Goode Lab Media and Undergrad Jobs (a.k.a., The Media Bible)
Last updated: 4/26/05
I.
Lab Jobs and Descriptions
1. Autoclaving (general)
2. Glassware/plastics
3. Culture tubes
4. Glass pipets
5. Pipet tips (for Pipetman)
6. Glass beads
7. Wooden dowels and toothpicks
8. Squeeze bottles
9. Velvets
10.Changing soak bins/waterbaths
11.Other assorted lab jobs
II.
Making and Pouring Plates
1. Pouring and testing plates (general)
2. LB (and antibiotics)
3. YPD
4. Dropout media
5. Other plates
III.
Making Liquid Medias and Lab Stocks
1. LB
2. YPD
3. Dropout media
4. Sugar solutions
5. Amino acid solutions
IV.
Other stocks
1. Amino acid powders
2. Frozen stocks
a. Antibiotics
b. Other stocks
Section 1. Lab jobs and descriptions
Using the autoclave
There are four autoclaves on the second floor. The left-most autoclave is broken. You
will probably use the middle two autoclaves most frequently, but you may also use the
very large manual autoclave for big batches of items or if other autoclaves are occupied.
General notes:
 Use as little autoclave tape as possible! It is expensive.
 Make sure that the chamber pressure is at 0 before you open any autoclave.
Ignoring this item is guaranteed to leave you with a nasty steam burn.
 If someone else has left their items in the autoclave, you may remove them and
place them on the shelves on the right side of the room (or the counter if they are
too large/unwieldy to put on the shelves). Be nice with other people’s things,
since you would want them to be nice with yours!
 Make sure that the chamber door is closed all the way (as tight as you possibly
can turn the knob) before beginning an autoclave cycle.
 Dry items may be stacked on top of one another inside the autoclave, but liquid
items should not be.
 Keep an eye on your time – leaving items in the autoclave for too long can result
in autoclave tape getting too sticky (at best) or media going bad (at worst).
 If dry items are not dry upon removal from the autoclave, you may move them to
the hood or the 37°C room to dry, but remember to take them out the next day!
Using the automatic autoclaves:
 Set the “sterilize” and “dry” times manually on both machines.
o All items should be sterilized for 20 min.
o Dry items (flasks, pipet tips, glass pipets, etc.) should also be dried for 20
min. Longer dry times are ok, but do not monopolize the autoclaves!
 Press the red reset button (if necessary).
 Put your items into the autoclave and close the door tightly.
 Turn the machine on. On autoclave #1, press the button matching your cycle
(liquid or dry) and then the on button; on autoclave #2, press the appropriate
button (liquid or gravity). Set a timer/reminder to return in 40 min.
 Make sure the autoclave is off and the chamber pressure is at 0 before opening.
Using the manual autoclave:
 Put your items in the autoclave and close it up.
 Turn the knob to “sterilize.” Set a timer/reminder to return in 20 min.
 Once the sterilize cycle is finished (20 min.), turn the knob to
“slow exhaust” (for liquid items) or “fast exhaust” (for dry items). Set a
timer/reminder to return in 20 min.
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The Media Bible / Section 1: Lab jobs

Check the chamber pressure. If it is at zero, turn the knob to “off”, open the
autoclave and remove your items. If it is not yet at zero, turn the knob to “fast
exhaust” and wait until the pressure comes down before opening.
Glassware and plastics (“dishes”)
Our lab uses separate soak bins for glass and plastic items. Glass/plasticware should be
removed from these bins at the end of every day, placed on the wire cart (or the white
plastic cart if absolutely necessary) and brought down to the 2nd floor to be washed.

Empty the beaker full of spatulas and spoons (in the weigh room) and include
those items in the “dishes” for each day.

Small items (caps, stir bars) should be placed in a wire basket; spoons and
spatulas should be placed in a beaker.

Remove all labels and autoclave tape from glass/plasticware (and caps) before
bringing dishes downstairs. This may be aided by the use of a razorblade.

Hand-wash flasks and beakers 2L or larger.
When the dishes have been cleaned, bring the carts upstairs and put items away as
appropriate:

Bottles should be capped and shelved.

Flasks 1L and smaller should be autoclaved. 125ml and 250ml flasks are capped
with the white caps (taped down with a small piece of autoclave tape). Larger
flasks are “sealed” with aluminum foil, with a piece of autoclave tape on top.
Autoclave 20 min./20 min. on a dry cycle. When cooled, move to the autoclaved
flask cabinet on the 3rd floor.

Beakers should NOT be stacked when wet – plastic beakers will stick together
and glass beakers may break.

Spoons and spatulas should be sorted and returned to the weigh room.

Extra caps should be stored in the appropriate drawer in Bay 1.

ASK if you are not sure where an item belongs!
The glass and plastic soak bins should be washed out as necessary, usually every week.
See section below (pg. 9) for more details.
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The Media Bible / Section 1: Lab jobs
Culture tubes

Cleaning culture tubes – daily
1. Check all bays for empty tubes. Ask before you take if you are not sure
which tubes need to be washed!
2. Remove all tape and stickers from test tube caps.
3. Remove caps from test tubes and place them in the ice bucket under the
Bay 2 sink.
4. Rinse out test tubes. It is important that no cell pellet remains in the
bottom of the tube! Usually it is sufficient to fill the tube halfway with
water and shake several times – you will probably want to wear gloves for
this step. If the pellet is particularly stubborn, you can use a test tube brush
to loosen it, but make sure that you rinse out any soap residue that the
brush might leave.
5. Deposit tubes in the tube soak bin under the sink in Bay 2.

Bringing culture tubes to be washed – twice weekly
1. Obtain a large plastic tray (usually under the bench in Bay 1).
2. Remove tubes from the tube soak and drain into the sink – you will
probably want to wear gloves for this. Stack the tubes in the plastic tray.
Be alert for pieces of broken glass! Broken tubes should be discarded in
the glass waste bin between Bay 1 and Bay 2.
3. Place caps in a wire bin.
4. Bring tubes and caps downstairs (on a cart), along with a note reading:
“Please do not use soap on caps and tubes, thanks!”
5. After these have been washed, you should autoclave them (see below).

Autoclaving culture tubes – twice weekly
1. Make sure tubes and caps are at least mostly dry before you begin.
2. Place culture tubes in wire racks – do not use racks that are too big for
these culture tubes! – and cap them. If a tube cannot be capped tightly, it is
probably a smaller size tube, which should be separated and used for
autoclaving toothpicks.
3. Place ONE long strip of autoclave tape diagonally across the top of the
rack of culture tubes.
4. Autoclave 20 min./20 min. on dry cycle. When cooled, move to the
autoclave cabinet on the 3rd floor.
The glass culture tube soak bins should be washed out as necessary, usually every other
week. See section below (pg. 9) for more details.
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The Media Bible / Section 1: Lab jobs
Glass pipets

Bringing glass pipets down to be washed – weekly
1. Obtain a large plastic tray (usually under the bench in Bay 1).
2. Each bay has one or two pipet soak bins, which should be emptied at least
once a week. Remove pipets from the pipet soaks – you will probably
want to wear gloves for this. Stack the pipets in the plastic tray. Be alert
for pieces of broken glass! Broken pipets should be discarded in the glass
waste bin between Bay 1 and Bay 2 or between Bay 3 and Bay 4.
3. Bring pipets downstairs (on a cart), along with a note reading: “Please do
not use soap on pipets, thanks!”
4. After these have been washed, you should sort and autoclave them (see
below).

Sorting glass pipets – weekly
1. Once pipets are clean, they should be sorted by size. It is usually easiest to
pull out the 25ml pipets first (since they are longer than the other sizes)
and then sort the rest of the pipets one handful at a time.
2. Pipets should be left out to dry in the wire sorting racks in the 2nd floor
media room.

Autoclaving glass pipets – weekly
1. Make sure pipets are at least mostly dry before you begin.
2. Fill a metal pipet canister full of one size of pipet. Do not fill the canister
so full that you must force the pipets in, but do not leave so much room in
the canister that the pipets can move around and break each other.
3. Cap the canister with the appropriately marked (e.g., “10ml”) metal top.
Do not force the top on if it does not fit! If the size marking is difficult
to read, re-mark neatly with a black Sharpie.
4. Place a small piece of autoclave tape over the seam between the canister
and the top, and another small piece of autoclave tape in the center of the
canister top.
5. Autoclave 20 min./20 min. on dry cycle. When cooled, move to the
appropriate drawer in on the third floor in Bay 1.
The glass pipet soak bins should be washed out as necessary, usually every other week.
See section below (pg. 9) for more details.
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The Media Bible / Section 1: Lab jobs
Pipet tips (for Pipetman)
There are four different types of pipet tips that you will re-fill on a regular basis:

P1000 tips (blue tips in the large individual blue boxes)

Yellow P200 tips (yellow tips in the large, stackable blue boxes)

White P200 “beveled” tips (white tips in the individual white/clear boxes)

P20 tips (white tips in the small individual blue boxes)
General notes:

Empty boxes should be collected from every bay each day, brought downstairs,
and sorted.

Remove any plastic inserts (purple or silver) from empty boxes. Purple inserts can
be recycled and should be disposed of in a plastic autoclave bag for this purpose.
When the bag is full, autoclave 20 min./20 min. on dry cycle. When cool, leave
bag in 2nd floor hallway.

Some tip refills are purchased from the stockroom; others are stocked in the lab
(see technician for location). Ensure that there is a good backup supply of refills
downstairs so that we will not run out of tips even if the stockroom does.

Re-fill empty boxes with the appropriate size tip, including the rack. For P200 tips
(and smaller), use the silver insert to guide the tips into the rack. This makes the
job infinitely easier.

For individual boxes, close and place a small piece of autoclave tape over the
seam between the top and bottom of the box. For the large stackable P200 tip
boxes, use a long strip of masking tape to hold the layers together and put a small
piece of autoclave tape on the top of the box.

Autoclave 20 min./20 min. on dry cycle. When cooled, move to the autoclave
cabinet on the 3rd floor.
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The Media Bible / Section 1: Lab jobs
Glass beads
Each person in the lab (2-3 per bay) has a small beaker of dirty glass beads on his/her
bench. These beads should be collected, washed and autoclaved once per week.
1. Collect dirty glass beads from all bays in a medium/large (500ml/1000ml) glass
bottle.
2. Fill the bottle 1/2 to 3/4 full with water and a small squirt of dish soap. Cap and
shake the bottle.
3. Dump out the soapy water in a sink, being careful not to lose any beads. A piece
of cheesecloth attached to a funnel can be very useful for this purpose!
4. Rinse the beads with distilled water, cap the bottle, shake, and pour off again.
Repeat these steps until the water is completely clear after shaking. It is
important that no soap residue remains on the beads. Wash twice with
ethanol (EtOH) to ensure that there is no soap residue.
5. “Aliquot” approximately 125ml of beads into small (250ml) bottles. Seal loosely,
mark bottle with a small piece of autoclave tape, and autoclave 20 min./20 min.
on a dry cycle. When cooled, move beads to the autoclave cabinet on the 3rd floor.
Wooden dowels
Each person in the lab (2-3 per bay) has a large beaker of dirty wooden dowels/empty
dowel tubes/caps on his/her bench. There is also a large beaker of wooden dowels in the
weigh room. These should be collected and brought downstairs at least once per week
(ideally every day), and autoclaved when there are enough to (almost) fill a rack of large
dowel tubes.
1. Collect dirty wooden dowels and empty dowel tubes/caps from each bay in a
large (2L) plastic beaker. Do not remove a collecting beaker from anyone’s
bench! These dowels should be brought downstairs and may be left there until
there are enough to autoclave.
2. Fill the empty tubes with the sticks. Tubes should be filled as full as possible
while still making sure that it is easy to remove a single dowel from the tube.
3. Remove any old autoclave tape from the tube caps and re-cap the tube. Place a
small piece of (new) autoclave tape on each cap.
4. Place each capped tube in a large tube rack. Autoclave 20 min./20 min. on a dry
cycle. When cooled, move rack to the autoclave cabinet on the 3rd floor.
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The Media Bible / Section 1: Lab jobs
Toothpicks
Each bay in the lab has a small beaker of empty tubes used to autoclave toothpicks (these
tubes are smaller than the regular culture tubes; the caps are smaller as well). These
should be collected and re-filled when the rack of autoclaved toothpicks is nearly empty
(< 5 tubes).
1. Collect empty toothpick tubes/caps from each bay in a small (500ml) plastic
beaker. Do not remove a beaker from anyone’s bench! These tubes should be
brought downstairs and may be left there until you are ready to autoclave a batch
of toothpicks. (Note: We do not re-use toothpicks, so there is no need to collect
dirty toothpicks.)
2. Fill the empty tubes with new toothpicks (purchased from the stockroom).
Toothpicks should be put in the tube skinny side up and fat end pointing to the
bottom of the tube. Fill the tubes as full as possible while still making sure that it
is easy to remove a single toothpick from the tube.
3. Remove any old autoclave tape from the tube caps and re-cap the tube. Place a
small piece of (new) autoclave tape on each cap.
4. Place each capped tube in the metal toothpick tube rack. Autoclave 20 min./20
min. on a dry cycle. When cooled, move rack to the autoclave cabinet on the 3rd
floor.
Squeeze bottles
Each bay in the lab has a set of three squeeze bottles (water, bleach and ethanol) that
should be refilled weekly. Each bottle should be filled to the marked line and also
checked for leaks or cracks. Make sure that the bottles are labeled properly and are in the
correct bay!

Water (ddH2O): Fill with deionized water from the large carboy in the centrifuge
core.

Bleach: Fill about 1/8 inch from the bottom with bleach, and fill the rest of the
way up with deionized water. If the bottle is more than half full, refill the bottle
almost to the fill line with water and add a small amount of bleach at the end.

Ethanol (EtOH): Fill from the large jug (1 gallon) of 90% EtOH (sometimes
labeled only as “TAX FREE ALCOHOL”) in the Alcohols cabinet underneath the
hood. DO NOT USE the 100% EtOH from the small plastic bottle!
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The Media Bible / Section 1: Lab jobs
Velvets
Our lab occasionally uses squares of velvet material to “stamp” identical patterns from
one plate to another. Dirty velvets are stored in a specially marked plastic tray under the
bench in Bay 1. You will have to wash, dry, sort and autoclave these velvets every two
months or so.

Washing and drying velvets
1. Remove the tray of dirty velvets from under the bench in Bay 1.
2. Bring the velvets to the washing machine in the ice machine room on the 2nd
floor. [The washing machine has a combination lock: 02-20-14] If the load of
velvets is especially large, you may want to sort them (lights/darks) but it is
okay to combine the colors. Wash on a normal cycle with NO SOAP.
3. Lay the velvets flat in another plastic tray to dry (overnight or longer).

Sorting and autoclaving velvets
1. Once the velvets are dry, they should be de-fuzzed. There is a lint brush
downstairs that may be used for this purpose, or you can use a large wad of
masking tape. While de-fuzzing, you should also sort the velvets by size and
color.
2. Make packets of 5-10 velvets of the same size and color in a thick aluminum
foil packet. These re-usable aluminum foils may be found in the far-left
cabinet underneath Bay 1 (closest to BG’s office).
3. Autoclave the packets 20 min./20 min. on a dry cycle. When cooled, the
packets should go on the top-most shelf of the autoclaved flask cabinet.
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The Media Bible / Section 1: Lab jobs
Changing soak bins/waterbaths

Dish soak bins. Change the water in these bins as necessary, usually every week.
1. Empty the bins. Use an ice bucket to remove most of the water and carefully
dump the rest into a sink.
2. Wipe out the bottom and sides of the bins with some clean water and a paper
towel or sponge. Make sure to get any labels or other gunk out of the bin! You
may need to repeat this step if the bins are very grimy.
3. Fill the bin to the fill-mark with deionized/distilled water from the tap, using
the nozzle/hose hookup in Bay 1.
4. Add a small amount of bleach (200-500ml) to the glass soak bin; use 7X
detergent (soap) solution (100-200ml) in the plastic soak bin. Both the bleach
and 7X detergent are stored under the sink in Bay 1. DO NOT USE
BLEACH IN THE PLASTIC SOAK BIN!

Culture tube soak bins. Change the water in these bins as necessary, usually every
other week.
1. Empty the bins by carefully dumping the water into a sink.
2. Wipe out the bottom and sides of the bins with some clean water and a paper
towel or sponge. Be very careful as there may be fragments of broken glass
in the bins! You may need to repeat this step if the bins are very grimy.
3. Fill the bins 1/2 to 3/4 full with deionized/distilled water from the tap, using
the nozzle/hose hookup in Bay 1.
4. Add a small amount of bleach (100-200ml) to the pipet soak bin.

Glass pipet soak bins. Change the water in these bins as necessary, usually every
other week.
1. Empty the bins by carefully dumping the water into a sink.
2. Wipe out the bottom and sides of the bins with some clean water and a paper
towel or sponge. You may need to repeat this step if the bins are very grimy.
3. Fill the bin 1/2 to 3/4 full with deionized/distilled water from the tap, using
the nozzle/hose hookup in Bay 1.
4. Add a small amount of bleach (100-200ml) to the pipet soak bin.
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The Media Bible / Section 1: Lab jobs

Hot water baths. There are two hot water baths in Bay 1 and one in Bay 2. Check
the water level in these daily – the water should come about 1 in. from the top of
the bin. Change the water in these as necessary, usually once per month. To
change the water:
1. Make sure that heating unit is cool. Remove the tube holders, thermometers
and heating unit from the waterbath. Be very careful with the thermometer
and heating unit and make sure to set the heating unit on its flat side.
2. Empty the bath by carefully dumping the water into a sink.
3. Wipe out the bottom and sides of the bath with some clean water and a paper
towel or sponge. You may need to repeat this step if the bath is very grimy.
4. Rinse the bath 1-2 times with deionized/distilled water.
5. Fill the bath to 1 in. from the top with deionized/distilled water from the tap,
using the nozzle/hose hookup in Bay 1.
6. Add a small amount of algaecide (10-25ml) to the bath (stored under the sink
in Bay 1) – WEAR GLOVES. Make a label with colored tape that says
“algaecide added / date / your initials” and stick the label to the front of the
bath.
7. Wipe down heating unit with a damp paper towel. You should also clean the
tube holders if necessary.
8. Replace heating unit and tube holders.

55°C water bath. This is the large metal bath at the end of Bay 1. Check the
temperature and water level in this daily – the water should come about 1-2 in.
above the wire rack. Change the water in this as necessary, usually once per
month. To change the water:
1. Unplug the water bath and turn it so that its back end is facing out.
2. There is a spout on the back end of the water bath; unscrew the spout and
allow the water to drain into a large (2L) plastic beaker. You will need to
change the beaker 2-3 times to drain all the water.
3. While bath is draining, remove the metal rack from the bottom of the bath and
clean with a coarse scrubby or paper towel.
4. After the bath has fully drained, wipe out with a damp paper towel and rinse
1-2 times with 500ml deionized/distilled water (drain as before).
5. Replace the rack, close the spout and fill the bath 1-2 in. above the wire rack.
Plug the bath in and position it normally on the bench.
6. Add a small amount of algaecide (25-50ml) to the bath (stored under the sink
in Bay 1) – WEAR GLOVES. Make a label with colored tape that says
“algaecide added / date / your initials” and stick the label to the front of the
bath.
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The Media Bible / Section 1: Lab jobs

25°C water bath. This is the large bath underneath Bay 3 for growing small (125ml500ml) cultures. Check the water level in this daily – the water should come about 12 in. above the metal rack. You should also remove any loose debris from the bath
(screws, paper towels, caps, etc.).
You should not need to change the water in this very often, but the procedure is
similar to changing the 55°C water bath (see previous page) in that it is drained
through a spigot. When changing the water, drain and wipe out the bottom of the bath
and the metal rack. You do not need to add algaecide to this bath.
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The Media Bible / Section 1: Lab jobs
Other assorted lab jobs
Keeping the 2nd floor media area clean
The media area should be “tidied up” every day before you leave.
 Keep liquid amino acids, sugars, etc. stocked.
 Wipe down the bench every day with water and then ethanol (using a paper
towel).
 Keep “dirty” (non-sterile) sticks separate and closed off from the rest of the room.
 Bag and move plates to storage (either upstairs or downstairs) in a timely manner
(i.e., as soon as they are dry).
 Autoclave or otherwise deal with any items as soon as you possibly can.
Autoclaved water
Refill purple-cap ddH2O bottles to 1L mark with water from the carboy in the centrifuge
core. Cap loosely and mark with a small piece of autoclave tape. Autoclave 20 min. on
liquid cycle. When cooled, move to the shelf in Bay 1.
Clean weigh room
The weigh room should be cleaned every week.
 Refill small containers of peptone, yeast extract and agar from large bins.
 Turn off and wipe down the large balance with a damp paper towel.
 Remove and wipe out the plexiglass “cage” around the balance with a damp paper
towel.
 Turn off and CAREFULLY clean out the analytical balance – wipe off the
windows and front panel with Windex and very gently clean out the weighing
apparatus with a damp paper towel.
 Wipe down the marble surface with a damp sponge.
 If necessary, also wipe down the wall behind the weigh table.
 Remove the absorbent paper (underneath the large balance only) and replace
with new piece.
 Wash the test tube holder and collect any dirty wooden dowels/spatulas. Rinse
and wipe dry the spatula/dowel collection beakers.
 Check the box of recipe cards and organize properly if necessary.
Clean out microwave
The microwave should be cleaned every month or so. Remove and wash the spinning
plate and wipe out the inside of the microwave.
Change diaper paper on shelves
The shelves for storing glassware are lined with diaper (absorbent) paper. These should
be replaced every month or so. Measure out an appropriate length of diaper paper and
tape onto the shelves with masking tape.
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The Media Bible / Section 1: Lab jobs
Wipe down/clean off carts
 The white plastic cart should be wiped down every other week with a damp paper
towel. Use coarse scrubby if the cart is particularly dirty.
 The wire cart used for dishes should be cleaned every month or so. Wipe down
with a damp paper towel, and clean the bottom tray (solid metal) thoroughly.
 The metal cart (marked as “do not use for dishes”) should also be cleaned every
month or so. Remove the plastic mats and wash with a dilute bleach solution.
DRY THE MATS THOROUGLY before replacing on the cart!
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Section 2. Making and Pouring Plates
General plate pouring procedure
1. Weight out dry ingredients and combine in a 2L flask. Most ingredients should be
weighed on the large balance, but weights smaller than 2g should be measured on
the analytical balance.
2. Fill flask with 1L water.
3. Add a medium-sized stir bar to flask.
4. Add additional liquid ingredients (NEVER sugars or antibiotics!) to flask.
5. Seal mouth of flask with tinfoil with a small piece of autoclave tape on top.
6. Autoclave 20 min. on liquid cycle.
7. Remove media from autoclave and allow to cool slightly while stirring gently
(avoid bubbles!) on a stir plate. The flask should be cool enough to hold
comfortably before pouring.
8. Add all the heat-sensitive ingredients (i.e., sugars, antibiotics) immediately before
pouring, allowing some time for mixing these into the media.
9. Pour the stack of plates being careful to avoid air bubbles and keeping the plate as
flat as possible. Try to keep the flask of media covered or at least somewhat
horizontal to avoid contamination.
10. Mark the plates. Label every plate with appropriate marker stripes down the
side (dragging the marker up the stack quickly is the best way to do this). Also
label the stack with a piece of masking tape on the top: plate contents / your
initials / date poured / batch # (if you have poured more than one liter worth of
the same type of plate that day).
11. Write down each stack that you have poured on the sheet on the 2nd floor.
12. Allow the plates to cool for at least two days before testing and bagging (see
below).
Testing plates
New batches of plates should be tested and bagged on the same day, at least two days
after they have been poured.
You should test one plate from each liter of plates poured. Once one plate has been
removed for testing, bag the remaining plates and store the stack in the 4°C room
adjacent to the plate-pouring room. Make sure that the stack is clearly and appropriately
labeled!
Yeast and bacterial plates are tested with separate sets of strains (see next page). All test
strains are stored in the –80°C freezer in the centrifuge core.
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The Media Bible / Section 2: Plates
Testing bacterial plates (eg: LB, LB Amp, LB Kan Cap, etc):
Mark the bottom of each plate to be tested with the following six “patches”:
WT
Amp
Kan
Cap
AmpCap
KanCap
Using a wooden dowel, streak the appropriate strain in each patch directly from the
frozen stock. Incubate the plates at 37°C overnight and check them the next day.
The pattern of growth for each type of plate (“X” = growth, gray = no growth):
Plate type
WT
Amp
Kan Cap AmpCap KanCap
LB
X
X
X
X
X
X
LB Amp
X
X
LB Kan
X
X
LB Cap
X
X
X
LB AmpCap
X
LB KanCap
X
Testing yeast plates (eg: YPD, dropout):
Mark the bottom of each plate to be tested with the following six “patches”:
WT —URA
—LEU
—HIS
—ADE
—TRP
—LYS
Using a wooden dowel, streak the appropriate strain in each patch directly from the
frozen stock. Incubate the plates at 25°C and check them in 2-3 days.
The pattern of growth for each type of plate (“X” = growth, gray = no growth):
Plate type
WT —URA —LEU —HIS —ADE
—TRP
—LYS
YPD
X
X
X
X
X
X
X
YPD-G418
X
X
doURA
X
X
X
X
X
X
doLEU
X
X
X
X
X
X
doHIS
X
X
X
X
X
X
doADE
X
X
X
X
X
X
doTRP
X
X
X
X
X
X
doLYS
X
X
X
X
X
X
doURA LEU
X
X
X
X
X
doURA HIS
X
X
X
X
X
doLEU HIS
X
X
X
X
X
15
The Media Bible / Section 2: Plates
Taking plate inventory





Count the stacks of plates that are stored in the 4°C room on the 3rd floor, checking
for contaminated plates (REMOVE and THROW AWAY any contaminated stacks).
Mark this number as “stocks.”
Count the stacks of plates that are stored in the 4°C room on the 2nd floor, checking
for contaminated plates (REMOVE and THROW AWAY any contaminated stacks).
Mark this number as “spare.”
Count the stacks of plates that are in the plate pouring room on the 2nd floor. Mark
this number (use tally marks) as “new poured.”
If the 3rd floor boxes are not full (i.e., “stocks” number does not match the ideal
number), bring the appropriate number of spare stacks (take the oldest stacks so they
will be used first) from the 2nd floor to the 3rd floor. Adjust the stocks/spare numbers
accordingly.
A list of plates and the appropriate stocks follows on the next page.
Other notes on plate storage
 Boxes should always be organized such that the oldest plates are used first (i.e.,
toward the front).
 Store “unchecked” stacks of plates (i.e., tested but not confirmed) in the 4°C room
(adjacent to the plate-pouring room). Keep these stacks in a box that is separate
from stacks that have been confirmed.
 Once stacks have been tested and confirmed, mark them with a check and add them to
the box(es) of confirmed stacks.
16
The Media Bible / Section 2: Plates
Plate
YPD
LB Amp
LB Kan
LB Cap
LB Kan Cap
LB
-5
doUra
doLeu
doHis
doAde
doTrp
doLys
YPD-G418
YPGal
Gal-5
doUraGal
doLeuGal
doUra-doHis
doUra-doLeu
doLeu-doHis
Number of plates to stock*
stocks
spare
10
5
5
5
5
2
3
1
2
1
5
2
5
2
5
5
5
5
5
2
3
3
2
2
3
1
3
1
3
1
2
1
2
2
2
* Please stock twice this number of stacks prior to long breaks/holidays.
17
The Media Bible / Section 2: Plates
Plate recipes
LB
10g tryptone
5g yeast extract
5g NaCl
20g agar
1L ddH2O
2.5ml 1N (1M) NaOH
Weigh out and combine ingredients (in this order) and autoclave 20 min. on liquid cycle.
Allow media to cool slightly while stirring. Pour.
LB + Antibiotic (Amp, Kan, Cap)
10g tryptone
5g yeast extract
5g NaCl
20g agar
1L ddH2O
2.5ml 1N (1M) NaOH
Weigh out and combine ingredients (in this order) and autoclave 20 min. on liquid cycle.
Allow media to cool slightly while stirring. In the meantime, thaw an aliquot of your
antibiotic (Amp, Cap, Kan). Add 1ml of 1000X antibiotic (if you are making a
multiple antibiotic plate, e.g. LB Kan Cap, simply add 1ml of each one). Allow the
antibiotic to stir into the media. Pour.
YPD
20g Peptone Y (Bacto-Peptone)
10g Yeast extract
0.15g L-tryptophan
20g agar
1L ddH2O
Weigh out and combine ingredients (weigh tryptophan on the analytical balance) and
autoclave 20 min. on liquid cycle. Allow media to cool slightly while stirring. Add 50ml
40% glucose and allow the sugar to stir into the media. Pour.
Variations:
 For YPD-G418, also add 1ml 1000X G418 (frozen aliquot).
 For YPGal, omit glucose and add 100ml sterile 20% galactose after
autoclaving.
 For YPGlycerol, omit glucose and add 67ml sterile 50% glycerol after
autoclaving.
 For YPDSorbitol, use only 620ml of ddH2O. Add 330ml of sterile 2.4M
sorbitol after autoclaving.
 For YPEG, omit glucose. Add 30ml 100% EtOH and 30ml sterile 100%
glycerol after autoclaving.
18
Dropout plates (-5, doUra, doLeu, doHis)
1.5g Yeast nitrogen base
5g Ammonium sulfate
2.5g Amino acid mix
25g agar
1L ddH2O
Weigh out and combine ingredients. The amino acid mix used will depend on the type of
plate that you are making (e.g., use the –5 mix if you are making –5 plates, use the –Ura
mix if you are making –Ura plates). Autoclave 20 min. on liquid cycle. Allow media to
cool slightly while stirring. Add 50ml 40% glucose (or 100ml 20% galactose, if making
doGal plates) and allow the sugar to stir in. Pour.
Dropout plates (doAde, doTrp)
1.5g Yeast nitrogen base
5g Ammonium sulfate
2.5g –5 a.a. mix (–6 for –Lys)
25g agar
1L ddH2O
Weigh out and combine ingredients. Autoclave 20 min. on liquid cycle. Allow media to
cool slightly while stirring. Add 50ml 40% glucose, then add the following liquid
amino acids (leaving out Ade or Trp as appropriate):
3ml 100mM His
17ml 100mM Leu
10ml 20mM Ura
10ml 30mM Ade
10ml 40mM Trp
Allow the sugar and amino acid liquids to stir into the media. Pour.
Double dropout plates
1.5g Yeast nitrogen base
5g Ammonium sulfate
2.5g –5 amino acid mix
25g agar
1L ddH2O
Weigh out and combine ingredients. Autoclave 20 min. on liquid cycle. Allow media to
cool slightly while stirring. Add 50ml 40% glucose, then add the following liquid
amino acids (leaving out two or more as appropriate):
3ml 100mM His
17ml 100mM Leu
10ml 20mM Ura
10ml 30mM Ade
10ml 40mM Trp
Allow the sugar and amino acid liquids to stir into the media. Pour.
19
The Media Bible / Section 2: Plates
5-FOA plates
1.5g yeast nitrogen base
5g ammonium sulfate
2.5g –Ura amino acid mix*
25g agar
1L ddH2O
Weigh out and combine ingredients. Autoclave 20 min. on liquid cycle. Allow media to
cool slightly while stirring. Add 50ml 40% glucose. Add 0.5g FOA (wear gloves!).
Add 12ml 20mM Ura liquid. Allow the sugar, FOA powder and amino acid liquids to
stir into the media until the FOA is completely dissolved. Pour.
* You could also make dropout 5-FOA plates, but they must contain the noted amount of Ura liquid.
Benomyl plates
This recipe will give you 10-plate batches of benomyl plates of seven different
concentrations (about 70 plates altogether). You will probably need to pour these plates
on the 3rd floor.



Make a 10mg/ml stock of benomyl in DMSO (0.1g benomyl + 10ml DMSO). Do
not make this more concentrated or the benomyl will come out of solution.
Combine the following in a 2L beaker and stir until uniform:
o 35g Peptone Y
o 35g agar
o 17.5g Yeast extract
o 1750ml ddH2O
o 0.2625g L-tryptophan
Aliquot 250ml into each of seven 1L bottles and autoclave 20 min. on liquid
cycle.

Allow media to cool to 55°C in the waterbath (Bay 1 on 3rd floor).

Add 12.5ml of 40% glucose to each bottle.

Then, add benomyl and DMSO to each bottle as follows, being careful to label
the bottles appropriately:

1. 0mg/ml plates
0µl benomyl
500µl DMSO
2. 2.5µg/ml plates
63µl benomyl
438µl DMSO
3. 5µg/ml plates
125µl benomyl
375µl DMSO
4. 7.5µg/ml plates
188µl benomyl
313µl DMSO
5. 10µg/ml plates
250µl benomyl
250µl DMSO
6. 12.5µg/ml plates
313µl benomyl
188µl DMSO
7. 15µg/ml plates
385µl benomyl
125µl DMSO
Pour one stack of plates from each bottle. Make sure that you label each stack
with the concentration of benomyl it contains! Store these plates in the 2nd floor
4°C room in a specially marked box.
20
The Media Bible / Section 2: Plates
GNA plates
30g nutrient broth
20g agar
10g yeast extract
1L ddH2O
50g glucose (dextrose)
Weigh out and combine ingredients. Autoclave 30 min. on liquid cycle. Allow to cool
slightly. Pour plates separately on counter and do not stack them until they have
solidified!
SPOR plates
First, make sure that you have fresh Spore Mix I and II (recipes below). Then, make the
media:
2.5g Yeast extract
20g agar
15g Potassium acetate
900ml ddH2O
Weigh out and combine ingredients. Autoclave 20 min. on liquid cycle.
After autoclaving, Add:
0.5ml Spore Mix I
50ml 4mg/ml Threonine
40ml Spore Mix II
2.5ml 40% glucose
Mix well and allow to sit in 55°C water bath 15 min. Pour.
Spore mix I
1g leucine
1g phenylalanine
1g lysine
1g methionine
1g arginine
1g histidine
1g tryptophan
100ml ddH2O
Weigh out and combine ingredients in a glass milk bottle. Autoclave 20 min. on liquid
cycle. Store at room temperature for up to 3 months.
Spore mix II
0.4g adenine
0.08g tyrosine
0.4g uracil
400ml ddH2O
Weigh out and combine ingredients in a 1L glass bottle. Autoclave 20 min. on liquid
cycle. Store at room temperature for up to 3 months.
21
Section 3. Making Liquid Medias and Lab Stocks
General media making procedure

Weigh out dry ingredients and combine in a 4L beaker. Allow to stir until
uniform.

Make 10 labels: “name of media / date / your initials” and adhere them neatly to
1L bottles. Also mark the cap of each bottle with a small piece of autoclave tape.

Aliquot 300ml media into each bottle. Autoclave 20 min. on liquid cycle. When
cool, move to the liquid media cabinet in Bay 1.

NOTE: This is just a guideline. Many liquids DO NOT and SHOULD NOT
follow this protocol, so make sure you use the correct recipe!
Liquid Medias and Stocks
Media type
Please stock*:
2X YEP
10 x 300ml
2X LB
10 x 300ml
2X –5
20% galactose
10 x 300ml
5 x 500ml
10 x 100ml
2 x 250ml
20% raffinose
1 x 250ml
50% PEG-3350
2 x 250ml
50% glycerol
1 x 250ml
20mM Ura
5 x 100ml
100mM Leu
5 x 100ml
40mM Trp
5 x 100ml
100mM His
5 x 100ml
30mM Ade
5 x 100ml
4mg/ml Lys
1 x 100ml
4mg/ml Thr
1 x 100ml
40% glucose
* Please stock twice this number prior to long breaks/holidays.
22
The Media Bible / Section 3: Liquid Media and Stocks
Recipes for liquid medias
2X YEP
120g Peptone Y (Bacto-Peptone)
0.9g L-tryptophan
60g Yeast extract
3000ml ddH2O
Weigh out ingredients and mix until dissolved (in a 4L beaker). [Note: To make 1X YEP,
use half of all ingredients except ddH2O; proceed as for 2X.] Aliquot 300ml into each of
10 labeled 1L bottles. Autoclave 20 min. on liquid cycle. When cool, move to liquid
media cabinet in Bay 1.
2X LB
60g tryptone
3000ml ddH2O
30g yeast extract
15ml NaOH
30g NaCl
Weigh out ingredients and mix (in this order) until dissolved (in a 4L beaker). [Note: To
make 1X LB, use half of all ingredients except ddH2O; proceed as for 2X.] Aliquot
300ml into each of 10 labeled 1L bottles. Autoclave 20 min. on liquid cycle. When cool,
move to liquid media cabinet in Bay 1.
2X –5
30g ammonium sulfate
12g –5 amino acid mix
9g yeast nitrogen base
3000ml ddH2O
Weigh out ingredients and mix until dissolved (in a 4L beaker). [Note: To make 1X -5,
use half of all ingredients except ddH2O; proceed as for 2X.] Aliquot 300ml into each of
10 labeled 1L bottles. Autoclave 20 min. on liquid cycle. When cool, move to liquid
media cabinet in Bay 1.
4% agar
Weigh out 12g agar into each of 10 (or more) labeled 1L bottles. Aliquot 300ml ddH2O
to each bottle. Autoclave 20 min. on liquid cycle. When cool, move to the agar/ddH2O
shelf in Bay 1.
YNB
1.5g yeast nitrogen base
1000ml ddH2O
5g ammonium sulfate
Weigh out ingredients and mix until dissolved (in a 2L beaker). Aliquot 100ml into each
of 10 labeled milk bottles. Autoclave 20 min. on liquid cycle. When cool, move to liquid
media cabinet in Bay 1.
23
The Media Bible / Section 3: Liquid Media and Stocks
Recipes for sugar stocks
40% glucose






Combine 800g glucose (dextrose) and 1200ml ddH2O in a 2L beaker.
Microwave 8 min. on high or until dissolved.
Stir until sugar is completely dissolved (i.e., solution is clear).
Pour into a 2L graduated cylinder and adjust volume to 2000ml with ddH2O. Return
solution to beaker and stir until well mixed.
Aliquot 500ml into each of four labeled 1L glass bottles, or 100ml into each of 20
milk bottles (or some combination of the two).
Autoclave 20 min. on liquid cycle. When cooled, move to media cabinet in Bay 1.
20% galactose





Combine 100g galactose and 400ml ddH2O in a 1L beaker.
Mix until fully dissolved and the solution is clear. DO NOT HEAT. This process
takes a while, sometimes >1 hour.
When solution is uniform, pour into a 1L graduated cylinder and adjust volume to
500ml with ddH2O. Return solution to beaker and stir until well mixed.
Filter sterilize 250ml of solution with each of two (separate, brand new) sterile
Nalgene filter units.
Label and store in media cabinet in Bay 1.
20% raffinose





Combine 100g raffinose and 400ml ddH2O in a 1L beaker.
Mix until fully dissolved and the solution is clear. DO NOT HEAT. This process
takes a while, sometimes overnight. If you are not returning the next day, ask a fulltime lab member to finish the solution for you.
When solution is uniform, pour into a 1L graduated cylinder and adjust volume to
500ml with ddH2O. Return solution to beaker and stir until well mixed.
Filter sterilize 250ml of solution with each of two (separate, brand new) sterile
Nalgene filter units.
Label and store in media cabinet in Bay 1.
24
The Media Bible / Section 3: Liquid Media and Stocks
Amino acid liquid stocks
20mM Ura
Combine 1.1g uracil and 500ml ddH2O in a 1L glass beaker. Microwave on high for 2
min. and stir until fully dissolved. Aliquot 100ml into each of five labeled milk bottles
and autoclave for 20 min. on liquid cycle. When cool, move bottles to liquid media
cabinet in Bay 1.
100mM Leu
Combine 6.6g L-Leucine and 500ml ddH2O in a 1L glass beaker. Microwave on high
for 2 min. and stir until fully dissolved. Aliquot 100ml into each of five labeled milk
bottles and autoclave for 20 min. on liquid cycle. When cool, move bottles to liquid
media cabinet in Bay 1.
40mM Trp
Combine 4g L-Tryptophan and 500ml ddH2O in a 1L glass beaker. Microwave on high
for 2 min. and stir until fully dissolved. Aliquot 100ml into each of five foil-wrapped
and labeled milk bottles and autoclave for 20 min. on liquid cycle. When cool, move
bottles to liquid media cabinet in Bay 1.
100mM His
Combine 10.5g L-Histidine and 500ml ddH2O in a 1L glass beaker. Microwave on high
for 2 min. and stir until fully dissolved. Aliquot 100ml into each of five labeled milk
bottles and autoclave for 20 min. on liquid cycle. When cool, move bottles to liquid
media cabinet in Bay 1.
30mM Ade
Combine 2.8g adenine and 500ml ddH2O in a 1L glass beaker. Microwave on high for 2
min. and stir until fully dissolved. Aliquot 100ml into each of five labeled milk bottles
and autoclave for 20 min. on liquid cycle. When cool, move bottles to liquid media
cabinet in Bay 1.
4mg/ml Lys
Combine 2g L-Lysine and 500ml ddH2O in a 1L glass beaker. Microwave on high for 2
min. and stir until fully dissolved. Aliquot 100ml into each of five labeled milk bottles
and autoclave for 20 min. on liquid cycle. When cool, move bottles to liquid media
cabinet in Bay 1.
4mg/ml Thr
Combine 2g L-Threonine and 500ml ddH2O in a 1L glass beaker. Microwave on high
for 2 min. and stir until fully dissolved. Aliquot 100ml into each of five labeled milk
bottles and autoclave for 20 min. on liquid cycle. When cool, move bottles to liquid
media cabinet in Bay 1.
25
The Media Bible / Section 3: Liquid Media and Stocks
Other liquid stocks
Coomassie blue stain
0.5g Coomassie blue (powder)
400ml ddH2O
500ml methanol (MeOH)
100ml acetic acid
Measure MeOH directly in a pre-labeled 1L glass bottle (you will usually be refilling a
bottle that had previously contained Coomassie). Dissolve the Coomassie blue powder in
the MeOH and add remaining ingredients in the order above (it is okay to measure by the
markings on the side of the bottle). Mix by gently inverting or shaking. Store solution on
shelves in Bay 4.
Destain
800ml ddH2O
100ml acetic acid
100ml methanol (MeOH)
Combine ingredients in order given above directly in a pre-labeled 1L glass bottle (you
will usually be refilling a bottle that had previously contained destain solution). It is okay
to measure by the markings on the side of the bottle. Mix by gently inverting or shaking.
Store solution on shelves in Bay 4.
50% glycerol
 Mix 250ml 99% glycerol (do NOT use ultrapure!) with 250ml ddH2O in a 1L
glass beaker until solution is uniform in appearance.
 Wash the graduated cylinder used to measure the glycerol with this solution,
return to beaker and mix well.
 Aliquot 250ml into two labeled 500ml glass bottles.
 Autoclave 20 min. on liquid cycle. When cool, move to the liquid media cabinet
in Bay 1.
50% PEG-3350
 Mix 250g PEG-3350 (do NOT use PEG-8000) with 250ml ddH2O in a 1L glass
beaker until PEG is fully dissolved. This may take >1 hour.
 When solution is completely clear, filter sterilize 250ml of solution with each of
two (separate, brand new) sterile Nalgene filter units. The PEG solution is
extremely viscous so this may take a long time, i.e., overnight. If you are not
returning the next day, ask a full-time lab member to finish the solution for you.
 When PEG has finished sterilizing, move bottles to the liquid media cabinet in
Bay 1.
200mM Inositol
 Combine 3.6g inositol and 100ml ddH2O in a milk bottle.
 Autoclave 20 min. on a liquid cycle. When cool, move to the liquid media cabinet in
Bay 1.
26
The Media Bible / Section 3: Liquid Media and Stocks
KOAc/yeast extract mix
12.5g yeast extract
500ml ddH2O
75g potassium acetate
Weigh out ingredients and mix in a 1L beaker until dissolved. Aliquot 100ml into each of
5 labeled milk bottles. Autoclave 20 min. on liquid cycle. When cool, move to liquid
media cabinet in Bay 1.
100ml of this media may be added to 900ml ddH2O/20g agar (autoclaved) to make the
base media for SPOR plates.
1N (1M) NaOH
Dissolve 20g NaOH pellets (wear gloves) in 500ml ddH2O in a plastic bottle (NaOH will
corrode a glass bottle). When pellets are mostly dissolved, mix gently by swirling. You
can increase the concentration of NaOH up to 10M (200g pellets in 500ml).
10X PBS
320g NaCl
9.6 KH2PO4
8g KCl
4000ml ddH2O
57.6g Na2HPO4
Mix salts together and dissolve in 2000ml ddH2O (use 3L beaker). When dissolved,
aliquot 500ml into each of four 1L glass bottles. Adjust volumes to 1000ml with ddH2O.
27
Section 4. Other Stocks
Amino acid powders (mixes)
20g alanine
20g arginine
20g asparagines
20g aspartic acid
20g cysteine
20g glutamine
20g glutamic acid
20g glycine
20g isoleucine
20g methionine
20g phenylalanine
20g proline
20g serine
20g threonine
20g tyrosine
20g lysine
5g adenine
20g histidine
100g leucine
20g tryptophan
20g uracil
Add together amino acids (through tyrosine) in a 2L plastic beaker. If making –6 powder,
omit all italicized amino acids. If making –5 powder, add lysine and omit other italicized
amino acids. To make a specific dropout powder, add all italicized amino acids except the
one you are “dropping out” (e.g., omit Ura in doUra powder).
Once you have added all the amino acids, parafilm the beaker and shake well. “Aliquot”
the powder into labeled plastic containers.
Notes:
 You can use this base powder to make several batches of dropout powder at once
(e.g., make –5 powder, divide in half and make one half –Ura). Just be sure to
adjust the weights accordingly!
 Since there are so many amino acids to combine, it can be helpful to check off
each one as you go along, for example, you might reprint this page and cross off
each ingredient as it is added.
28
The Media Bible / Section 4: Other Stocks
Frozen stocks
100mg/ml (1000X) Ampicillin (AMP)
Mix 5g ampicillin powder in 50ml ddH2O in a 50ml conical falcon tube. Allow to
dissolve with shaking (use the gel shaker/”bellydancer” in Bay 4). While powder is
dissolving, label and date 50 1.5ml Eppendorf tubes. Filter sterilize the solution with a
small Nalgene unit and aliquot 1ml liquid stock into each tube using a repeat pipetter.
Store aliquots in the –20° freezer box.
200mg/ml (1000X) G418
Mix 10g G418 powder in 50ml ddH2O in a 50ml conical falcon tube. Allow to dissolve
with shaking (use the gel shaker/”bellydancer” in Bay 4). While powder is dissolving,
label and date 50 1.5ml Eppendorf tubes. Filter sterilize the solution with a small Nalgene
unit and aliquot 1ml liquid stock into each tube using a repeat pipetter. Store aliquots in
the –20° freezer box.
60mg/ml (1000X) Kanamycin (KAN)
Mix 3g kanamycin powder in 50ml ddH2O in a 50ml conical falcon tube. Allow to
dissolve with shaking (use the gel shaker/”bellydancer” in Bay 4). While powder is
dissolving, label and date 50 1.5ml Eppendorf tubes. Filter sterilize the solution with a
small Nalgene unit and aliquot 1ml liquid stock into each tube using a repeat pipetter.
Store aliquots in the –20° freezer box.
15mg/ml (1000X) chloramphenicol (CAP)
Mix 0.75g (750mg) chloramphenicol powder in 50ml EtOH in a 200ml glass bottle.
Allow to dissolve with shaking (use the gel shaker/”bellydancer” in Bay 4). While
powder is dissolving, label and date 50 1.5ml Eppendorf tubes. Aliquot 1ml liquid stock
into each tube using a repeat pipetter. Store aliquots in the –20° freezer box.
1mM dNTPs
Combine 30µl each dNTP (dATP, dTTP, dCTP, dGTP) with 2.88ml ddH2O in a 15ml
conical falcon tube and mix well. Label and date 50 250µl Eppendorf tubes. Aliquot 60µl
liquid stock into each tube. Store aliquots in the –20° freezer box.
2mg/ml salmon sperm DNA
Mix 0.1g (100mg) salmon sperm DNA (looks like a cotton ball) in 50ml TE in a 50ml
conical falcon tube. Allow to dissolve with shaking (use the gel shaker/”bellydancer” in
Bay 4) overnight. Label and date 50 1.5ml screw-cap tubes. When solution is fully
uniform, aliquot 1ml liquid stock into each tube using a repeat pipetter. DO NOT
FILTER STERILIZE! Store aliquots in the –20° freezer box.
29
Section 5. Lab map (or, ‘Where can I find this?!’)
NaOH (1M / 1N): Top shelf, right side of Bay 1
7X soap solution: Under Bay 1 sink
Acetic acid: Acids cabinet under hood
Agar (powder): Under reagent shelves
Algecide: Under Bay 1 sink
Amino acid mixes: Reagent shelf #2
Amino acids (liquids): Media cabinet in Bay 1
Amino acids (powders): Reagent shelf #2
Ammonium sulfate: Reagent shelf #4
Ampicillin: –20°C freezer
Bacto-Peptone: Reagent shelf #1
Beakers: Clean items shelf #3
Benomyl (powder):
Bleach: Under Bay 1 sink
Cheesecloth: In funnel drawer in Bay 4
Chloramphenicol (liquid): 4°C refrigerator
Chloramphenicol (powder): 4°C refrigerator
Coomassie blue powder: Reagent shelf #3
Culture tubes (autoclaved): Autoclave cabinet #2
Culture tubes (dirty): Under sink in Bay 2
Diaper Paper: Weigh room
DMSO: Chemical shelves
Dowel tubes (empty): On benches (collect), stored in plate pouring room
Ethanol (70%): Alcohols cabinet under hood
Flasks (autoclaved): Autoclave cabinet #1
FOA: -20°C freezer
Funnels: In funnel drawer in Bay 4
G418: –20°C freezer
Galactose (20%): Media cabinet in Bay 1
Galactose (powder): Under reagent shelves
Glass beads (autoclaved): Autoclave cabinet #2
Glass beads (dirty): On benches (collect), wash and autoclave immediately
Glass bottles: Clean items shelf #4 and #5
Glass pipet canisters: On benches (collect), stored in plate pouring room
Glass pipets (autoclaved): Sterile pipet drawers in Bay 1
Glass pipets (dirty): In pipet soak bins in every bay (collect), wash and autoclave
immediately
Glucose (40%): Media cabinet in Bay 1
Glucose (powder): Under reagent shelves
30
The Media Bible / Section 4: Other Stocks
Glycerol (50%): Media cabinet in Bay 1
Glycerol (99%): Chemical shelf
Inositol:
Kanamycin (liquid): –20° freezer
Kanamycin (powder): 4°C refrigerator
KCl (potassium chloride): Reagent shelf #5
KH2PO4: Borrow from lab member
Lint brush: Labels and tape drawer in Bay 1
L-tryptophan: Reagent shelf #2
Methanol: Alcohols cabinet (under hood)
Na2HPO4: Borrow from lab member
NaCl: Chemical shelf
NaOH (pellets): Chemical shelf
Nutrient broth:
Orange caps (for glass bottles): Misc. caps drawer in Bay 1
PEG-3350: Chemical shelf
Peptone: Reagent shelf #1
Pipet soak bins: Under each bay
Pipet tip boxes (empty): On benches (collect), store in media room
Pipet tips: From stockroom, store in media room
Plastic tray: Under bench in Bay 1
Potassium acetate (KOAc): Chemical shelves
Purple caps (for ddH2O bottles): Misc. caps drawer in Bay 1
Raffinose (20%): Media cabinet in Bay 1
Raffinose (powder): Reagent shelf #4
Razorblades: Taped to wall next to glass soak bin
Sorbitol (2.4M): Borrow from lab member
Spatulas: Weigh room
Spoons: Weigh room
Squeeze bottles: On benches (refill); empties on shelf in Bay 2
Stir bars: Metal cabinet in weigh room
Tester plates: 25°C incubator
Tester strains: -80° freezer in weigh room
Toothpick tubes (empty): On benches (collect), store in media room
Toothpicks: From stockroom
Tryptone: Reagents shelf #1
Velvet foil packets: Replica Plating cabinet in Bay 1
Velvets (autoclaved): Autoclave cabinet #1
Velvets (dirty): In bin under bench in Bay 1
White caps (for flasks): Misc. caps drawer in Bay 1
Windex: Under sink in lunch room
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The Media Bible / Section 4: Other Stocks
Wire bins: On carts
Wire tube rack: On benches (collect), store in media room
Wooden dowels: On benches (collect), store sealed in media room
Yeast extract: Reagents shelf #1
Yeast nitrogen base: Reagents shelf #2
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