Get an ice bucket. Fill with ice. Remove competent cells from -80C freezer and thaw on ice 5 min. The competent cell you will be using is from the “In-fusion” kit. Add 50 uL of competent cell to a 1.5 mL microcentrifuge tube.
Remove agar plate from the cold room and let it warm to room temperature by placing it in lab bench or putting it in the 37 C warm room
Get DNA from from plasmid box
Quickly spin it down in centrifuge to collect all of the DNA into the bottom of each tube prior to use. A quick spin of 20-30 seconds at 8,000-10,000 rpm will be sufficient. Note: There should be 50 µL of DNA in each tube sent in the Kit and you will only be using 1 uL of it.
Add 1 uL DNA to competent cells and mix gently (i.e. by tapping tube with finger).
Incubate on ice for another 20 min
Heat shock at 42C for 45 seconds,Be careful to keep the lids of the tubes above the water level, and keep the ice close by.
Immediately transfer to ice for 2 minutes
Add 1 mL LB OR SOC media to each transformation tube as you take it from the ice
Incubate in 37C shaker for 30 min
Plate 150 microliters on LB amp plates (or corresponding antibiotic). If DNA is from ligation reaction, centrifuge transformation rxn tube for 3 min at 8.5g (approx. half of max speed of centrifuge). Remove ~1 mL of supernatant (such that ~250 mL of supernatant remains in the tube). Resuspend pellet by pipetting up & down. Finally, plate as normal.
Incubate overnight at 37C (ideally for 14-18 hours)

Fly Lab Procedure

Knocking Flies Out with CO2

You will observe that there is piping on the wall, with a black turn valve that is pointing to the right by default. This piping is connected on the right side to a upturned bottle of water, with a knob on the side and a gauge on top. Begin by turning the black turn valve counterclockwise, from pointing to the right to pointing upwards. Then, adjust the knob on the right side until the gauge reflects a value of around 7-10. This represents a good concentration of CO2 to knock flies out.

Then, you will notice a syringe with a trigger, usually to the left. Direct the CO2 syringe down along the cork/cap, into the bottle. Turn the bottle upside down (so that flies fall onto the lid), and squirt CO2 into the bottle in 2-3 bursts, or until the flies mostly stop moving. Then, remove the lid and shake/tap unconscious flies onto the CO2 pad. This pad is porous and pumps CO2 through, so flies will remain unconscious while on the pad.

Starting New Bottles of Stock Flies

Begin with a previous bottle of stock flies. Knock flies out (see above), and observe under microscope. Sort males and females into two separate piles, using a brush to move them without causing any damage. Males typically have darker and more curved abdomens, such that the stripes terminate more early. Females will have slightly lighter stripes that go all the way to nearly the end, and more straight abdomens. Select 5 males and 5 females for each bottle. Keep the bottles on their sides, and using a brush, scoopula, or folded cardstock to pick up flies, deposit flies onto the side wall of the bottle. This is to keep flies from getting stuck in the food.

Keep bottles on sides for a few minutes, until flies regain consciousness and begin moving. Then, flip bottles upright, and label the sides of the bottle with “iGEM” and the date, to identify the stock bottles.

Fly Dissection

Begin with a dissection petri dish, which appears as a petri dish with a thick layer beginning halfway through. This has a soft/elastic bottom which will allow you to safely pin flies to the bottom without doing too much damage. Take a tube of phosphate-buffered saline (PBS), and two pairs of tweezers. If putting fly gut onto glass slides, see below for relevant materials.

In dissection room, uncover the dissection petri dish. Create a puddle of PBS on an edge (around size of a quarter, or a bit more). Put petri dish under microscope, if you have not done so already. Then, take a bottle of stock flies. Knock flies out (see above), and deposit onto CO2 pad. Pick up chosen fly using tweezers, and submerge into PBS puddle. Move your petri dish and adjust zoom/focus on microscope until you can see the fly quite closely and sharply in focus, when pinned at the bottom of the dish.

Begin the dissection process. First, note the appearance of various parts of the abdominal organs. Most notable are (1) The gut. Appears long and is somewhat randomly curled up, segmented. Translucent and white, usually. (2) Malpighian tubules. Very thin strands, often beaded and distinctly yellow. Usually surrounding other organs. (3) Testes. Long, very tubular (with no segments). Often, translucent with a yellow tinge. Curled up like a snail (and will maintain this shape). Easy to mistake for the gut, but note the yellow tinge that is distinctive, as well as the well-defined shape, and confusing the two should be avoidable. (4) Ovaries. Large and white, hard to mistake for much else. Will explode into eggs all over plate if accidentally disturbed. Are not easily confused for the gut.

There are multiple approaches to dissection. One is: (1) Tear off the head and thorax, being careful that no gut accompanies or is torn in the process. Then, attempt to pull the exoskeleton on the abdomen apart, so that the innards are exposed. Keep doing so until you observe something that may resemble the gut, which may be clumped together with other parts. Gently pull organs apart until the gut is separate from other organs. (2) Tear off the head and thorax, as above. Then, gently squeeze abdomen, as most of contents are the guts and will squeeze out. Then, gently pull organs apart. More may be added as different things work for different people; anything can be valid, as long as you try to avoid damaging any organs and try to get as complete of a gut as possible.

If you will be putting gut onto glass slides, try to stretch the gut out to a long linear tube (instead of curled up), so that it may be observed/imaged more easily subsequently.

Putting Fly Gut onto Slides

First, do all steps in fly dissection above. Bring along a tube of glycerol and a pipette, as well as glass slides and slide covers. Once the gut has been separated from the fly, use the pipette to put a droplet of glycerol onto the glass slide. Then, use tweezers under the microscope to pick up the gut and deposit it into the droplet of glycerol. You may attempt to visually confirm the location of the gut in the droplet, then put on the slide cover and again visually confirm that the gut is present and its location is known to you.

Making Arabinose Stock

Weigh out 2g Arabinose and add to 10mL water (or, scale down if you have less Arabinose).

Feeding Flies

Similar to Starting New Bottles of Fly Stock. Begin with a previous bottle of stock flies. Knock flies out (see above). Count out the best flies with a brush, and scoop onto brush/scoopula. Put feed tube to its side to avoid flies falling into the liquid and drowning, and transfer flies into feed tube. Gently push flies down into the tube with a brush so that they aren’t squished by the reinsertion of the cork/lid, then put the lid back in. Keep tube on its side until flies awaken.

Putting Flies into Insect Cages

You may put whatever food or liquid you need into the cages. If you put petri dishes in, keep the lid on, push the dish through the tubular entrance, place the dish down, and remove the lid through the entrance. Then, gently tap incapacitated flies onto the floor of the cages, being careful not to crush them. Tie the entrance into a knot, and then put the cages into the constant-temperature chamber.

Removing Flies from Insect Cages

Remove insect cages from constant-temperature chamber. Put on nitrile gloves, and hold onto a folded piece of cardstock and a brush with one hand. Gently unknot the tubular entrance to the cages, and stick the hand with the cardstock and brush into the cage. Use your other hand and hold the cardstock, or put the cardstock on the floor of the cage. Then, use the brush to collect or sweep up dead flies into the folded cardstock. This is important to be able to distinguish living flies from dead ones after putting in the cold room. Remove cardstock and brush from cage, and making sure that no flies are trapped in the entrance tube, tie it back up into a knot. Count dead flies if necessary, then put flies into trash.

Put insect cages into cold room for 5-10 minutes or until all of the flies are immobile in their habitats. When you remove them from the cold room, they will be on the floor of the cages, and you may once again unknot the entrances, remove a few to dissect or whatever other procedure you need, and tie the cages back up, and put back into the constant-temperature chamber.

Team:Princeton/Experiments

Princeton

Protocols

E. Coli Transformation Protocol