha tublin. Note bands present at 50kDa
Experiment 4:
Primary 1
 Mouse beta tublin 1:1000
 Rat alpha tublin 1:1000
o 45 kDa
Primary 2
 Mouse Sir 2 1:500
o Sir2 92kDa
 Rat alpha tublin 1:1000
Secondary
 Goat anti mouse AP
 Goat anti rat HRP
In this experiment a strong antibody of Sir2 was tested. Results that wanted to be seen, of this
protein, is it necessary to have a large or small concentration to do immunoflorescent analysis.
Another method of HRP was added to the development of the western blot. It was seen that beta
and alpha tublin bands didn’t show upon completion of regent addition. AP regents were added
first to blot then HRP. AP regents gave Sir2 bands present at about 110kDa at a high visibility. It
was later concluded that HRP & AP should be tested again in the next experiment to see if it
made a difference in order of which they should have been completed.
Mouse beta tublin 1:1000
Rat alpha tublin 1:1000
Mouse Sir 2 1:500
Rat alpha tublin 1:1000
Western Blot 2: Analysis of Sir2 with HRP and AP regents
Experiment 5:
This experiment was a repeat of number four. Same primary and secondary antibodies where
used with only experimental variations of doing HRP or AP first. The same experimental results
had occurred in this experiment as it did in experiment 4. The alteration of HRP or AP first
didn’t make a difference. It was concluded that maybe the regents where bad, the antibody was
old or a stronger expression of tublin was needed.
Western Blot 3: Analysis of Sir2 and tublin with alterations given to HRP and AP regents
Experiment 6:


Primary:
o Mouse Sir2 1:25
o Rat alpha tublin 1:250
Secondary:
o goat anti-rat 1gG Alexatluor 488 Green 1:250
o goat anti-mouse IgG Texas Red 1:250
It was known that Sir2 was an active protein in high amounts per dilution. Sir2 was put to the test
of Immunofluorescence since it was known to be highly active antibody and we could hope to
get a good result. Final analysis showed that there was a presents of secondary antibody and no
presents of the Sir2 pattern. It raised concern that Sir2 may not be binding correctly to the
embryo or antibody matching was wrong; needed different primary or secondary matching. With
seeing a constant product of secondary in high concentration it leaves to question can we dilute
the secondary antibody to a degree that it allows for visibility of primary antibodies?
Image 6&7: Sir2 embryo analysis
Experiment 7:
Primary:
•
•
Rat alpha tublin 1:250
Mouse anti even-skipped 1:250
o 38kDa
Secondary
•
•
CO66 Rho mouse red 1:250
goat anti-rat 1gG Alexatluor 488 Green 1:250
In this experiment, antibody of even-skipped was analyzed. This antibody gives a nice banding
pattern as shown below (shown below for interactive fly website). It has a molecular weight of
about 38kDa. A different secondary antibody had to be used here. Texas Red antibody was left
out and unknown if still useable so Rho CO66 Red was used. It was also not known if the scope
used can detect outside the Texas Red spectrum. Rho red is pinker in color then Texas Red and
higher in the spectrum of the scope. It was noted when looking under the scope that the chorion
was not removed giving notion that the bleach was old. The slide for this experiment was thrown
out since the chorion needed to be removed in order to have proper absorption of antibody thus
another experimental trial was preformed.
Image 8&9: Even-skipped immunoflorscent analysis of embryos. Note the chorion still around embryo
Image 10: Interactive fly website
picture of even-skipped band
patterns.
Experiment 8:
This experiment was a repeat of experiment 7. Outcome that we wanted to see was the stripping
pattern on the embryo. The experiment slide had dried out over a long break. This can be due to
the use of a large cover slip. This cover slip allowed for drying of the plate and this no images
have been obtained.
Experiment 9:
Primary 1:
• Mouse Sir2 1:25
• Rat alpha tublin 1:250
Primary 2:
•
•
Rat alpha tublin 1:250
Mouse anti even-skipped 1:250
Secondary
•
•
CO66 Rho mouse red 1:250
anti-rat 1gG Alexatluor 488 Green 1:250
The last experiment was to see if changes in secondary antibody and time would help express the
antibodies in the desired pattern of even-skipped and Sir2. Sir2 had the secondary antibody of
Texas red in experiment 6 then changed to Rho Red. The time variant for this experiment was
also changed. It was tested to see if primary antibody can be left for only two hours and not
overnight while secondary was left overnight and not for two hours. It was concluded that Sir2
banding was still not present in the images. Even-skipped showed a slight pattern but was not
conclusive enough to get a result from. It is still not known though if timing can have a greater
affect and more experiments will have to be done with this at a later time.
Image 11,12&13: Even-skipped results
from immunoflorscent studies.
Image 14: Sir2 results of
immunoflorscent study
Based off of the experiments completed there is still a lot more experiments that need to
be done in order to obtain the desired results. Some future experiments that can help us in this
search would be to: have a variant in antibody concentrations, embryonic staining techniques for
fly development, secondary dilution methods, different antibody variants, testing of HRP & AP
with western blots (answering the overall question of Can we make this work?), and other
techniques and tools to bring to a lab setting on the bases of making it easier to teach to students
quicker and easier. The experiments that were done can be the building blocks toward future
developments for this objective. The use of Drosophila embryos will act as the backbone of all
developments and the key will be through the methods of both Immunofluorescence and western
blotting.
Drosophila melanogaster & Modern
Protocol Analysis for the Classroom Setting
By
Amelia Irene Dosio
A Thesis Submitted to the Graduate
Faculty of Rensselaer Polytechnic Institute
in Partial Fulfillment of the
Requirements for the Degree of
MASTER OF SCIENCE
Major Subject: BIOCHEMISTRY & BIOPHYSICS
Approved:
___________________________________
Dr. Donna E. Crone
Thesis Advisor
Rensselaer Polytechnic Institute
Troy, New York
November 2010
(For Graduation December 2010)
i
Copyright 2010
By
Amelia Irene Dosio
All Rights Reserved
ii
CONTENTS
LIST OF TABLES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v
LIST OF FIGURES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . vi
ACKNOWLEDGMENT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . viii
ABSTRACT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .ix
INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1
1.
Drosophila is a Good Model Organism for High School Classroom. . . . . . . 1
2.
Embryonic Development in Drosophila. . . . . . . . . . . . . . . . . . . . . . . . . . . . . .3
3.
Studying Aspects of Drosophila Development. . . . . . . . . . . . . . . . . . . . . . . . 5
4.
Drosophila Embryonic Development Requires Even-Skipped. . . . . . . . . . . .9
5.
Assaying for Eve Expression in the Classroom. . . . . . . . . . . . . . . . . . . . . . . 11
METHODS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .12
1.
Fly Maintenance and Handling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ..13
1.1
Fly Vials/Bottles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ..13
1.2
Apple Juice Agar Plates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .13
1.3
Fly Cages. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14
1.4
Embryo Collection. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .15
2.
General Reference Strain Embryo Fixation. . . . . . . . . . . . . . . . . . . . . . . . . .15
3.
General Immunofluorescence of Reference Strain Embryos . . . . . . . . . . .16
4.
Imaging of Reference Strain Embryos . . . . . . . . . . . . . . . . . . . . . . . . . . . .. 18
5.
DNA & General Cell Staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18
RESULTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20
iii
1.
2.
Visible Stains. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20
1.1
Coomassie Blue. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .20
1.2
Trypan Blue. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22
1.3
Nile Blue A. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23
1.4
Methylene Blue. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .24
Immunostain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25
2.1
Block 1: Modification of Embryo Fixation and Primary Antibody
Incubation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28
3.
2.2
Block 2: Secondary Antibody. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30
2.3
Block 3& 4: Post Antibody Processing and Imaging. . . . . . . . . . . 30
Test of Modified Protocol by Students. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33
CONCLUSION/DISCUSSION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35
1.
Visible Stains. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35
1.1
Coomassie Blue. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .35
1.2
Trypan Blue. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37
1.3
Nile Blue A. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37
1.4
Methylene Blue. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .38
2.
Immunostain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .38
3.
Considerations for Classroom Use of Laboratory Protocols. . . . . . . . . . . . 40
REFERENCES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .42
iv
LIST OF TABLES
1.
Common Abbreviations Used In Text. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .12
2.
List of Buffers and Percent Composition of Solutions . . . . . . . . . . . . . . . . . . . . .. 12
3.
Table of Primary and Secondary Antibody Combinations and dilution used in
experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17
4.
Antibody and Reagent Source . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .17
5.
List of DNA &Cell Stains Used in Experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . . .19
6.
Flow Chart for Three 1.5 Hour Time Blocks for Modified Immunofluorescence
Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26
v
LIST OF FIGURES
1.
Stages of Embryonic Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4
2.
Drosophila Embryogenesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4
3.
Nuclear Division Cell Cycle of Wild-Type Embryo 2 Hours after Fertilization . . . . 5
4.
Different Layers of Drosophila Embryo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6
5.
Activation of the Even-Skipped Gene at during Development . . . . . . . . . . . . . . . 9
6.
Band Labeling for an Embryo Treated with Even-skipped . . . . . . . . . . . . . . . . . . 10
7.
LacZ Reporter Assay of Regulatory Elements Required for Stripped Pattern of
Even-Skipped Expression.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11
8.
Embryos Stained with Bio-Rad Fast Blast DNA Stain. . . . . . . . . . . . . . . . . . . . . . .21
9.
Embryos Stained with Bio-Rad Fast Blast DNA Stain. . . . . . . . . . . . . . . . . . . . . . .21
10.
Embryos Stained with Bio-Rad Fast Blast DNA Stain. . . . . . . . . . . . . . . . . . . . . . . 21
11.
Embryo Stained with Sigma-Aldrich Trypan Blue Stain. . . . . . . .. . . . . . . . . . . . . 22
12.
Embryo Stained with Sigma-Aldrich Trypan Blue Stain. . . . . . . . . . . . . . . . .. . . .22
13.
Fixed Embryo Stained with Carolina Biological Nile Blue A. . . . . . . . . . . . . . . . . .23
14.
Embryos Stained with Sigma-Aldrich Methylene Blue. . . . . . . . . . . . . . . . . . . . ..25
15.
Embryos Stained with Sigma-Aldrich Methylene Blue. . . . . . . . . . . . . . . . . . . . ..25
16.
Fixed Embryo Placed in Methanol as a Control for Comparison to Other Visible
Stains. . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . .. . . . 25
17.
Embryo Processed with Original Immunofluorescence Protocol and Control
Antibodies Viewed At 20X Under Fluorescence. . . . . . . . . . . . . . . . . . . . . . . . . . 27
18.
Comparison of Embryos with Primary Antibody Reuse. . . . . . . . . . . . . . . . . . . . .29
vi
19.
Embryo Processed with Modified Immunofluorescence Protocol and Antibodies
of: 1o Rat Anti-Alpha Tubulin and Mouse eve Even-Skipped. . . . . . . . . . . . . . . . 31
20.
The Seven Stripes of The Protein Encoded eve Gene In A Developing Drosophila
Embryo. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .32
21.
Embryo Processed with Modified Immunofluorescence Protocol and Antibodies
of: 1o Rat Anti-Alpha Tubulin and Mouse Anti-Even-Skipped . . . . . . . . . . . . . . . .33
22.
Embryo Processed with Modified Protocol By A Control Antibodies and
Modified Immunofluorescence Protocol by Cell and Development Student At
RPI. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34
vii
ACKNOWLEDGMENT
I would like to thank my thesis advisor Professor Donne E. Crone, Ph. D. for her
help and guidance throughout my research. This thesis would have not been possible
without her guidance and broad knowledge of Drosophila research. I would also like to
thank Professor Eric M. Rutledge, Ph. D. for his help and guidance with biochemistry
techniques that broaden my research abilities to relate them to my Drosophila research.
I would like to show my gratitude to all the undergraduates and graduates whom I have
met during my research studies in the teaching and research labs. Thank you for
allowing me to share my research experiences with you and to exchange the knowledge
that was learned in and out of the classroom with great respect. All your help has
enabled me to write this thesis today with greater appreciation for Drosophila. I would
also like to thank my family and close friends for giving me the support through the
rough times that I have faced throughout my undergraduate and graduate years at
Rensselaer. If it wasn’t for these people my studies wouldn’t have gotten this far and I
greatly appreciate of all the help and guidance that I have received. For all others that I
may have forgotten that deserve to be acknowledged I am sorry and would like to say
thank you for all your help also
viii
Abstract
Drosophila is a model organism that gives many advantages that include the ease
of visualization of embryonic development through protocols and techniques such as
DNA staining and immunofluorescence. In research protocols, a significant amount of
time and materials are used to observe the aspects of interest in the common fruit fly.
In this study, protocols for Drosophila studies were tested and modified to fit the time
constraints and chemical use restrictions of an advanced placement high school
classroom. Modification to these protocols will enable the study of development while
keeping the interest of the students, preserving the scientific aspect of the study and
making the protocols more economical. With the use of a reference strain,
modifications to embryonic analysis protocols were generally successful. Modifications
of the protocols will enable Drosophila use in the high school level setting where time
and material constraints are a general factor.
ix
Introduction
1. Drosophila is a Good Model Organism for High School Classrooms
Drosophila melanogaster, the common fruit fly, is a prime model for viewing
embryonic developmental stages using light microscopy. Drosophila has a long history of
use in the areas of development, behavior, aging and evolution. Thomas Hunt Morgan,
Nobel Prize winner, studied Drosophila’s phenotypic variations from genotype
mutations. By mating white-eyed male with a red-eyed female, he was able to invent
heredity. Heredity allows genes on the chromosome to be expressed in later
generations of Drosophila (Krock, 2001). Drosophila’s swift development is a favorite
among geneticists and is a popular organism studied for its vast developmental aspects.
Drosophila is an excellent organism to use in a classroom setting for students of all
grade levels to study development. Drosophila’s short life cycle also makes it an ideal
organism for classroom study setting since time restraints are a big factor (Ashburner,
Hawley & Sullivan,2000).
Drosophila is not only used in basic research but is a common teaching tool.
Protocols from all aspects of developmental biology, in Drosophila, are used to study a
number of human neurological diseases such as Parkinson and Alzheimer disease. Many
protocols can help to screen for chemical compounds linked to these diseases and have
the potential to prevent or to reconstruct the disease for termination in later mutations.
While there are numerous research protocols that focus on Drosophila embryogenesis
in detail, modifications need to be done to adapt protocols for the classroom setting.
1
Not all protocols can be adapted for many reasons which include cost, efficiency and
safety.
Genetic crosses are often at the core of Drosophila research but it requires the
skill of collecting virgin female flies. Companies have developed kits that enable
experiments to become a little quicker with protocol modifications that can be used for
teaching. Such kits include a monohybrid cross inquiry-based kit that allows more time
for in-depth observation and analysis of phenotypes. This kit, from Carolina Biological,
doesn’t require a collection of virgin flies for controlled crosses. Students are able to
study a monohybrid inheritance using wild type and an “Easy Fly” sepia Drosophila
strains. A heat shock treatment prior to shipment of flies allows fly lines to only produce
female virgins ready to cross with wild type males (“Carolina Easy Fly Monohybrid Cross
Kit”, 2010).
Drosophila protocols are modified to target key aspects that one wants to study.
In a high school classroom setting time is a big factor; in a research setting
experimentation can be done over longer blocks of time. Attempting to use protocols
already published can cause teachers to stray from using Drosophila for developmental
studies due to this time constraint. The key factor to minimizing effect of time constraint
would be to modify established protocols. These modified protocols would be designed
for different classroom settings, a more economical approach based on what materials
and equipments are readily available and, most importantly, time restraints. Visible
staining of cells, embryonic imaging and immunostaining are some basic techniques that
2
can be modified to bring to an advanced high school classroom. These techniques will
not only allow students to learn the aspects of development with Drosophila under high
school scheduling time constraints but will prepare them for a college lab class.
Techniques brought into a classroom setting not only need to be important scientifically
but will need to be visually pleasing to establish and maintain student excitement.
2. Embryonic Development in Drosophila
Embryogenesis takes place after fertilization. The process starts as mitosis or
cellular division in the embryo. In the early stages of Drosophila embryo, the first 9
nuclear divisions occur but the cytoplasm does not divide resulting in a syncytial
blastoderm. This blastoderm is multinucleate and centered in the center of the embryo.
The cytoplasm plays a role in gradient formation that influences body pattern formation
in later embryo stages. At the 10th nuclear division, the nuclei move from the center to
the posterior portion of the embryo and cellularization occurs forming a cellular
blastoderm. At about 10 hours after fertilization, the major body segment boundaries
become defined. After 16-20 hours post fertilization, the embryo hatches to yield a
larva. Larva, pupa and adult stages have head, thoracic and abdominal segments that
were derived from segmentation formed during embryogenesis. (Alberts et al.,
2007).Figures 1&2 show the stages of embryogenesis. Figure 1 shows an embryo in
lateral view in different stages of development. Endoderm midgut, mesoderm, central
nervous system and pole cells are shown at different stages. Figure 2 shows an embryo’s
3
journey through embryogenesis with emphasis on the formation of nuclei, cellular
blastoderm and body segments (“Drosophila Development”, 2010).
4
3. Studying Aspects of Drosophila Development
Drosophila embryogenesis is well studied. The ease of preparation of embryos
and the collection of a large number of embryos makes Drosophila an attractive system
to study in the classroom. Numerous strains are available for investigation of normal
and altered embryogenesis. Early stage embryos, provide a series of nuclear divisions
that establish a certain pattern as a normal part of development. Embryos at a variety
of developmental stages can be stained with a variety of DNA stains including
DAPI/Hoechst 33342, 7AAD (7-Aminoactinomycin D) and propidium iodide and then
analyzed under a fluorescent microscope. Nuclei would be too small to be viewed under
a dissection microscope. Figure 3 shows a textbook image of early nuclear division of a
Drosophila embryo stained with propidium iodide, a nuclear stain.
The embryo has a shell like protection barrier known as the chorion. When doing
experiments using Drosophila embryos, the chorion must be broken with chemicals and
reagents in order to allow reagents to penetrate and interact with cellular components
5
for later detection under a fluorescent scope. Figure 4 shows the outer chorion and
internal layers of a Drosophila embryo.
Once the chorion has been removed, embryos can be fixed at the stage of
development prior to cellularization as in figure 3. Once an embryo is fixed, the second
protected layer known as the vitelline membrane must be removed. Removing the
membrane must be done quickly since the membrane can trap the embryo and cause
later optical obstructions when viewing under the microscope (Ashburner, Hawley &
Sullivan, 2000). Once protective layers are removed, embryos can be dehydrated in
methanol and stored over a long period of time. The use of stockpiled fixed embryos can
be used to shorten protocols and techniques in later times. Dehydrated embryos need
to be rehydrated in aqueous solution for a number of downstream applications
including DNA staining and immunostaining. Increasing proportions of methanol and
PBS are used to rehydrate embryos from an alcohol solution to an aqueous solution.
Once rehydrated, embryos are ready to be processed. Embryos can also be nuclear
stained with Hoechst dye simultaneously with antibody (Ashburner, Hawley &
Sullivan,2000).
6
More specific visualization of cellular components can occur with the use of
antibodies. Immunochemistry uses antibody to detect a specific antigen or protein
which is located in a cell. In order for an antibody to access the interior of a cell, there
must be holes in cell membrane. This occurs during the fixation and processing steps in
Drosophila embryo fixation and is why a detergent is added to solutions. A blocking
solution of BSA and PBT is used to coat the embryo in protein and prevents non specific
binding of antibodies.
Antibodies to cellular components are called primary antibodies. Most are not
able to be detected directly; detection requires a second step. Secondary antibody is
necessary for the detection of primary antibody through fluorophores, an enzyme linked
or a radiolabel. If primary antibodies were generated in mouse then the secondary
antibody that detects mouse IgG should be used. Fluorophores are light sensitive and
bleaching can occur if left under ultraviolet light for an extended period of time (Asai,
2008).
Antibodies can be at low concentrations but not too low to have nonspecific
binding. When using secondary antibody tagged with fluorescent molecule, tiny
amounts of antigen can be detected. The molecule is excited by specific wavelength of
energy and then emits photons at a lower wavelength to give a fluorescent image.
Alternatively, the use of enzymatic labeled secondary and colorimetric staining solutions
developmental aspects can be seen without fluorescence. By tagging with enzymes that
are visible under bright light visualization under a fluorescence bulb can be eliminated.
7
Immunofluorescence is a powerful technique. Visualization of fluorescently
labeled cells or embryos requires specialize equipment; in particular, a fluorescence
microscope is needed. A microscope with fluorescent capabilities cause these
techniques to become expensive, however, microscopes can be shared and borrowed
from a university that is in partnership to a high school classroom to help ease the cost.
In immunofluorescence, primary and secondary antibodies need to be chosen to
carefully to double label and observe two proteins simultaneously. Similar principle
would apply for the use of enzyme labeled antibody. When double labeling, one must
chose two primary antibodies that were generated in different animals and secondary
antibody with distinct fluorescent properties. For example, double labeling can be
examined with fluorescein (emission at 519nm) and Cy5 (emission at 670nm) channels
in a fluorescent microscope (Ashburner, Hawley & Sullivan, 2000). Filters are available
that separate emission spectra that for fluorophores allowing them to be both seen.
For these studies, primary to alpha and beta tubulin (generated in rat and
mouse respectively) of antibody was available. Actin and beta-tubulin are cytoskeletal
proteins present in large quantities in a cell allowing them to be controls. In a classroom
setting, these controls would enable students to compare with other antibodies specific
to that of developmental affects.
8
4. Drosophila Embryonic Development Requires Even-Skipped
Even-skipped (eve) encoded 376 amino acids protein (EVE). That is a marker for
pattern formation at the cellular blastoderm and the development of a small neuron set
(Brody, 1998). Even-skipped is a pair rule gene; pair rule genes send signals to cells to
make 14 abdominal parasegements and have a role in head formation. Figure 5 shows
the activation of even-skipped gene at different stages of fly development.
At the early stages of development eve is expressed at very low levels. In later
stages, a higher level of expression is seen as a single broad band located near the tip of
the embryo. This band can be detected before nuclear division of the nuclei is complete.
Approximately 3 hours after fertilization, the band narrows and becomes dense in cells.
This first band becomes classified as A1 and the bands that follow will appear broad and
undeveloped. At approximately 4 hours after fertilization bands A3, A5, A7, A9, A11 &
A13 narrows and becomes asymmetric. Figure 6 shows classification of bands in an
even-skipped treated embryo. Positioning of the stripes is a big part of development
through their functioning of growth in the embryo (Lawrence, 1992).
9
Mutation of eve causes every other segment of the embryo abdomen
development to eliminate a certain part of the cuticle pattern. Larvae that are
homozygous for weak alleles lack the even abdominal segments. These segments are
classified as A2, A4, A6, A8, A10, A12 and A14 and the general pattern formation can be
accounted for cell death (Lawrence, 1992).
Control of eve expression has been studied in detail using reporter genes and
upstream DNA fragments from the eve gene (Lawrence, 1992). These pieces of DNA
(putative eve control regions) were placed in front of a promoter and a lacZreporter
gene. Reporter genes that were constructed were inserted into the embryo and then
transformed into an adult fly. Transgenic flies that express beta galactosidase when
expression occurs from the promoter of interest. For eve expression, it was found that
700 base pairs of upstream DNA must be linked to the promoter to have regulated
10
expression for stripe formation. Figure 7 illustrates how the regions required for the
different eve stripe formation in the lacZ reporter assay.
5. Assaying For Eve Expression in the Classroom
Stripped pattern of eve expression has important developmental affects as
shown by phenotype mutants (Lawrence, 1992). Eve is an excellent protein of study in
the classroom. Observation of eve expression in the classroom, not only shows
developmental affects on an embryo, but it can also preserve the interest of students
with its unique banding pattern. Modified protocols for Drosophila study within time
constraints of a high school laboratory can bring new aspects to biology studies. These
protocols will enable students to have broader exposure to research based material and
allow them to become prepared for challenges in advanced educational and industry
settings.
11
Methods
12
1. Fly Maintenance and Handling
1.1 Fly Vials/Bottles
Flies were cultured in 6oz Polypropylene square bottom bottles and narrow
Drosophila vials with Droso-Plugs (Genesee Scientific, San Diego, CA) containing 1.5oz 3.0oz of Formula 4-24 plain Drosophila medium (Carolina Biological, Burlington, NC).
Medium was rehydrated with an equal amount of distilled water following the
directions from the supplier. During the 1-2 minutes of the hydration process, the flies
were anesthetized with carbon dioxide and a Foot Valve Complete System (Genesee
Scientific, San Diego, CA). A sprinkle of yeast was added to the bottles and the flies were
then placed into the containers. A range of approximately 10-25 flies were placed in
each vial to ensure a good male to female ratio. Flies were kept at 25oC in an incubator
or at room temperature with 12 hour light cycle. Vials and bottles were changed every
two weeks.
1.2 Apple Juice Agar Plates
Protocol was modified from Drosophila Embryo Preparation Protocols
(Ashburner, Hawley & Sullivan, 2000). In a 1000mL bottle, 750mL of distilled water
(dH2O), 27.0 grams of agar with a stir bar were added and the solution was autoclaved
at 121oC for full cycle in an autoclave (Tuttnauer 3870E). After autoclave, 250mL of
sterile apple juice or grape juice was added. To prevent fungal growth, a 0.05%
Tegostept (Apex) stock solution (1gram Tegosept dissolved in 20mL of ethanol) was
added once solution was cooled to 45-50oC. Solution was poured into 60mm X 15mm
13
petri dishes filled half way and allowed to solidify on the bench top. Plates were stored
at 4oC until use.
1.3 Fly Cages
An apple juice agar plate was obtained and a 2mm drop of yeast paste was
placed into the center of the plate. Yeast paste was made from 10 grams of baking yeast
(Fleischmann) with 6mL of water. Mixture was stirred for three minutes and allowed to
sit at room temperature for 20 minutes. Mixture was stirred again until desired viscosity
was obtained and stored at 4oC.
Small (60mm) embryo collection cages (Genesee Scientific, San Diego, CA) were
used to collect embryos on apple juice agar plates. Flies were anesthetized with carbon
dioxide and added to cages from fly bottles and vials. To do so, cages was placed upside
down on the carbon dioxide pad and anesthetized flies were added to cage. An apple
juice agar plate was placed at the bottom of the cage and a secure cap ring was placed
over plate to lock plate in place. Cage was flipped over into an upright position and kept
at 25oC in an incubator or under a lamp with a fluorescent bulb at 12 hour cycles. Plates
were changed daily or hourly depending on desired embryo collection. After 10 days,
flies were disposed of by placing them in a -20oC freezer. After 1-2 hours flies were
placed in trash.
14
1.4 Embryo Collection
Protocol was modified from Drosophila Embryo Preparation Protocols
(Ashburner, Hawley & Sullivan, 2000). An apple juice agar plate containing collected
embryos were harvested. The drop of yeast was removed with a kimwipe avoiding as
many embryos as possible. Embryos were rapidly transferred from the apple juice agar
plate to a 50mL beaker using distilled water and a fine point paint brush. A 10mL
solution of 50% bleach was placed into the beaker containing the embryos and swirled
for 3 minutes at room temperature. To ensure chorion removal, bleach solution was
made fresh before each collection. Embryos were strained through a 100uM nylon cell
strainer (BD Falcon, Sparks, MD) placed on top of a 250mL flask then rinsed with a small
amount of 0.7%NaCl/ 0.1% Triton X solution to remove bleach. Embryos were then
rinsed with a 5mL of dH2O and strainer was then placed on a kimwipe to slightly dry the
embryos.
2. General Reference Strain Embryo Fixation
Protocol was modified from Drosophila Embryo Preparation Protocols
(Ashburner, Hawley & Sullivan, 2000). Dechorionated embryos were transferred into a
scintillation vial containing 10mL of heptane using a small natural bristle paint brush.
Embryos were allowed to sit for 30 seconds to allow for heptane permeation and 2.5mL
of 4% formaldehyde solution was then added to the vial. Embryos were allowed to fix
for 20 minutes. After fixation, solution was removed and embryos were washed twice
with fresh heptane. Once washed, between 5-10mL of heptane was added to vial
15
depending on amount of embryos obtained. An equal amount of 100% methanol was
quickly added and vigorously shaken for 3 seconds. Fixed embryos were in the
interphase between methanol and heptane layers. When removal of both heptane and
methanol layers were completed, a rehydration series of methanol and PBS steps
followed for 2 minute durations. Rehydration series included: 90% methanol/10%
1XPBS, 75% methanol/25% 1XPBS, 50% methanol/ 50% 1XPBS, 25% methanol/75%
1XPBS. Fixation process was completed with two rinses in a 1X PBS/0.1% Tween 20
solution and placed in a microfuge tube for downstream applications.
3. General Immunofluorescence of Reference Strain Embryos
Protocol was modified from Drosophila Embryo Preparation Protocols
(Ashburner, Hawley & Sullivan, 2000). Embryos hydrated in PBT were resuspended in
one pipette volume of 1% BSA in PBT for blocking. Embryos were placed on a rocker at
room temperature for one hour. After removal of blocking solution, 250uL of primary
antibody solution was added and allowed to incubate overnight on a rocker. After
incubation, the used (also called preabsorbed) primary antibody was removed and
placed in a separate microfuge tube for later use. Antibodies were reused 4 times and
then replaced with new antibody. Embryos were washed in PBT in three increments of
five minutes each. After the final wash, PBT was then removed and embryos were
incubated with 250uL of secondary antibody for two hours on a rocker in a dark room.
Table 3 indicates primary and secondary double labeling combinations and dilutions
16
used in this protocol. Table 4 indicated sources and protein concentrations of primary
and secondary antibodies used.
After incubation, embryos were washed with PBT in three increments of 5
minutes and then replaced with 100uL of Hoechst stain at a 1:5000 dilution. Hoechst
stain was allowed to absorb for 10 minutes on a rocker to nuclear stain. Hoechst dye is a
DNA stain that is excited by ultraviolet light with blue light emission at 460-490nm.
17
Hoechst 33342 can stain of nuclei in a fixed or living embryo by binding to the adeninethymine region in the minor groove of a DNA strand. Embryos were then rinsed and
washed with PBT for another three increments of five minutes. To prepare embryos for
mounting, all traces of PBT were removed and embryos were washed twice with one
pipette volume of 100% methanol for 5 minutes. Methanol was removed and 100uL of
mounting solution (Sigma,USA) was added to the embryos.
4. Imaging of Reference Strain Embryos
Protocol was modified from Drosophila Protocols (Ashburner, Hawley & Sullivan,
2000). Four small dots of clear nail polish were placed onto the microscope slide in an
arrangement of a square for embryo placement. A 20uL drop of embryo mounting
solution mix was placed into the center of the dots with placing a cover slip over the
bubble of solution. A thin perimeter of nail polish was placed around the cover slip to
adhere it to the glass slide. Images of embryos were taken with a Zeiss Axio Observer Z1
compound research microscope with Imaging Solution AxioVision Rel 4.6.3.0 software.
This compound microscope transmits light and epifluorescence. Microscope has DAPI,
GFP-FITC and Cy3-TRITC filters.
5. DNA & General Cell Staining
Microscope slides (3” X 1” X 0.1mm) were covered with parafilm. Two small
squares about 5mm X 5mm were cut out of the parafilm. Parafilm was flattened against
the microscope slide using another microscope slide to allow parafilm to adhere to slide.
Embryos processed from fixation step were used for this technique with 20uL of
18
embryos in 100 % methanol placed in each square opening and 25uL of DNA stain added
to embryos. The embryos were allowed to sit for a period time depending on stain used.
Once embryos were stained, they were washed three times in 100uL of distilled water at
room temperature. Embryos were viewed under bright .Table 5 indicates stains and
dilutions used.
19
Results
Initial studies were undertaken to find a method for colorimetric staining of
Drosophila embryos. A series of slides for embryogenesis was originally sought but no
company was found to offer a series of such slides. Colorimetric staining was found to
be an easy and inexpensive technique. If the protocol can be adapted it would be a good
tool for high school students. Many staining protocols were available and a few were
chosen for use on Drosophila embryos. Dyes were chosen based on the availability and
reported staining protocols found.
1. Visible Stains
1.1 Coomassie Blue
Drosophila embryos were incubated in DNA stain with combinations shown in
Table 5. All embryos were all collected at early stages of development (1-2 hours post
fertilization) to concentrate on cellular development. Coomassie Blue (“Fast Blast DNA
Stain Instruction Manual”, 2010) is used in biology as a safe alternative to ethidium
bromide to detect DNA in agarose gels. Fast Blast uses positively charged dye molecules
to attract and bind to negatively charged dye phosphate groups in DNA molecules. Fast
Blast stain will stain DNA a deep blue in gels (“Fast Blast DNA Stain Instruction Manual”,
2010). Fast Blast was used to stain fixed embryos to see if this safe alternative to using
ethidium bromide gels to detect DNA was possible to bring into a high school classroom.
Embryos were stained and diluted in methanol or hydrogen sulfate, as suggested by
supplier, with destaining both in distilled water. Figures 8,9 &10 show embryos stained
under different conditions with Fast Blast DNA stain and visualized at 20X and 40X using
20
bright field. As shown, embryos were stained but were not sufficiently destained to
show sufficient developmental detail.
21
1.2 Trypan Blue
Fixed embryos were next stained in Trypan Blue. Trypan Blue is excluded from
live cells. Trypan Blue is typically used for cell viability. When the membrane of a cell is
compromised there is an uptake of dye into the cell. Living cells are impermeable to
Trypan Blue yielding clear and opaque (unstained) cells. When cells are nonviable they
appear dark blue in color (Long-Sorbello, Saydam, Banerjee & Bertino, 2006). For these
experiments, Trypan blue is used for general staining properties. Embryos were stained
at 1:10 dilution with methanol and destained with distilled water. Some embryos were
not destaining to observe the concentration of Trypan Blue present in the cells of the
embryo. Figures 11 &12 are images taken at 20X and 40X of Trypan Blue stained
embryos. The embryos took up the dye but there was not sufficient destaining leaving
the embryo to be black in color and lacking in visible detail.
22
1.3 Nile Blue A
Nile blue A is a stain that is used in biology as a lysosome detector (“Nile blue A”,
2010). Nile blue A colors the lysosome red. The solution preferentially favors lipid
staining in the solution as a red color. Overall Nile blue A stains the nuclei and
background blue in contrast and the lipids red when mixed in sulfuric acid solution
(Llewellyn, 2005). Since sulfuric acid is known as a corrosive acid and not recommended
for a high school lab setting, an alternate substrate was used. Fixed embryos were used
with Nile Blue A dissolved methanol, a less toxic solution, to see if staining of the nuclei
and or lipids could be detected. Figure 13 shows an embryo treated with Nile blue A in
methanol and viewed at 40X under an compound scope with bright field. Treated in this
manner, Nile Blue A appeared to stain the entire embryo blue to black. Since methanol
is an alcohol and not an acid it is unlikely to have the same effect as was described in
the original protcol. Acetic acid would have been a better choice over methanol and
sulfuric acid.
23
1.4 Methylene Blue
In the final attempt at staining embryos with visible dye fixed embryos were
stained with methylene blue. Methylene blue is used for qualitative examination of RNA
and DNA present in a cell (“Methylene Blue Stain”, 2010). Methylene blue is a substitute
for ethidium bromide when visualizing nucleic acid in gels or directly staining of an
embryo. Fixed embryos were stained with methylene blue diluted 1:30 in methanol. A
higher dilution ratio of methanol to Methylene Blue was used since the stock solution
was at a high concentration of stain. Stock solution was 60% methylene blue/ 40%
distilled water. As shown in Figure 14 &15, embryos treated with methylene blue in
methanol and viewed under bright field at 20X appear blue in color with no detectable
features. In contrast Figure 16 shows a control embryo placed only in methanol and
viewed under bright field. This control (unstained) embryo was used for comparison to
all stains. Since stains were found to be unsuitable to enable visualization of the
developmental aspects of an embryo, a new set of protocols were tested. The new
approach was to screen a series of antibodies via immunofluorescence to find ones that
would be suitable for classroom use to highlight the developmental aspects of
Drosophila embryos.
24
2. Immunostain
Immunofluorescence microscopy is commonly used in cell biology labs. Existing
protocols were designed for research labs to optimize results. However, in research labs
workers work devote a full day to lab work. Modifications to the immunofluorescence
protocol were made to bring the experiment time frame to better fit a high school
classroom setting. In the research setting immunofluorescence of Drosophila embryos
would take a period of two days. The first day of experimentation takes approximately 4
25
hours with an overnight incubation and the second day takes approximately 6 additional
hours of processing to complete the protocol. The protocol has been modified to span
over three 1.5 hour blocks, a typical double period in a high school setting, with time
between blocks varying 1-3 days. Table 6 shows a flow chart of the three 1.5 hour
blocks.
The original protocol taken from Drosophila Protocols (Ashburner, Hawley &
Sullivan, 2000) was used as a control for comparison to the modified protocol. Primary
antibodies of mouse anti-actin and rat anti-alpha tubulin diluted at 1uL:250uL with
26
1%BSA/PBT and secondary antibodies of goat anti-rat IgG Alexafluor 488 and goat antimouse IgG Texas Red diluted at 1uL: 250uL with 1%BSA/PBT were also used as controls.
Since actin and tubulin are present throughout the embryo and their presence and
localization is well established its makes excellent control for compare to antibodies that
give different expression patterns. Figure 17 A-D shows an early stage embryo
processed with the original immunofluorescence protocol and control antibodies. Red
stain in the embryo indicates actin localization, blue stain indicates DNA stained with
Hoechst dye and green stain indicates tubulin. An image overlay was also included to
observe where proteins were localized with fluorescence.
27
2.1 Block 1: Modifications of Embryo Fixation and Primary Antibody Incubation
Modifications to the immunofluorescence protocol were taken in a series of
small blocks. In the embryo fixation, the amount of fixation chemicals added was varied
based on the amount of embryos obtained on the apple juice agar plate and the time
needed to fix was cut by 30%. The addition of heptane and 4% formaldehyde amounts
was cut by 50% to 5mL of heptane and 1.75mL of 4% formaldehyde in a scintillation vial.
The time for fixation was also cut from 20 minutes to 15 minutes. This elimination of
over 50% of the hazardous chemicals in this step allows for less chemical waste and the
decrease of possible hazards to the students in a classroom setting. By also reducing
the amount of heptane and methanol in the removal of the vitelline membrane by 50%
also eliminates waste. However, this step is unlikely to be permitted in a high school
classroom so the technique of stockpiling fixed embryos for use in immunofluorescence
was tested.
Ashburner etal. noted in Drosophila Protocols (Ashburner, Hawley & Sullivan,
2000) that stockpiling embryos in methanol can be done after removal of the vitelline
membrane and dehydration of the embryos was complete. This protocol was tested and
stockpiling of embryos was done over a period of three weeks with collection and
processing of embryos daily. All embryos used in this experiment were not kept more
than 5 months with this stockpiling technique due to high turnover and use. When
ready for immunoassay, embryos were rehydrated. Rehydration of the embryos was
kept as the same as the original protocol and the blocking step with 1%BSA/PBT for an
28
hour was maintained in original form. The original protocol called for an overnight
incubation in primary and this was used as a stopping point for block one. At this point,
embryos can remain in antibody at 40C for several days.
Primary antibody was found to be able reused for a range of 7-8 times before
antibody recovery rate was too low for use (figure 18). This was compared to the normal
4 times of use (Blackman, 1999) Too little antibody should not cause background
reactivity but may make it difficult to see specific signal. Since antibodies can be
expensive (Sigma alpha tubulin at $200.00 for .5mL), primary antibodies are often
reused. Ability to reuse antibody is highly dependent on the primary antibody and the
dilution at which it is used also. Reusing primary antibody depended on the antibody
being used and the combination of proteins being tagged. Figure 18 A&B show a
comparison of reference strain embroys treated with primary antibody that was resued
4(A) and 8(B) times. It was seen that with the prolong use of primary antibodies made it
is difficult to see specfic signaling of eve.
29
2.2 Block 2: Secondary Antibody
Second experimental block of the modified protocol consisted of removing
nonspecific bound primary antibody followed by secondary antibody incubation.
Modifications to this process were done to the washing steps. Primary antibody was
removed and placed in a microfuge tube to save for later use. Embryo wash times were
reduced from 5, 10, and 20 minutes in PBT to three washes in PBT at 5 minutes each.
This cut the total washing times from 45 minutes to only 15, a 70% difference for each
washing step. In most embryo immunofluorescence protocols, DNA is stained to provide
an additional visible marker. Staining with Hoechst 33342 was kept the same as the
original protocol of Drosophila Protocols (Ashburner, Hawley & Sullivan, 2000).
Secondary antibody incubation was changed from two hours at room temperature to an
overnight incubation at 4oC on a rocker to end day two. In theory secondary antibody
can be used over again but the secondary antibody was not reused in these
experiments.
2.3 Block 3 & 4: Post Antibody Processing and Imaging
Block three of the modified protocol consisted of removing secondary antibody,
washing, mounting and imaging of the embryos. This part of the experiment was kept
the same as the original protocol except depending on the availability of different
microscopes to view embryos. Figure 19 A-D shows a late stage embryo processed with
the modified immunofluorescence protocol. Primary antibodies of rat anti-alpha tubulin
and mouse anti eve protein and secondary antibodies of goat anti-rat IgG Alexafluor 488
30
and goat anti-mouse IgG Texas Red were used in the testing of the modified protocol.
All antibodies were diluted at 1uL:250uL with 1%BSA/PBT. Red signal indicates eve
protein localization; blue signal indicates DNA and green signal indicates alpha tubulin.
An image overlay was also included to observe overlapped of localization. The presence
of eve protein is shown on the outer edges of the embryo indicated by arrows in the
figure below. Eve protein is known to be expressed in a striped pattern across the
embryo. In these images, eve protein bands can only be seen on the outer edges. Figure
20 shows a textbook image of eve expression. Typically, generated textbook images use
a confocal microscope to show banding pattern.
31
The modified protocol was repeated with a different red fluorescent secondary
antibody (replacing goat anti-mouse IgG Texas red with donkey anti-mouse Rho-TRITC)
using the same primary antibodies. Introducing this variation provided a test of the
modified protocol but maintained visualization of eve protein using red fluorescence.
Figure 21 A-C shows a later stage embryo processed with the modified
immunofluorescence protocol using mouse anti-even-skipped primary antibody donkey
anti-mouse IgG Rho TRITC secondary. Red signal indicates eve protein localization; blue
signal indicates DNA stained with Hoechst. An image overlay was also to make the
banding pattern of eve distinct. Arrows indicate strong banding of eve expression.
32
3. Test of Modified Protocol by Students
Modified protocol was tested by students in a Cell and Development Biology
class at RPI. Even though these students were at the college level, none of the students
had previous experience with Drosophila. By allowing these students to test the
modified protocol, it helped to assess the protocol from a “non expert” point of view.
Figure 22 A-D was produced by one group of students using primary antibodies of
mouse anti-actin and rat anti-alpha tubulin and secondary antibodies of goat anti-rat
IgG Alexafluor 488 and goat anti-mouse IgG Texas Red and compared to control group
obtained in previous experiments. Red stain in the embryo indicates actin localization,
blue stain indicates DNA stained with Hoechst dye and green stain indicates tubulin. An
image overlay was also included. The student embryos tend to be very bright, probably
33
due to less than optimal image capture settings (figure 22), but show common signal
expression to that of the control embryos (figure 17).
34
Conclusion/Discussion
Before the start of these experiments, the goal was to generate a set of slides of
Drosophila embryo development that could be used for reference in the teaching
laboratory. None of the typical scientific suppliers had Drosophila embryo development
slides (stained or unstained) although slides for larvae to adult flies where found (for
example, Carolina Biological). The initial focus for this thesis was to find a stain and
protocol that could yield series of slides for embryogenesis of the Drosophila embryo.
More than 20 were available in the laboratories that were intended for a variety of cell
to SDS gel stains. Protocols were looked up in a variety of textbooks and recommended
staining techniques from the supplier. We decided that four preliminary stains and
techniques would be used to attempt to develop a set of teaching slides on Drosophila
embryogenesis. When these attempts were unsuccessful, a different approach was
taken. Instead of creating a new protocol, modifications were made to existing
Drosophila protocols that have been successfully used to visualize developmental
aspects of the embryo. Protocols and other visible stains maybe looked at in greater
detail in future studies to see if a successful set of slides can be obtained.
1.Visible Stains
1.1 Coomassie Blue
Coomassie Blue stained the entire embryo dark blue in color even after
destaining for 5 and 10 minutes (figures 8 & 9). While staining with Coomassie Blue it
appeared that embryos seemed to become weak around the outer membranes after
35
chorion and vitelline membrane removal. Many of the embryos broke open near the
center and caused leakage of cells around the embryo. These embryos were deemed
unusable. Additional chemicals used processing prior to Coomassie stain such as bleach,
heptanes and methanol could have aided in the weakness of the embryo wall. The weak
outer membrane of the dechorioned and fixed embryos may have cause Coomassie Blue
to enter the entire embryo and cause staining throughout. Even with the use of
methanol to dehydrate the embryo and shrink the membrane to allow less staining,
embryo was stained in its entirety. Even with the addition of water to destain and
rehydrate for imaging, embryo remained entirely stained with a large concentration of
stain at the center of the embryo. Embryos stained with Coomassie blue diluted in
hydrogen sulfate had the same outcome as that of the embryos diluted with methanol
(figures 8&9 versus figure 10).
It was concluded that perhaps a better technique can be used in future studies
to destain the embryos to reduce the overall staining of the embryo. A variation to
decrease the amount of stain taken up would be to dilute the stain at 1:1000 or higher
with longer destains (overnight) or longer destain in larger volumes. Another possible
technique to stain the embryo could be to not remove the chorion or the vitelline
membrane. Embryo would be stained directly after collected from fly cages and then
processed through the fixation method to removed membranes and then observe if this
method aids in staining the embryo without breaking the delicate membranes.
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1.2Trypan Blue
Trypan blue was easily able to enter fixed embryos but was inconclusive in aiding
differentiation of the intracellular structures. Staining covered the entire embryo
making it impossible to distinguish developmental features (figures 11&12). Trypan blue
indiscriminately stained the embryo since the fixed embryo has all “dead” cells. Embryos
were dark blue to black in color when not destained and with only a slight difference
when destained for 5 minutes or 10 minutes (figure 11 &12). After the 10 minutes of
destaining, Trypan blue was pooled to the center of the embryo as a black color while
the rest of the embryo was a slight dark blue color. While destaining with distilled
water, embryos seemed to clump and stick to one another and to the microscope slide.
The uneven staining may have been due to improper washing. If Trypan blue were to be
tried as a general stain in the future a diverse list of solutions would be needed to test if
destaining with other possible solutions would cause the embryos to stick together or
cause a pooling of stain to the center of the embryo. Also increasing the dilution of the
Trypan blue stains could tested.
1.3 Nile blue A
Nile blue A was found to be unsuccessful in staining the nuclei and lipids in the
early stage embryos. This result was not unexpected since the solvent in the staining
was changed from an acid (in the original protocol) to an alcohol. The color of Nile blue
A in stock solution was very rich in blue color. With the addition of 1uL of Nile blue A to
10uL methanol, it was noted that the diluted solution was as dark in color as in the
37
original bottle. After destining, the embryos were a dark blue to black color and
developmental aspects were not discernable (figure 13). To change this outcome, future
studies could use a much higher dilution of stain. In addition, instead of changing the
solvent to an alcohol a less acidic acid could be used. Another possible solution can be
to change the destaining method. Destaining the embryos in larger volumes for longer
times and/or changing the destaining solution frequently during destaining may, help
the overall staining of the embryo and give a better possible outcome.
1.4 Methylene Blue
The last visible stain tested was methylene blue. This also yielded unsatisfactory
results. Methylene blue is reported to stain DNA (“Methylene Blue”, 2010). Staining
with methylene blue yielded similar results to Nile blue A of dark blue embryo (figure
17). Having a dilution ratio of 1:30, it was hypothesized that staining would occur at a
less intensely compared to the original solution. After destaining the embryo, embryos
were remained a dark black in color. The embryos seemed to clump together in the
distilled water destaining solution. When viewed under the microscope, it was apparent
that some of the embryos were not stained at all while others were completely stained.
2. Immunostain
Since it was proving difficult to develop a protocol for generating teaching slides
with embryos with visible dyes, new approach was taken. This new approach was to
modify existing protocols for Drosophila embryo immunofluorescence, a well known
technique. Drosophila embryo immunofluorescence protocols were successfully
38
modified to enable them to be used in a classroom setting. The modified protocol not
only brought the experiment time frame to better fit a high school classroom setting but
it also enabled visualization of the developmental aspects of the embryo. This was in
contrast to the poor resolution seen with the visible stains that were tested. When an
antibody to a developmentally important protein is used, the modified
immunofluorescence protocol yields results that show developmental aspects of the
embryo. The control of actin and tubulin allow for comparison to the development of
important eve protein.
The binding of eve antibody was visualized with the secondary antibodies labeled
with the fluorophore Texas Red or rhodamine/TRITC. The unique banding pattern eve
expression was seen in embryos about 8 hours after fertilization (figure 20). All
parasegments of eve expression are seen on the outer edges of the late stage embryo
(figure 19 & 20). Based on published data the expected stripes should be seen to span
the entire embryo but published data is often obtained using more sophisticated
techniques (such as confocal microscopy). Confocal images which enable imaging at
multiple planes would show the full striping pattern of eve (figure 20) (Alberts, 2002).
When imaging with a compound microscope, the embryos are relatively thick so you are
only able to see the concentration of eve protein at a single plane. At the edges of the
embryo, the entire section is in the focal plane enabling visualization. Different stage
embryos can also be treated with the even-skipped antibodies to observe the complete
cycle that eve makes.
39
3. Considerations for Classroom Use of Laboratory Protocols
A major advantage of modifying the Drosophila embryo immunofluorescence
protocols is in limiting the amount of hazardous chemicals used in the classroom for
student health and to limit the amount of waste being put into the environment. Being
able to stockpile embryos by a teacher can allow for these benefits. A question comes
about on how long these stockpiled embryos can be kept. A future study could examine
the usable life span of stockpiled embryos the time frame used here (less than 8 months
after fixation and dehydration).
In a classroom setting, commonly used solutions can be cross contaminated due
to high volumes of use. Small amounts of solutions should be made or placed to smaller
bottles for use and thus one must allow for a little longer preparation time. Different
secondary antibodies can be used depending on availability of antibodies and much
more importantly what filters are available on the fluorescence microscope. In order to
use the technique of immunofluorescence, you must have access to a fluorescence
microscope. Fluorescent microscopes are costly (around $2500), but used microscope
may be obtained for less via source such as Ebay only buying one or two microscopes,
cost might not prohibitive. Alternatively, there are secondary antibodies that have
enzyme tags of alkaline phosphatase and/or HRP which yield chromogenic results when
incubated with the proper substrate. Now that antibody and protocol development
have been modified for fluorescence the next step would also be to work out protocols
for use with secondary antibodies of alkaline phosphatase and HRP.
40
The modified Drosophila embryo immunofluorescence protocol was tested by
students in a Cell and Development Biology class with positive results. These college
students had no previous experience with Drosophila, yet they achieved good results
with the modified protocol. In addition two students in the Cell and Developmental
biology class, tested a protocol for colorimetric detection of enzyme (lacZ) using a
special transgenic fly line. These students showed that although colorimetric techniques
are less sensitive than fluorescence, they could visibly detect regions where beta
galactosidase was expressed (Lee and Rubel, personal communication). This result
suggests that the use of enzymatic tagged secondary antibodies should be feasible to
make immunolabeling of Drosophila embryos even more accessible to high
school/college students where; visualization requires simple compound microscope
found in many high school biology classrooms.
In general, this thesis has shown that modifications to Drosophila protocols can
be used in a high school classroom setting. By careful selection of antibodies the
developmental aspects of Drosophila can be observed. Results that are important
scientifically and also visually interesting spark and keep the interest of young and
growing minds. Additional variations of the immunolabeling protocol may increase its
utility in the classroom. Many biology classes use Drosophila to show genetics. This
thesis provides a protocol to enable exploration of cell and development biology as well.
41
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