The Making of
Transgenic
Drosophila guttifera
Made possible by
Team Spot 21
Written by Thomas Werner
Drosophila guttifera
1 General notes about D. guttifera
Drosophila guttifera is a very rare mycophagous species which is native to the Midwest of the USA. In
the lab, it can be reared on fresh (maximum 2 weeks old) sugar food with some additional sprinkles of dry
baker’s yeast. As a rule of thumb, the generation time and sexual maturation time of D. guttifera are about
50% longer as compared to D. melanogaster. The egg production rate of D. guttifera is one to two orders
of magnitude lower than that of D. melanogaster. In a typical injection experiment, the first flies hatch
about 19 days post injection.
Drosophila guttifera is a very demanding species in terms of breeding conditions. If breeding parameters
are slightly off the optimum, the culture can be lost rapidly. It is important to know the three big enemies
D. guttifera. They are 1) Contaminations with D. melanogaster females, which can produce enough
offspring to out-compete the slower-growing D. guttifera larvae, leading in the worst case to a complete
nonappearance of the next D. guttifera generation. 2) Mold. Especially during spring and summer, there is
a high risk of green mold infestations of freshly started bottles, which can exterminate the culture if not
discovered early. 3) Ethanol. It is important to know that ethanol is lethal to Drosophila guttifera. In
conditions of poor aeration, the ethanol fumes produced by active yeast can kill the flies within a few
hours.
D. guttifera cultures are best kept at room temperature in a place where a regular visual inspection
(especially for fungus growth) can be done conveniently. D. guttifera eggs dry out very much faster than
D. melanogaster eggs. The larvae tolerate wet conditions, while pupae and flies prefer a moderate
humidity.
Whenever any D. guttifera fly work is done, all surfaces and tools have to be cleaned thoroughly with
anti-fungus spray (defined later). High quality work is the key to keep a D. guttifera stock alive, while
sloppiness will result in the loss of the culture.
2 Two culture systems of D. guttifera
Two independent D. guttifera culture systems are needed: One stock-maintaining culture and one egglaying cage. The stock-maintaining culture takes 1 year to establish from a handful of starter flies and has
to be maintained and kept healthy at all times. The egg-laying cage can be put out of function when
micro-injections are not planned within the next months. However, it is advisable to keep the egg-laying
cage populated as a backup system whenever possible.
The stock-maintaining culture
The stock-maintaining culture has to be large to compensate for the low egg-laying capacity of the
females when micro-injections are done, and to keep a solid backup of healthy flies when fungus
infections hit some bottles of the stock. The basic principle is to have four groups containing 20 culture
bottles each, which are 0-1, 1-2, 2-3, and 3-4 weeks old, respectively. Once a week (on a fixed weekday),
20 sugar food bottles are taken out from the cold room and are allowed to warm up, evenly spaced and
not touching each other (to prevent the formation of condensation in the bottles). The bottles are then
supplied with some sprinkles of dry baker’s yeast and a piece of anti-fungus paper stuck between the
glass wall and the food. Now, the emerged flies from a 3 week old bottle are distributed into 3 freshly
prepared bottles. The remaining 17 fresh bottles are populated by combining the flies from the 1 week old
bottles. If the flies in the 3 week old bottles haven’t hatched yet, use a 4 week old bottle instead. The 20
oldest bottles are kept for one more week to allow more flies to hatch, which can be used to either repopulate the egg-laying cage (see next chapter) or to collect virgin flies for crosses.
As fungus growth usually starts out from the bodies of dead flies, it is advisable to empty out ALL flies
from the one week old bottles (especially the dead ones!).
The egg-laying cage
The egg-laying cage is made of Plexiglas and allows good aeration through the nylon entrance at the front
and the plastic mesh at the back of the cage. Aeration prevents the accumulation of toxic ethanol fumes
produced by the yeast. The cage measures 300 x 200 x 200 mm and holds tens of thousands of flies.
Before a clean cage can be populated, six small (35 x 10 mm) plastic Falcon Petri dish bottom halves are
taped upside-down (using double-sided tape) onto the bottom of the cage. They will serve as “tables” for
the bigger sugar food plates and have to be spaced out evenly. Medium-sized empty dishes can be used to
see if the spacing is good. Without these “tables”, the sugar food plates would smash lots of flies on the
ground with every food exchange.
Feed the flies in the cage three times a week with six sugar food plates at a time, which are medium-sized
(100 x 15 mm) Petri dish bottom halves, filled to the top with sugar food. Right before feeding the flies,
add some dry baker’s yeast to the top of the food, and wipe off any free water from the plates. When
replacing old against new plates, follow the three rules below to minimize the killing of flies:
1) Bump off the flies before taking out a plate. Use the thumb and middle finger to hold the dish, while
holding the index finger in place to prevent the food from falling out.
A) If the gap between the food and the Petri dish is large due to excessive drying of the food, then
bump towards the large gap. The food won’t move much downwards and no fly will be trapped.
B) If the gap is narrow, bump towards the side where the food still sticks to the plastic. This
prevents most of the flies in the gap from being squeezed to death.
2) One out, one in! Always take only one plate out, then put one fresh plate inside. If there are flies on the
food “table” in the cage, they can be pushed away with the front edge of the fresh food plate when putting
it down. Then take the other plates out and replace them one by one.
3) Discard the used food because flies developing from it will be of poor quality and a source of fungus
infestations.
An egg-laying cage can be in operation for up to 10 weeks before the flies have to be transferred to a new
cage. If fungus grows (pay particular attention to places where the food comes in contact with the wall),
change the cage instantly. Perform the following steps to move the flies to a new cage:
1) Take out all sugar food plates, close them and continue working at a fly bench with CO2 gas.
2) Cover an area with unfolded paper towels to your right and put a 5 L beaker on it.
3) Have a new cage with a fresh entrance ring plus nylon ready.
4) Put the populated cage to the left of the new cage (not on the towels), the entrance facing upwards.
5) Open the CO2 gas, lead it through the nylon to anesthetize the flies, and then close the CO2 gas supply.
6) Remove the ring with the nylon of the old cage and pour the flies into the 5 L beaker.
7) Flies which fall on the towels can now be thrown into the beaker as well.
8) Gas the old cage again to blow loose the remaining living flies, and shake the cage over the beaker.
9) Pour the flies from the beaker into the new cage.
10) Close the freshly filled cage, bring it to its designed place in the lab and supply the flies with food.
11) Clean the old cage and the nylon with tap water. Do not use soap or bleach. Rinse with distilled water.
3 Drosophila guttifera injection
General remarks
D. guttifera females lay comparably few eggs within a given period of time. For this reason, a large egglaying cage filled with thousands of flies is used to obtain at least 200 - 300 freshly fertilized eggs per
hour. The cage is kept on the lab bench with a lamp placed 1 m above it which is timed to turn on at
6 AM and off at 6 PM. Females need to be reared on fresh food with dry yeast at all times in order to
make their ovaries develop, and to achieve a continuous egg production. If the food is in poor condition,
females will retain their fertilized eggs inside of their bodies and lay them when the embryos are too old
for micro-injections. The cage has to be supplied with fresh food three times a week and especially one or
two days before injections are carried out. Eggs which are laid in the morning are often already
cellularized and tend to have a thicker corona, which is disadvantageous for injections. The egg quality
increases towards the afternoon. The best time to start the egg-laying is 11 AM. The eggs are to be
harvested in 1-hour cycles, starting at noon. It is advisable that one person lines up 2 slides (200 embryos)
during a 1-hour cycle, while another person injects them. All wash steps of the embryos are done with
distilled water from the tap. Since this water turns quickly hot, 10 liters should be collected the day before
and stored at room temperature. Never wash Drosophila guttifera eggs with ethanol, because it seems to
harden the chorion and prevents the injection needles from penetrating it. The Skilcraft cellulose sponges
used to collect the eggs can withstand autoclaving and treatment with anti-fungus spray.
Egg collection and preparation for microinjections
A starter yeast paste has to be made and activated one or a few days before using it, and it has to be mixed
again with additional dry yeast and water in the morning of the injections. The yeast stays at room
temperature during the injections and is only good when it smells like fermented fruit. At 11 AM, all
sugar food plates in the cage are replaced by four moist Skilcraft cellulose sponges, which are to be
placed into medium-sized (100 x 15 mm) Petri dish bottom halves and covered with a thin layer of fresh,
active yeast paste. A second set of four sponges will be needed. The best way to prepare the sponges is to
first make them too wet and then to carefully squeeze off the excess water with two thumbs pressing and
moving from the top to the bottom throughout the surface of the sponges, until no drops are running out
easily. The sponges should be dry enough that no water runs out when flies are bumped off from them,
but wet enough to supply the flies with enough liquid. Sponges which are too dry seem to result in thicker
coronas of the eggs, while sponges which are too wet cause the accumulation of liquid in the cage. The
yeast is then supplied evenly over the sponge surface. Care should be taken when yeast is handled,
because contaminating flies will be quickly attracted to the site. The sponges are to be placed very close
to the wall inside the cage but without touching it or squeezing the flies. The females in the cage are
allowed to lay eggs on the 4 sponges for one hour. The flies prefer to sit on the sponge part which is
closest to the wall. Sponges placed too far away from the wall are often ignored by the flies.
To collect the eggs, the sponges are one by one removed from the cage and replaced by fresh sponges
with yeast paste. The eggs-containing sponges are gently squeezed out one by one in a 2 L glass jar with
distilled water (room temperature!). After all 4 sponges are squeezed out and while the eggs sink to the
bottom of the jar, the sponges and Petri dishes are washed with distilled water and prepared for the next
round. To avoid contamination, wash and reassemble in the following order: One bottom half, one
sponge, one lid, and so on. The sponges should be squeezed hard when washing them in order to kill the
possibly remaining eggs inside of them. After all 4 sponges are washed, put the yeast paste on them and
store them in a drawer (away from contaminating flies) until they are used again.
Now the first half of the egg collection water can be poured right into the sink. The second half is swirled
and poured through a double filter system, consisting of two fine-meshed embryo collection baskets, each
stuck to a funnel. The finest filter is sitting underneath and retaining the eggs, while the groove filter
above is trapping sponge fibers and flies. The glass jar is then rinsed once with 100-200 ml of distilled
water which is also poured through the filter system. With a squeeze bottle of distilled water, wash the
eggs of the upper filter and allow them to slip through into the filter below. After obtaining a sufficient
amount of eggs in the lower filter, remove the basket from the funnel and wash the eggs thoroughly with
distilled water from a squeeze bottle. Then put the basket with the washed eggs into a bottom half of a
small Petri dish, which is half way filled with distilled water. Line up the embryos under a dissection
scope.
A VWR micro cover glass (18 x 18 mm) is now placed onto a VWR micro slide (25 x 75 mm, 1 mm
thick) with a small droplet of water so that it sticks. Dry off the edges with a piece of tissue. The washed
embryos are now transferred with a medium-sized white haired brush onto the cover glass and kept wet at
all times, while only the white and structureless appearing embryos are lined up and also kept wet at all
times. It is advisable to put a clump of more than 100 embryos into the upper middle of the cover glass.
Then use a very fine brown haired brush, make it wet, quickly dry it on a tissue paper, pull the first
embryo into position, dry the brush, pull the next embryo to extend the line, and so on. The first 10
embryos shall be moved with a relatively dry brush, or otherwise the surface tension will make it
impossible to position them correctly. Move the clump of embryos down while progressing downwards
with the line. In the end, the row of embryos consists of about 100, which shall have their posterior end
towards the right edge of the cover glass, and the row of embryos should be positioned about 5 mm away
from that edge. Keep the embryos moist at all times by supplying water with the brush to the clump of
unaligned eggs and also to the top, middle, and end of the row. When the row is completed, remove all
unused eggs and let the row dry while carefully monitoring the eggs under the microscope. The row can
now be rearranged slightly U-shaped to make them fit better to the sugar food surface later on. When the
embryo line appears completely dry, wait about 3-5 more seconds and then cover them quickly with the
Halocarbon oil mixture (7 parts of type 700 + 1 part of type 27), supplied by a syringe. The Halocarbon
oil stops further desiccation and allows breathing. If the embryos become too desiccated (oil added too
late), they die, and it becomes visible when the injection needle penetrates the chorion but not the plasma
membrane at the first try. If, however, the oil is put on too early, then the embryos won’t stick to the glass
and move around when the injection needle is poked against them. Only a few seconds lay between both
extremes.
DNA preparation for microinjections
The DNA is prepared with the Qiagen Midi kit, eluted with 5 ml QF buffer, precipitated with 3.5 ml isopropanol, centrifuged and re-dissolved in 350 L distilled sterile water, precipitated again with ethanol,
washed with 70% ethanol, dried, and dissolved with EB buffer to a final concentration of 1 g/l. The
DNA is centrifuged for 20 min at maximum speed and a mixture of the clear DNA solution is prepared as
follows: 20 L construct, 5 L helper plasmid, and 15 L sterile Milli-Q water. The mixture is
centrifuged again for 20 min at maximum speed, and 39 L are transferred to a fresh tube for immediate
use.
The piggyBac system (Wimmer and Horn, 2000) results in 1 transformation event every 50 D. guttifera
embryos with empty vector, and 1 every 500 embryos with an 8 kb insert. The exact plasmid names are:
“pBac{3xP3-EGFPafm}” (piggyBac backbone) , and “phspBac” (piggyBac helper).
Needle preparation, microinjections, and heat shock of the embryos
We use a Flaming/Brown micropipette puller Model P-97 and FHC capillary tubing (Borosil 1.0 x 0.75
mm ID/Fiber with Omega dot fiber) to obtain standard needles (program 46). Extreme care should be
taken in order to avoid damage to the platinum heating filament of the machine. The needles are loaded at
least one hour before using them with approximately 0.5 L DNA mix and can be stored in a moist
chamber at 4°C for a few days. We use an automated injection system based on nitrogen pressure, a
micro-manipulator for the needle, and an inverted microscope. The tip of the needle is broken by carefully
touching the chorion of an embryo while repeatedly pushing the “Clear” button until the DNA flows out.
Embryos are injected with a very fine needle tip and either an automatic stream of DNA into the very
posterior part of the embryo or a push (if necessary) by hitting either the “Clear” button (when the needle
is still very fine) or the foot pedal (when the needle is wearing out). In order to prevent the needle from
getting plugged, the “Clear” button or foot pedal should be hit frequently while being outside of the
embryos. Also, moving the embryos away from the needle at high speed can help to make the DNA
flowing again. When using the foot pedal, adjust the amount of microseconds on the machine between 25
and 4 microseconds to maintain a constant injection volume. The needles have to be changed when the
embryos start leaking out granulous cytoplasm, whereas clear droplets are normal.
For effective transformations, the eggs may not have undergone cellularization yet. Any cellularized
embryos should be left alive to prevent fungus growth on the leaking cytoplasm of killed cells.
Wipe off the oil: After all 100 embryos on a slide are injected, remove the cover glass (a droplet of water
can help) with the embryos and quickly wipe off some excess oil against the micro slide. Stick the cover
glass into a fresh sugar food vial, positioning the posterior ends of the embryos right above the food.
Close the vial with a cotton ball and squeeze some drops of distilled water between the vial and the cotton
to provide immediate moisture. Place the vial vertically into a wet chamber (a beaker with a little bit of
water on the ground and 2 wet paper towels stuck against the inner wall). Hold multiple vials together
with a rubber band. Avoid water flowing into the vials by keeping them away from the wall. Close the
chamber with aluminum foil and store it in the 18°C incubator.
The next morning (16 – 18 hours after injection), fill the wet chamber with 42oC warm water to a height
that covers the food and heat shock the embryos at 37°C for 120 minutes. The heat shock should happen
with the lid half way lifted so that air can circulate through the chamber. After the heat shock, empty the
chamber completely (including the paper towels), fill it with some cold water and put the vials in it to
lower the temperature in the food. After some minutes, discard the cooling water, and fill the beaker with
about 1 cm of distilled water (no paper towels required), put the vials back in, place the lid back on and
store the embryos in the 18°C incubator for an additional 4 days.
After-care and fly crosses
When all pupae have formed and no more wandering larvae are seen, collect the pupae with a brush from
the food and with two pairs of forceps from the cotton plugs. Making the cotton very wet helps to
dissolve the glue with which the pupae stick to it. Do not leave much cotton left on the pupae, but a few
fibers are allowed to remain attached to them. Collect the pupae temporarily on a wet tissue. Take a clean
and empty glass vial, place a long piece of anti-fungus paper along one side of the wall and moisten the
paper with a wet brush (distilled water). Transfer the collected pupae with a moist brush from the wet
tissue paper onto the anti-fungus paper in the vial. Close the vial with a fresh cotton ball and store it
horizontally in a moist chamber so that the pupae lay on the anti-fungus paper. Moisten the anti-fungus
paper inside the glass vials when necessary.
Start collecting wildtype virgin flies of both sexes for back-crosses when you see the first pupae coming
up from the injection experiments. The collected virgins are stored in sugar food vials with some dry
baker’s yeast and have to be bumped to fresh vials every 3 or 4 days. They will be sexually mature when
your first injection survivors hatch.
The hatched flies from the injections are collected once a day in the afternoon on a CO2 pad and crossed
with wildtype virgins of the opposite sex. Use sugar food vials without adding extra yeast and combine 3
males or 10 females of the injection survivors with 10 wildtype virgins of the opposite sex in one vial.
The combining is done to produce enough larvae to keep microorganisms in the culture down. Bump the
crosses to fresh food every 3-4 days.
EGFP screen in the larval eye-disks
Look for EGFP in the eye disks, but it is often even visible in the labial disks, in the brain, and sometimes
in the genital disks. The expression is usually strong and clearly visible in all larval instars. Perform the
screen after all larvae are hatched, which is usually the case 3 days after the parents have been removed
from the vial. EGFP can be detected latest in white pupariums. Multiple positive larvae per vial are
common. Also, many lines are obtained in later screening sessions, so it is worth the time bumping the
crosses for 2 or 3 weeks until declaring them as negative. The larval EGFP screen is carried out in the
dark room as follows:
1) Switch on the UV lamp 10 minutes prior to screening.
2)
- Remove the larvae from the food vials with distilled water (squeeze bottle) and a white-haired
brush, and pour them into a medium-sized Petri dish bottom half.
- Collect positive larvae with a pair of forceps into 2 mL tubes filled with 1 mL sugar food.
- Discard water with negative larvae into an empty 5 L beaker.
3) Microscope settings: Light filter = “GFP”; Bottom filter = “Oblique”; Objective =1 x; Shutter open.
4) Switch on the red light and switch off the top light.
5) Clean up and DO NOT FORGET TO TURN OFF THE UV LAMP.
Recipes
Sugar food (Double anti-fungus action)
Add 79 g agar to 8500 mL boiling water, let dissolve, add 275 g ICN baker’s yeast, 520 g cornmeal,
110 g granulated sugar, and 20 g Sorbic Acid. Stir until boiling, turn off the heat and stir for 15 min. Add
23.8 g Tegosept dissolved in 91.8 mL 95% ethanol, mix and pour 50 mL into each bottle and fill Petri
dishes up to the top.
Anti-fungus paper towels
Dissolve 2 g of Sorbic Acid and 0.6 g of Methyl Paraben in 200 mL of 95% ethanol. Roll 20 - 30 paper
towels and stick them into a 1 L glass beaker. Slowly pour the anti-fungus solution over the towels to
soak them evenly. Unfold the towels and let them dry completely. Anti-fungus paper does not go bad and
can be stored for years.
Anti-fungus bench spray
Dissolve 2 g of Sorbic Acid and 0.6 g of Methyl Paraben in 50 mL of 95% ethanol and add 70% ethanol
to a final volume of 1 L.
Literature:
Horn, C. and Wimmer, E.A. (2000) A versatile vector set for animal transgenesis. Development Genes
and Evolution. 210, 12, 630-7.
The egg-laying cage.
Putting yeast paste on a sponge.
Eggs in the lower filter.
Putting oil on lined-up embryos.
Embryo and injection needle.
Wet chamber for injected embryos.
Collecting pupae from a cotton plug.
Pupae in a moist chamber.
A transgenic D. guttifera larva.
Transgenic pupae reveal DsRed spots.
Adult wing.
Pupal wing: vein spot CRE (EGFP), and intervein shade CRE (DsRed).