Version: 7/1/96
A. General Policies.
1. Before researchers can use the tissue culture lab, they must carefully read this entire document. The next step is to perform the various required techniques for the first time while being supervised in person by Gelb or another certified lab member. Only authorized users will be given a key to the tissue culture lab, and only authorized users are allowed to enter the lab. Researchers from other groups cannot use the tissue culture lab unless they have permission from Gelb. In most cases, permission will not be granted. These steps are to insure that sample contaminaton is kept to a minimum as loss of samples due to contamination is very expensive and time-consuming.
2. Items that belong in the tissue culture lab shall remain there. It is very frustrating when you are working in the lab and then you reach for an item and find that it is not in the lab.
3. Individual users of the tissue culture lab are to seggregate their sterile expendable items (plasticware, bottles and tubes of fluids, etc....), and such items are to be labeled with the researcher's name. Without exception, such items are not to be handled by other workers. By doing this, each lab worker maintains the responsibility of keeping their items sterile and needs not to be concerned about contamination beyond their control. Exceptions are items that are individually wrapped in sterile packages (i.e. pipets, cell scrapers, etc...) since such items can be inspected by workers to insure that their packagings are not damaged. Items such as squirt bottles of ethanol that essentially cannot become contaminated can be used by all workers and need not be seggregated or labeled with worker's names. Such items should include TCL on the label to designate that it is to be kept in the tissue culture lab. Bottles of serum, media, trypsin, glutamine, antibiotics, etc... from vendors are to be stored unopened in the appropriate place. If a worker wishes to use these items, they should label them with their name and seggregate them with their supplies. Often such items are to be opened and aliquoted into sterile tubes. In this case the worker will aliquot the entire contents of the bottle into tubes that are labeled with the worker's name and seggregated. Do not open a bottle of commercial product, aliquot part of it, and return the remaining portion to the general stock. This is to insure that when a worker opens a bottle of commercial agent, they can be sure that it has the sterility provided by the vendor. Aliquoting a portion of one item to one worker and another portion to another worker can be carried out as long as both workers agree on this together. In this case, all aliquoting should be done at one time so that partially used commercial supplies are not left without seggregation and without labeling with lab worker names. Frozen items must be stored in the -20°C freezer dedicated for tissue culture reagents and 4°C items must be stored in the refrigerator in the tissue culture lab.
4. If you notice that we are running low on certain items, order them well in advance of running out. It can take up to 2-3 weeks to receive certain items, for example from VWR or Fisher.
5. Biohazardess Waste is handle in the following way. The big white Nalgene tank should have an autoclavable bag liner. All used plasticware should be put into this tank. When the tank is full, the bag should be closed, removed from the tank, and autoclaved. Immediately put a new bag in the empty tank. Liquid waste should be poured into the wide-mouth jar that contains bleach. Do not put containers or pipets with excessive liquid directly into the white tank. Instead, first pour the liquid into the bleach jar and then put the plasticware into the white tank. Solid waste such as gloves, can go into the white tank. Bleach-treated waste can be poured down the sink. Pasteur pipets should be put into the red sharps container. When full this container is autoclaved and the Pasteur pipets are crushed and the glass is poured into a carboard box in the labs designated for glassware waste. The aspirator trap contains bleach and when full it can be poured down the sink. Thus, there are four, and only four waste containers: bag-lined white tank, Pasteur pipet container, wide-mouth jar and aspirator trap. No additional waste containers are allowed.
6. A set of glassware labeled TCL (graduate cylinders, beakers) is dedicated for making media. This glassware should not leave the tissue culture lab, and it should only be used for tissue culture work.
B. Use of the Laminar Flow Hood.
The UV lamp should be turned on approx. 15 min before use of the hood. There is no need to leave the lamp on for longer periods such as overnight. Never turn the hood fan off. The fan switch should be taped in the on position so that it is not accidentally turned off. Adjust the stool height so that your chin is above the bottom of the glass hood sash. This will prevent contamination of the worker by hood contents. Some people may prefer to work at a lower stool position but then you must wear your own face shield (labeled with your name). Squirt some 70% ethanol onto the bench and wipe it with a paper towel. You should wipe most of the bench surface; only the surfaces within about 6 inches from the walls of the hood need not be wiped. Very few items should be left in the hood when you are finished using it. The electric pipettor is to be left on the right side of the bench and plugged in to the battery recharger. Glass containers of sterile Pasteur pipets can be left on the left side of the hood bench; make sure your container is labeled with your name. A few tube racks can remain in the hood as well. Do not leave packages of sterile plasticware in the hood when you are done using it. Such items are to be removed and keep seggregated in each workers designated area. Wipe the bench surface with 70% ethanol when done using the hood, and turn off the fluorescent light. The end of the aspirator tubing should be placed in the black clip that is attached to the hood sash. After use, be sure to squirt a bit of 70% ethanol into the aspirator tubing while the pump is running in order to sterilize the line. If you spilled any liquids on the bench, pour some 70% ethanol over the spill and scrubb with the green scouring pad over the spill area, then wipe it dry with paper towel.
If the power on the hood goes out, you need to restore it, and then wipe all inner surfaces of the hood with 70% ethanol (you can spray with ethanol and then wipe but do not spray the perforated celling; instead wipe it with a paper towel moistened with 70% ethanol. Be sure to wipe the grill along the front edge of the bench and the inside of the hood sash (you can unclip the locks and flip the sash open). Turn on the UV lamp and leave it for an hour or so before using the hood.
C. Sterile Techniques.
It is difficult to write down precisely all of the skills needed to maintain strict sterility. What follows are some general statements and some specific examples.
When you want to carry out a sterile technique, allow uninterupted time to do it, and don't rush through it. Patience will save you countless hours needed to recover from a contamination problem. Maintaining sterility relies a good deal on common sense. For example, any surface that you touch with your finger is no longer sterile and should not be allowed to come in contact with sterile liquids. You cannot assume that the air outside of the hood is sterile, and thus you should never open a sterile container outside of the hood, even for a brief moment.
Here are some specific examples:
1. Storing sterile liquids. Sterile liquids should be kept in tubes and bottles in a verticle position. Never lay falcon tubes or glass bottles of liquids in a horizontal position since prolonged contact with the cap can lead to leakage and thus contamination. Never fill containers more than about 80-90% full to insure that the solution does not stay in contact with the cap. Also frozen solutions will expand and you don't want frozen liquid pushing against the cap.
2. Autoclaving Liquids: Put it into glass tissue culture bottles (fill them about 50% full), put screw cap on loosely (don't screw down the cap more than about 1 turn) and secure it to the bottle with autoclave tape Autoclave 1 hr at 120°C liquid cycle. Take out of autoclave and leave caps loose. Let them cool and then tighten; if you tighten the cap while still hot, non-sterile air will be drawn in as it cools. Sterilize capped glass bottles in the same way except that you do not need to use the liquid cycle. To sterilize spinner flasks tighten the stirrer unit and center cap in place. Leave the side arm caps loose and secured with autoclave tape as above.
3. Opening Sterile Containers. Always loosen container caps when the container is in the hood and always tighen the caps before removing the container from the hood. When you have open vessels in the hood, never pass your hand over the top of the vessel. It is convenient to place a rack of tubes into the hood and loosen the caps and leave them in place on the tubes so that they can be removed with one hand. When you remove the cap, lift it vertically away from the tube and put it back vertically on to the tube. Do not remove it with an arc motion. Always grab the cap by touching only the top and upper side walls of the cap. Never touch the bottom edge of the cap. It is best if you hold the cap in one hand, use the other hand to transfer liquid, and then replace the cap. If you need to set the cap down, carefully lay it upside down on the hood surface and don't bump into it. When the cap is off, avoid touching the top edge of the container or the cap threading. While you use your other hand to transfer liquid make sure that the hand holding the cap does not leave the hood or bump into items in the hood. If you drop the cap, even in the confines of the hood, you should get a replace it with a sterile cap from an empty bottle.
4. Pipet transfers. Take the cap off of the bottle of liquid that you want to transfer by pipet and carefully place it upside down on the bench somewhere toward the back wall of the hood so that it will not get bumped. Open plastic-wrapped pipets in the hood. Rip the plastic at the end of the pipet away from the tip. Peel the plastic back a bit and insert the pipet into the pipettor. Then grab the plastic wrap near the tip end of the pipet and pull the plastic away from the pipet. It is OK to touch the end of the pipet that goes into the pipettor with your fingers since this part will not come in contact with sterile items. Note, when you peel the plastic back a bit, make sure that you peel it enough so that surfaces of the plastic that you have touched with your fingers are not in contact with the pipet. In this way when you pull the bag off of the pipet, sterile plastic will be dragged along the length of the pipet.
Keep you eye on the pipet at all times so that your sure you don't touch it to non-sterile surfaces like the outside of a tube or the hood bench. If you need to temporarily have two free hands, you can lay the pipettor on the bench with the pipet angling upward. Insert the pipet into the bottle opening making sure not to touch the outside of the bottle or the top edge of the opening. It is OK if the pipet touches the inner bottle surfaces. Suck liquid into the pipet and before pulling it all of the way out of the bottle touch the pipet tip to the inside wall of the bottle above the liquid level to remove liquid that can drip from the pipet tip and with your other hand lift the cap off of the receiving vessel. Remove the pipet and put the tip inside the receiving container. Whether you can touch the pipet to the inner surfaces of the receiving container depends on what you are doing. If you are aliquoting liquid from a single bottle into a bunch of empty tubes, it is OK to touch the pipet to the inner walls of the receiving vessels. If the receiving vessels contain liquid or cells that you don't want to end up in the bottle of liquid from which you filled the pipet then you should not touch any part of the pipet to any part of the receiving vessel. Also, do not quickly squirt the liquid in the pipet into liquid in the receiving vessel since splashing will cause the pipet to be contaminated with liquid in the receiving vessel. If liquid drips out of the pipet while you are moving it from one vessel to another and if the liquids hits the top edge of an open container you must assume that the liquid in that container is now contaminated. This is because part of the drop will run down the inside of the container and part will run down the outside (non-sterile) of the container, and since their is a single drop that contacts both surfaces before the drop splits, contamination is possible. If this happens you have to discard the container that got hit by the drop. This is another reason why you must keep your eye on the pipet during the transfer so that you don't miss drops that hit container edges. Drops that land on the hood bench do not create any problems; however, when done with the hood you must scrub such areas of the hood bench with the green scrubbing pad with some 70% ethanol. When you are done with the pipet, carefully put the cap back on the bottle from which liquid was drawn then pull the pipet out of the pipetor and place it in the white tank. If you have some doubts about whether you touched the pipet to a non-sterile surface, discard it and take a new one; they are not that expensive. Avoid sucking liquid into the pipettor. This will wet the filter cartridge and the pipettor will no longer function. If this happens, disassemble the black nozzle portion of the pipettor (where the pipet inserts into) and replace with a new sterile filter cartridge (these are purchased from Drummond).
5. Storing plasticware. Sterile dishes, tubes, etc... should be stored in your designated area outside of the hood, and they should be left in the bags that they come in with the bag taped shut. This helps to prevent dust build up on the outside of the plasticware (the dust will not get inside the device but dusty items should not be put into the hood since they carry microbes which can obviously spread by becoming airbourne).
6. Checking media. It is always a good idea if you can check your complete media for contamination before adding it to your cells. Simply pipet a small amount of media into a dish and leave it in the incubator overnight, 2 days is better, and check for contamination. By preparing your media in advance of needing it, you can do this check.
7. Pouring liquids. Pouring sterile liquids requires care, and so it is better to do multiple transfers by pipet. If you must pour liquids because the volume is too large, you have to pour the liquid in one motion and make sure the liquid stream goes through the center of the opening in the receiving vessel without touching the top edge of the vessel opening. You must let the liquid stream out fast enough so that it does not drain down the outside of the bottle that you are pouring from. Also, if you don't pour all of the contents of the bottle, when you are done pouring and you tilt the bottle back to a vertical position rotate the bottle 1 revolution before replacing the cap. If you are pouring liquid into a receiver that has virus (i.e. baculavirus) you may get your new media contaminated with virus. In this case, if you only want to pour part of the liquid, pour the amount that you need into a separate sterile container, and then pour all of the liquid from the new container into the receiver.
8. Pasteur pipets. Pasteur pipets are placed in glass canisters with stainless steel caps, and are autoclaved. Each lab worker should have their own cannister of sterile Pasteur pipets, and the cannister should be labeled with the worker's name. To use, place the canister in the hood, remove the cap and place it upside down on the bench toward the back of the hood and out of the way so you don't bump it. Gently shake the canister so that a single pipet protrudes slightly from the canister. Grab the pipet by its end and pull it out of the tube (if more than 1 pipet protrudes from the container, be sure only to touch the pipet you desire, if you touch other pipets they will contaminate the inside of the canister as they slide back into the canister).
Pasteur pipets that are used for drawing up cells (not aspirating but for transferring) must be plugged with cotton prior to autoclaving to protect the cells from contamination. Unplugged pipets should only be used for aspirating. The plastic serological pipets that are purchased should come plugged with cotton. In fact, any device that will be used to deliver sterile solution from the device to the desired vessel must be plugged with cotton.
9. Handling plates. When carrying plates of cells, keep them horizontal and don't slosh them around too much since this may cause fluid to lodge between the cover and the plate. If this happens, just continue to let them grow; they likely will not become contaminated. Again, respect other workers by not handling their plates unless you have permission to do so.
10. Contaminated cells. Once your cells become visibly contaminated, there is nothing you can do to save them. Remove the vessel of contaminated cells from the lab immediately, and add bleach before pouring the fluid down the drain.
11. Splitting Cells. When splitting cells, it is a good idea to leave 1 plate alone as a back-up. If the split cells become contaminated, you can go to this back-up plate and thus avoid going to frozen stock which takes a long time to start up. Put media, PBS, and trypsin-EDTA into the 37°C water bath to warm it to 30-37°C. Avoid having the container in the bath fall over since dunking the cap in the bath may lead to contamination. Wash the bottom half of bottles including the bottom with the squirt bottle of 70% ethanol holding it over the beaker on the floor to collect the ethanol. Put in hood. Put 10 ml of media into the desired number of 10 cm dishes that will be seeded with cells (20-25 ml into 15 cm dishes). To remove the dish lid, grab the lid without touching the bottom edge of the top and hold it in one hand while pipeting the media with the other hand and then replace the lid. Pipet off the media off of the cells you wish to split. Do this by leaning the plate on top of another plate, remove the lid, and hold it in one hand while removing medium with the other. The Pasteur pipet that is hooked up to the vacuum line is place on the bottom corner of the dish where the liquid settles (be sure the pipet tip does not touch non-sterile surfaces). Replace the lid. Add 3 ml of PBS to the cells, briefly swirl gently for a few seconds then pipet off PBS as above. Add 2 ml of trypsin-EDTA, swirl gently for a few seconds, then put in incubator for 1 min. Then quickly check with the microscope to see if cells have been dislodged from the plate (the dislodged cells are sometimes clumped together). If necessary put the plate back into the incubator and check it again with the microscope 1-2 min later. Many cells are dislodged after 1 min incubation with trypsin, but some will require 5-10 min. After a final quick check with the microscope put the plate in the hood and add 2 ml of media to the trypsinized cells. Again with the plate leaning against another plate, draw the liquid up with a serological pipet from the bottom corner and then discharge the solution at the upper corner sweeping across the plate so as to wash the entire plate surface. Do this pipet filling and discharging 6-7 times, and avoid excessive foaming or sucking air through the liquid. With the same pipet transfer the desired volume of cells to each of the plates containing fresh media. Put the cells in the incubator. The trypsin should get diluted a total of 20-fold; a typical split is to trypsinize one 10 cm plate with 2 ml of trypsin-EDTA followed by adding 2 ml of media (2-fold dilution of trypsin). Then 1 ml aliquots are added to four plates each with 10 ml of fresh media (a further 10-fold dilution of trypsin).
Some cells should be split differently as follows (you should find out which way the cells are to be split). Remove media, wash with PBS, and add trypsin as above. After the cells are dislodged, add 2 ml of fresh media and transfer the solution to a 15 ml Falcon tube. Spin the cells at about 500 x g for 2-3 min. Pipet off the media and resuspend the cells in the desired amount of fresh media (do this by pipetting the suspension up and down about 5-6 times, avoiding foaming and drawing air through the solution). Aliquot the suspension into plates of fresh media.
12. Feeding cells: Pipet off the media (see Splitting cells), then add 10 ml of fresh media by pipeting down the side wall of the dish, but be sure not to let the liquid contact the top edge of the dish. Discharge the liquid slowly enough so that no splashing occurs since splashing may lead to contamination. Also do not touch the pipet to the dish since it will get contaminated with cells and then if you reinsert it into the media bottle you will contaminate your media with cells. Some cells like to receive media that is pre-warmed to 37°C while others can tolerate the addition of media that is warmed a bit above room temperature.
13. Centrifuging Cells. Spin cells in a capped Falcon tube at room temperature in a clinical centrifuge for approx. 2-3 min and about 1/2 full speed. If you spin too fast you may rupture the cells; this will be evident if you look at the cells under the microscope after you deliver them to a dish of media.
D. Miscellaneous.
1. Cell Adherence: Some cells lose their adherence to the plastic plate or flask. There is no general rule for dealing with this problem. If you are aware of this problem for your cell line, try to find out what is done about it. Gelatin can be used to increase adherence. Prepare a sterile gelatin solution as follows: Wash some solid gelatin with a few portions of cold water and then add milli-Q water to the gelatin to give a 0.66 % solution, Autoclave the solution to sterilize it and to completely dissolve the gelatin. For a 10 cm dish, add 5 drops of gelatin solution and use a glass Hockey Stick to spread the gelatin over the surface of the dish. Replace the dish cover and let the plate air dry. Rinse the plate with a few ml of sterile PBS to remove gelatin that is not firmly attached to the plastic. You can prepare many plates in this fashion at one time and store them. The Hockey Stick is made by bending a Pasteur pipet about 1.5 inches from the tip into the shape of a Hockey Stick and flame sealing the tip so that gelatin does not enter the pipet by capillary action. Flame the pipet in a small area where the bend will occur to ensure that the part that skims across the dish of gelatin is flat.
2. Freezing and Thawing Cells. You should always have 4-5 vials of frozen cells as a back-up. If someone sends you a single vial of cells, make sure to make 4-5 frozen vial stocks as soon as the cells from the first vial are growing well. To freeze: Trypsinize the desired number of plates of cells and dilute the trypsinized. cells into fresh media Count an aliquot of trypsinized cells with the hemacytometer. Centrifuge the cells and pipet off the media. Resuspend the cells in freezing solution and transfer the suspension to cryo-vials. The cells should be at a concentration of about 3-5 x 106 per ml. You should aim to freeze 1 ml of cells. The components of the freezing solution should be mixed well before adding it to the cell pellet (in this way the cells never see a locally high concentration of DMSO for example). Place the vials in a styrofoam box with a cover (walls at least 1 inch thick) and place the box in the -80°C freezer overnight, and then transfer them to the liquid nitrogen dewar. The styrofoam box provides insulation so that the cells cool down slowly.
Freezing Solution: Most cells can be frozen in growth medium plus 5% DMSO (ATCC uses this for almost all of their cell lines). But, some cells may require different conditions.
Thaw the frozen vial containing cells by incubating in a 37°C water bath, and swirl until it is completely thawed (try to do this as soon as possible). When thawed, transfer 1 ml of cells to 10 ml of media in a flask or dish (In this way the DMSO is diluted 10-fold which is sufficient). The next day, evaluate the condition of newly plated cells .
3. Cleaning Glassware: To clean glassware for use in tissue culture: Soak in liqui-nox bleach (about 50 ml liqui-nox and 25 ml of bleach in about 5 gal of tap water for few hrs, scrub with brush, rinse 7 times with tap water and then 3 times with milli-Q water. Then autoclave (apply autoclave tape as necessary). Use the gravity cycle with a dry time of about 10-15 min. Place in oven if not completely dry.
4. Making Media: Most media is bought from a commercial source as a ready-made liquid form that comes sterile in bottles. Some media such as that use to grow insect cells are made in our lab from powder form since it is much cheaper and we use a lot of this media. To make media, we have a dedicated Nalgen Carboy that is to be used for making TCL media only, and this should be stored in the TCL lab. Before use, rinse the carboy several times with Milli-Q water. There is a special set of glassware stored and kept in the tissue culture lab (the items bear the label TCL). **as Kohei for details*.
When you filter media into storage bottles it is critical that you avoid letting liquid contact that outside walls of the threaded bottle opening. We have had a past incidence where a bottle of media that had liquid on its outer walls became infected with mold on the inside surface of the cap. Mold growth was not detected in the media stored at 4°C, but infection of insect cell cultures appeared 2-3 weeks into the growth of cells. This is time consuming and frustrating because it sends you back to the beginning, and you thus can loose several weeks of time especially since you have to start with frozen cells. So when you fill the media bottles from the tip of the filter, keep your eye on the outlet and make sure it does not wet the outside surface of the bottle. This is very very important!
E. Incubator. Avoid spills in the incubator. If a small spills occurs such that all of the wet areas can be identified, wipe it with a bit of dilute bleach and then with 70% ethanol. You do not have to remove the cells to do this. If a major spills occurs, you will have to do a major cleaning as described below.
1. Routine cleaning. Once per 6 weeks the incubator should be cleaned by the person who has been assigned the job of Tissue Culture Lab. Remove the cells and shelfs. Spray the inside walls of the rack, the inside glass door, and the door gasket with 70% ethanol, wipe, and spray lightly with antimycotic solution. Do the same with the shelfs and then put them back into the rack. Put the cells back into the incubator (you must work fast so that the cells remain outside of the incubator for < 15 min). Discard the pan of water and replace with milli-Q (doesn't need to be sterilized) and add about 15-20 ml of antimycotic solution.
If the incubator shows signs of infection (visible or it smells), remove the shelf and racks, wipe them with water, and autoclave them. Let them cool in the autoclave. While autoclaving, wipe all surfaces of the inside of the incubator and the door and door gasket with 70% ethanol and spray lightly with antimycotic solution. Put the cooled rack into the incubator and clean treat with ethanol and antimycotic as above. Do the same with the shelfs and then place them back into the incubator. Replace the water as above.
F. Reagents:
Anti-mycotic solution: Dissolve 3.88 g of p-hydroxybenzoic acid n-butyl ester (Sigma Cat # H9503) in 526 ml of 95 % ethanol, then add 474 ml of milli-Q water (add water after the solid has completly dissolved in the ethanol). Store in glass bottle at room temperature. No need to autoclave the storage bottle since the solution has lots of ethanol.
Gelatin: Sigma G9391
Glutamine: This needs to be added to media stored at 4°C every 2 weeks. Sterile glutamine (200 mM) is aliquoted into sterile tubes and stored at -20°C. Thaw what you need (do not refreeze), and add to media. Keep track of the dates of glutamine addition on the media bottle.
Media: Store media as it comes from the vendor at 4°C (observe the expiration date, GIBCO claims they have tested their media after the expiration dates and it performs less well). Complete media (i.e. with serum, antibiotics, etc...) can be stored frozen, and a working solution can be stored at 4°C for up to 1 month (if glutamine is needed, be sure to add it every 2 weeks). Frozen media that is thawed for use should not be refrozen.
PBS: All chemicals are reagent grade. For 10 liters: 80 g NaCl, 2 g KCl, 21.6 g Na2HPO4-7H2O, 2 g KH2PO4. Use milli-Q water, adjust to pH 6.8 with HCl and top off to 10 liters with milli-Q water. Autoclave it in glass tissue culture bottles. Store room temp.
Penicillin/Streptomycin/Fungizone: (JRH Biosciences 59606-77P). Store -20°C. Make sterile aliquots, store -20°C. You can thaw a bottle, prepare sterile aliquots, refreeze once. If needed, keep a working aliquot at 4°C for up to 2 weeks and then discard.
Sera: Store -20°C. You can thaw a bottle, prepare sterile aliquots, refreeze once. You may want to store frozen stocks of various volumes in case you need only a small amount since you don't want to freeze/thaw multiple times. Some sera come heat inactivated. If not heat inactivate a bottle by first thawing it in a 37°C bath and then placing it in a bath at 56°C +/- 1°C for 30 min (be sure bath water level goes above the level of serum in the bottle but not above the neck of the bottle. Aliquot the heat inactivated sera and freeze.
Trypsin/EDTA: (GIBCO/BRL Cat 25200-015). Store it -20°C. You can thaw a bottle, prepare sterile aliquots, refreeze once. If needed, store a working solution at 4°C for up to 2 weeks.
G. Supplies:
Cell Scrapers: (Costar 3008, individual wrapped, 100, 1-800-492-1110).
Cryo Storage Vials: (Nunc, from VWR).
Detergent: Liqui-nox (VWR 21837-027, case of 4 gallons).
Dishes (Culture): 10 cm diameter Falcon 3003 (VWR 25382-166, case of 500).
Falcon Tubes: J-wing stockroom.
Microscope Lamp: 6V 20 W bulb (Cat # B620A ) Bartels and Stout, Inc. P. O. Box 1994, Bellevue, WA 98009, 206-453-1705
Pasteur Pipet Canisters: Belco cat # 1290-38300
Pipets (Serological): Individually wrapped, cotton-plugged:
1 ml: VWR 53283-264, case of 1000
5 ml: VWR 53283-344, case of 200
10 ml: VWR 53283-366, case of 200
25 ml: VWR 53283-377k, case of 200
CO2: From University Stores, 50 lbs, size 6, 8709210. Note it takes a fews to get the tank so there should always be a full tank in addition to the one in use.
Bleach: purchase at grocery store since stockroom bleach is expensive.