In a large lab populated by people with very different backgrounds and levels of experience in various techniques, it is necessary to establish standard methods for carrying out important laboratory procedures. Since everyone in the lab uses the same chemicals, solutions, and so on, it is essential that everyone use the appropriate techniques to ensure that our stocks and instruments are not contaminated. Thus, there are no exceptions to the following. This may sound heavy-handed, but past experience has shown that no one feels that way after they have lost an
important sample due to their own or someone else's carelessness (lack of concentration) or arrogance ("I know how to be nuclease-free").
The primary worry in this laboratory is that ribonucleases (RNA nucleases) found in the oil on your skin will find their way into our solution samples. These nucleases destroy the mRNAs and tRNAs and rRNAs used in many experiments done in the lab, and nuclease contaminations have cost us many dollars and much time in the past. Thus, as far as lab procedures are concerned, THERE IS NOTHING MORE IMPORTANT TO THIS LAB THAN EVERYONE USING PROPER NUCLEASE-FREE TECHNIQUES. This is true for both those working on secretion/integration projects
and those working on clotting projects. Chemicals and Biochemicals: Under normal circumstances, NOTHING should be inserted into a chemical or biochemical bottle or container. To get material out of bottle, wipe the dust off of the bottle top and shoulders, and then pour (tap the outside of the bottle if necessary) the material onto the weighing paper or boat. Be careful not to pour out too much, because nothing should be placed back in the bottle (except in extraordinary circumstances and when AJ is told). If too much comes out of the bottle, simply throw out the excess. In some cases, it is necessary to use a spatula to break
material into smaller chunks. Also, with fluorescent dyes, antibiotics, etc., you may need a small spatula to obtain mg quantities. In each of these instances, the spatula must be cleaned with water and acetone thoroughly, then dried, using nuclease-free techniques, with a clean kimwipe before use. Pipetmen and tips: Remove tips from container using the micropipet barrel: do not use your hands. If the tips have been or may have been exposed to finger oils, then mark container as NNF=non-nuclease free. NF=nuclease free. To obtain accurate volume measurements and deliveries with the pipetman, always (i) suck up the desired aliquot into the tip, (ii) release the aliquot back into the stock solution, and (iii) refill the tip again. The second aliquot will have the correct volume when it is delivered. If you do not wet the inside of the tip first and then fill it, the volume that is delivered will be less than advertised because a thin film of solution remains on the inside of the tip after delivery. NEVER STICK A MICROPIPET INTO A SOLUTION BOTTLE!! Even if you think you can do this without touching the pipet (where everyone's hands have been) to the bottle opening, do not try it. Instead, pour some of the solution into a test tube and then use the micropipet. Never take chances with stock solutions: the time you think you save is not worth the risk of having yours or someone else's experiment fail because of a nuclease contaminant. It costs more than $100.00 to repair and recalibrate a micropipet, so please do not drop them, twist barrels too vigorously, or let solvent get inside the hole at the end of the barrel. YOU (please do not assume that someone else will do this for you) should routinely check micropipets as you use them: if solution leaks out of the tip when you hold the pipet vertically, or if there are always air bubbles at the tip, the pipet needs to be repaired. If you suspect that a micropipet is not working properly, then you should check how accurately and
reproducibly it delivers water by measuring the mass of the water placed on weighing paper using analytical balance. Please bring any faculty micropipet to the secretary, with a note describing its problem, so that the faculty micropipet is not used in any experiments. The micropipet will then be sent to the company to be repaired. Microcentrifuge tubes: Do not open tubes with your hand. Always use a plastic lever to open a microfuge tube, and thereby avoid getting finger oil on the inside top of the cap where it can contact the sample. Ask questions: Please ask questions without hesitation, so that you fully understand why and how something is done. Do not assume that you will be able to "figure out" how to do a procedure or to operate an instrument properly if you do not understand it. It is very important for the safety of both people and their experiments that you do not "guess" how to do something amid then end up inactivating some costly samples, solutions, glassware, or equipment. Our biggest problems in the lab have always been caused by someone assuming that he or she knew how to do
something when they in fact were just guessing, hoping, and /or afraid to ask. As for equipment, one should always read the owner's manual before using an instrument. If the manual does not provide the necessary information on a particular issue, or if it is unclear, then consult the list posted in rooms 107, 114, and 116 that identifies the person in training needed, or will send you to someone who can solve your problem. Priorities: Whenever you are doing an experiment, your first priority is to do the experiment accurately and quickly (e.g., so the enzymes don't die). Thus, under no circumstances should you answer the phone, answer someone's question, or otherwise get distracted while you are doing an experiment. It is OK to be rude. Or more accurately, to ignore someone who is rude. Everyone who is a researcher should know better than to interrupt when someone is in the middle of a procedure, but some apparently do not understand that doing research successfully requires total concentration.
If someone really needs to talk to you, they will call back or come back later. Keep the doors to the labs locked: Always check and lock all of the doors if you are the last person to leave at night. If you are the only person in the lab, it is best to keep the doors to all of the labs locked, simply to minimize passer0by access, especially to empty rooms. Do not lend you lab key to anyone: if someone does not have a key, then they are not supposed to have access to our labs; if they think that they should have access to our labs, then they can come to me and explain their needs. Radiation Safety: Call Radiation Safety to pick up waste when necessary. If there is a spill of radioactive materials, isolate the area (use signs), contain or clean up the spill, and alert AJ and the Radiation Safety office (5-1361). Equipment Malfunction: If a piece of equipment breaks or stops working when you are using it, take care of it right away. Do not let it sit until you or someone else needs it again before fixing it. First tell the person in the lab who is responsible for that instrument (consult list in rooms 107, 114, or 116), and let them arrange for repairs. If that person is not available and if the instrument is on a service contract, then call the company yourself. If the problem is very serious (time or money), or if the item is not repaired as soon as it should be, then tell AJ. Facilities problems: If there is a problem with the cold room temperature, with burnt-out light bulbs, with the hoods, electricity, plumbing, or whatever, call George Martin (5-7902) to report the problem. If the problem is not corrected promptly, then alert/ call AJ. Supplies: Items that need to be ordered should be listed on the Board in 114. Please list part number (it is best to get this from the old container before throwing it away), amount, supplier, and unit price. Also, before ordering, please check with others in the lab to see if they need anything from the same company; this will minimize our freight costs. Routine orders to replace materials should be made soon enough to avoid running out of something before the replacement material arrives. DO NOT PUT AN EMPTY OR NEARLY EMPTY CONTAINER BACK IN STORAGE WITHOUT ORDERING A REPLACEMENT. Check with AJ before ordering any expensive or unusual items; we may already have them. If you place an order, always ask what the price is, when the material will be shipped, and who you are talking to. Write this information down on the order sheet. If the price quote is not what you expected from the catalogue (note: Fisher, VWR, and other large suppliers should give us a minimum of a 10% discount on the catalogue price), then either you or the person on the phone used the wrong part number and you must correct this. If the material is back-ordered or not in stock, you may want to order it from a different source. General supplies should usually be ordered from
Fisher or VWR, whichever is cheaper. Some materials must be ordered only from particular suppliers. If unsure, ask AJ. Packing Slips: When and order is received, the packing slip should first be checked with the contents of the package to ensure that we have received all that was ordered and listed on the packing slip. If there is a mistake or a partial order was received or the contents were damaged when received, make a written note of the problem on the packing slip. The packing slip should then be dated and placed on the secretary's desk. Storage of samples in liquid nitrogen "refrigerator": Freeze samples first in liquid nitrogen. Before inserting a sample storage tube into an aluminum cane, squeeze the cane edges together slightly in order to ensure that the cane will hold the tube tightly. Tubes should be put onto canes as close to the bottom of the cane as possible. This means (i) tubes will be more likely to remain in the liquid N 2 as the level lowers, and (ii) tubes will tend to hold each other on the cane. Do NOT force canes into canisters! That results in tubes being knocked loose and lost from canes already in the canister. Your sample could be next. Storage of samples at -20ºC and -85ºC: Freeze samples first in liquid nitrogen. Please open freezer doors only long enough to get out or put in your samples, and do not open the doors unnecessarily. Let's try to lengthen the time between defrosting "parties". How many times has a sample been frozen and thawed? Whenever a sample is thawed, you should ALWAYS place a mark on the outside of the tube with a black making pen. The number of marks on the tube will then tell you how many times the material in that particular tube has been melted. This is very important when evaluating the success of an assay or titration, because many proteins are inactivated by repeated freezing and thawing. Of course, marking this mark is of no use unless you also always write down in your lab notes the number of times that a sample has been melted. Centrifuge rotors:Every centrifuge rotor should be washed and dried after each use. If the rotors are not cleaned, they will become pitted and corroded inside the tube holes, and then the tubes will break (and your samples will be lost) when they are pushed down on an irregular corroded surface at high speed. Since the ultracentrifuge rotors costs $8,000- $10,000 each, we cannot afford to ignore proper cleaning procedures. Rotors should be stored in the cold room. If a run is to be made at a temperature higher than 4ºC, place rotor at room temperature well ahead of time to warm it up. Please pay attention to maintenance procedures: O rings on tube caps and rotor plates should be coated with a thin film of vacuum grease; screw-on caps should periodically have a small amount of Spin-kote applied to the threads to ensure easy tightening and loosening (if you add too much, the excess graphite will end up in your sample). Centrifugation with open tubes: Whenever you are going to sediment a sample using swinging-bucket centrifuge tube that does not have a cap, you are going to sediment a sample using a swinging-bucket centrifuge tube that does not have a cap, you should nearly fill the tube (e.g., to about 3-4 mm from the top of a SW 28 or SW 41 tube). If there is not enough fluid in the tube, the tube may collapse at high speed and you will not get either your sample or the tube out of the rotor. Cleaning cuvettes:
After use, cuvettes should be rinsed with water, especially those which have contained radioactivity and /or protein. Cuvettes that have contained phospholipid must be rinsed in the hood with water (twice), cholorform (twice), ether (twice), acetone (twice), and water (twice).After this rinse, cuvettes should be gently lowered into a beaker containing Chromerge (room1070. Use pasteur pipet, if necessary to remove air bubbles from inside cuvettes.Cuvettes normally soak overnight in Chromerge. A shorter time may result in incomplete cleaning, but we have not really tested this, and a shorter time has been used in the past without any apparent problem. A longer time can result in acid-etching of the cuvette surface, especially if the cuvette stays in the Chromerge constantly. Hence, it is good practice to clean and dry your cuvettes (steps 4-9) the day after you use them.Following the Chromerge soak, pour Chromerge out of the beaker containing the cuvettes into an empty beaker, using your fingers (with polyethylene glove on) to make sure cuvettes do not fall into sink. To rinse acid off of cuvettes, gently (to avoid splashing) add water to beaker containing cuvettes and then pour off. Repeat until no yellow-colored acid is obvious on outside of cuvettes.With clean polyethylene gloves on, rinse cuvette(s) with distilled water at least 50 times. (Ann Dell found that 100 rinses were required to avoid artefacts, and I encourage you to rinse 100 times to ensure that all of the Chromerge is gone.) NOTE: BE CAREFUL. HOLD ONTO CUVETTES TIGHTLY, to avoid slippage and breakage. NOTE: A "rinse" only counts if all of water is poured out of cuvette before the addition of fresh water.Rinse each cuvette ten times with doubly distilled water.After shaking excess water out of cuvette, dry outside of cuvette with clean kimwipe and invert cuvette onto a clean piece of aluminum foil and kimwipes to let it drain.With a clean pasteur pipet on nitrogen gas tank, blow each cuvette dry.
Place cuvettes in rack to minimize chances of breakage during transport or storage.
Cleaning Microcells: The cleaning procedure is the same as for other cuvettes, except that the larger surface area per volume inside the microcell makes it essential that you do not cheat on the number of legitimate rinses of the cell (i.e., do not use less than 50). Since it is difficult to fill microcells completely (bubbles tend to form in cells unless water is added down one side of cell) and to empty them completely, it is best to empty microcells by using a teflon-tube-tipped pasteur pipet connected to the water aspirator ( using a trap ) to provide vacuum. If you use this approach, BE SURE that this teflon tip is kept clean and nuclease-free: do not allow it to touch other things, and store it in a protective test tube. Cleaning Cuvette Caps:
Cuvette caps should be rinsed with water before being placed in a micro soap solution.Soak in soap solution. As with cuvettes, it is probably best to do steps 3-5 the next day rather than letting the caps sit a long time in the soap (especially if the water evaporated out of the beaker and the soap dries on the caps).Using a polyethylene glove to hold caps, wash off soap using tap water. Then rinse caps with large amount of distilled water before rinsing them with acetone from the squirt bottle.Dry with clean kimwipes or blow dry with nitrogen gas (Kimwipe may leave fibers on the caps).
Store clean cuvette caps in a nuclease-free plastic tube with a cap.
Washing glassware and plasticware:
Pre-clean, by rinsing thoroughly, any item that was exposed to radioactive material, proteins, or unusual (=non-aqueous, such as dye reagents) material, before placing item in wash bucket. Also, remove all tape, residual parafilm, etc. from the item. Be sure to immerse pipets and glassware totally in the soap solution, but do so carefully: it is expensive to replace items.Please avoid letting soap dry onto glassware or plasticware. Keep large containers filled with water until washed; keep small containers totally submerged in wash bucket.Rinse blue dishwasher racks with distilled water. Items should be placed on racks rather than on sink during washing procedures.Rinse each item thoroughly with tap water to remove all soap. This means that the item should be rinsed until no more soap suds are visible, and then some more. As a guide, in the case of small-volume items (up to 50mL), "thoroughly" means completely filling and emptying the container a minimum of 10 times. For item 50 to 400mL, a minimum of 5 times. Larger items (e.g., 4 L Flasks) should also be rinsed at least 5 separate times, using enough water and swirling to rinse inside completely. Then place item on blue rack.After accumulating almost a full rack of items, put on a clean pair of polyethylene gloves and rinse each item with distilled water a sin step 4. Then place item back on rack to drain. Be sure to keep track of items, so that each receives a distilled water rinse.Change the water in the glassware soaking bucket and add Micro soap (~ 20mL) after each wash.Washed volumetric flasks and plasticware should be left to dry, and should then be treated as in steps 8 to 10. Other items can be readied for baking after draining.Cover openings on each container with aluminum foil, placing shiny side against the container. Use sufficient aluminum foil to cover the opening and then some. Do not cover the glassware with foil until it is dry, unless you plan to bake it that day.Items are baked at a very high temperature in order to destroy any nuclease enzymes that may have been transferred to its surface. For baking place glassware in oven and bake overnight at highest temperature (a setting of 9 on oven). NOTE: For baking loosely-capped bottles or bottles with plastic seals around rim, use a temperature setting of 6 on the oven.
After baking overnight and then cooling the items, distribute them to the appropriate storage locations in room 112.
Cleaning large (1mL-25mL) glass pipets:
Rinse glass pipets after use with water and /or acetone, if necessary (e.g., if there is fluorescein solution along pipet walls).Immerse pipet into soap bucket (containing Micro soap) with tip pointing up, and let soak.Using polyethylene gloves, transfer pipets from soaking bucket to washing frame (keep tips up), and insert loaded frame into pipet washer.Run cold water through pipets in pipet washer overnight to rinse out all soap.Fill up washer through distilled water and then drain. Repeat three times. Then fill up and let pipets soak in distilled water for an hour or so. Repeat.Using clean polyethylene gloves, CAREFULLY distribute pipets into appropriate pipet cans. If possible, fill cans only 1/2 - 2/3 full.
Bake overnight at a setting of 9 on the oven.
Use of glass micropipets:
NEVER touch the tip or barrel of a glass micropipet. Touch the micropipet only at the end where the numbers are printed on the barrel.Micropipets should NEVER be placed on a lab bench or other surface without proper precautions. If pipets need to be put on a surface, they should be placed on a clean sheet of aluminum foil.Clean micropipets properly if you use proteins or phospholipids or membranes; see 1 below.Place micropipets into cleaning solution gently after use. Do not force them into beaker because the glass wool does not compress very much, and you may therefore break the end off of the micropipet by pushing too hard.Please keep micropipet tips pushed up to the end of the storage containers, so that the pipet barrels do not end up adjacent to the places on the micropipets where people touch.
The tips are easily broken, so please be careful. Always check a micropipet before use to ensure that its tip is intact. Open and close the micropipets drawers gently: avoid having the micropipets slam their tips into each other and into the end of their container.
Cleaning of glass micropipets:
After use, every micropipet that has been exposed to either protein or to phospholipid or to salts that precipitate must be rinsed immediately with soap. Four to six up-and-down rinses of the insides of the micropipet with soap solution should be sufficient. Micropipets exposed to protein should then be immersed in the Chromerge or No-chromix (acid) solution. Micropipets exposed to phospholipid should be: (i) left in the soap solution, (ii) cleaned as soon as convenient in the hood with successive rinsed (use a rubber bulb, but be careful: do not let solvents get into rubber bulb and then drain back through the micropipet) of water (to times), chloroform (two times), ether (two times) and acetone (two times), and the (iii) immersed in No-chromix or Chromerge. Micropipets should not be left in the soap solution longer than overnight. Micropipets exposed only to salt solutions or to nucleic acids can be immersed in No-chromix or Chromerge for at least 8 hours (an arbitrary guess on my part; if necessary, use less time) before proceeding to the next step.With used polyethylene gloves on your hands, transfer micropipets that are to be washed (the apparatus handles 18 micropipets per load) into the plastic wash container. Tip container so that the micropipets rest on their bases (not on micropipet tips), and run tap water into and out of the container for a minute or so. This should wash acid and radioactivity off the outside of the micropipets.Turn on hot water tap until you get hot water. Pour enough Micro Soap into the bottom of the "SOAP" cleaning bucket to barely cover its bottom. Run hot water into bucket until level is about 1/2 inch below top of container. Blow off suds.With used polyethylene gloves on hands, transfer micropipets into base of washing set-up. Note: do this over the bench, because acid may leak out of the bottom of the base during the loading (and possibly onto you!). Mix sizes, if possible, so that base contains both small and large volume micropipets. Put each micropipet in base securely; there is no need to push the micropipet in 1 or 2 inches. Make sure that each bent tip is pointing towards the middle of the base.If you must set the base down, be sure that it is placed at all times so that no micropipet touches anything, including the bench top. In all of the following, make sure that the micropipets are kept clean.Attach base with micropipets (held vertically) to hose connected to aspirator. Turn aspirator vacuum to maximum. The No-chromix or Chromerge should be sucked out of each micropipet, and then out of the base.If No-chromix or Chromerge is not sucked out of a particular micropipet, brush its tip with clean kimwipe. This will dislodge any particle at the tip opening that may be blocking the micropipet. If the micropipet is still not sucking, place it back in the acid to soak. (If additional acid treatment does not open up micropipet, throw it away.) Replace the plugged micropipet in the base with another micropipet for cleaning.Immerse micropipets into soap bucket. BE SURE that tips are centered over bucket, so that no micropipet is sticking outside the bucket as you lower the base. Before completely immersing micropipets, make sure that each micropipet is sucking up soap solution. This is best done by dipping tips in and out of soap until you have seen each micropipet suck up solution. If a micropipet is not pulling soap solution, do step 7.While the soap is running through the micropipets, add 0.1mL (about 3 drops) of 1 N HCl to the "FIRST RINSE" bucket, and fill bucket to 1/2 inch below its top with distilled or nanopure water. This acid rinse is supposed to remove all of the soap from the micropipets.When the soap wash is completed, you will hear (in absence of other noise) a different sound from the aspirator because air will be going through the micropipets instead of fluid. Remove base from wash bucket and hold base with micropipet tips up until all fluid is sucked from base. Then rinse outside of micropipets with distilled (if possible) or tap water to remove all of the soap. Also rinse soap off of base where suds may have gathered. NOTE: be careful not to touch micropipets to water faucet or anything else. NOTE: We do not want soap dried
in the inside of the micropipets, so it is important that you do not let very much air run through the micropipets at the end of the soap wash. Please stay close by and change micropipets to the acid wash bucket when appropriate. In subsequent steps, it is OK to let air go through micropipets and you can change buckets when it is convenient.Immerse micropipets in acid wash. Be sure that all micropipets are pulling solution. If not, repeat step 7.Rinse soap bucket thoroughly (i.e., rinse along all sides for 2-3 minutes) with hot water before storing it or using it again.Fill "SECOND RINSE" bucket with distilled water.At end of acid wash, lift base out of bucket, invert to drain fluid out of base, and transfer micropipets to "SECOND RINSE" bucket.Rinse the "first rinse" bucket thoroughly with distilled water. Then fill it with distilled water for the "THIRD RINSE".After third rinse, drain fluid from base and then transfer micropipets to "FOURTH RINSE" bucket.Add acetone to a 1L beaker until volume is about 300mL. The acetone is used as a final rinse to remove organics and water, and thereby to speed the drying process.At end of the fourth rinse, lift base out of bucket and invert to drain all water out of base.Immerse micropipet tips in acetone, making sure that all tips go inside beaker. Be sure that acetone is being pulled through each micropipet.After a total of 200mL of acetone has been sucked through the micropipets, remove from acetone, and invert base so that all acetone is drained out of base.Find a clean spot on lab bench and place base with micropipets on bench. Let sit for 15 minutes or so with aspirator running to dry micropipets. Pull base off of aspirator hose before turning off aspirator (so water does not come back through hose and into micropipets). NOTE: it is VERY IMPORTANT to arrange base and hose so that the hose does not pull on base and cause it to tip, thereby letting micropipets touch bench. I suggest using buckets to stabilize the base. You may also want to put clean aluminum boil under micropipets. Also do not put
micropipets close to sink while drying, because someone using sink may carelessly splash water onto the clean micropipets.Rinse the "third rinse" and "fourth rinse" buckets thoroughly with distilled water before storage.
After drying is complete, using a clean glove, place micropipets gently into the appropriate storage location in drawers.
Ethanol precipitation of RNA and DNA: RNA and DNA can be precipitated out of solution by first increasing the salt in the solution to between 0.3M and 1.0M (this reduces the amount of water available to solubilize the RNA or DNA), and then adding ethanol so that the final solution is 66% to 75% (v/v) ethanol. In most cases, we add an equal volume of 2.0 M KOAc (pH 5.0) to a sample (this gives up 1.0 M salt) and then add between 2 and 3 volumes (based on the total aqueous volume of sample + salt) of ethanol. for example, if I had 2mL of sample, I would add 2mL of 2M KOAc (pH 5) and then 12mL of 100% ethanol. Variables: The most important variable is the concentration of the RNA or DNA in the sample. If your concentration of RNA is too low, then you will not get efficient precipitation, nor will you get precipitate particles that are large enough to collect easily by sedimentation or filtration. The minimum concentration varies, depending upon the length of the NA, but the minimum A 260 for efficient precipitation of tRNA is above 0.1. It is best to have an A 260 of 1 or greater in your sample before precipitation. If you have low concentration of NA, then you should use a higher concentration of salt and of ethanol. If your sample is fairly concentrated, then you can use a final salt concentration of 0.5M and add only 2 volumes of ethanol (to 66% v/v) to get efficient precipitation. Also, remember that these values do not need to be exactly. If adding three volumes of ethanol would make your sample volume too large to fit into a particular ultracentrifuge tube, then add less ethanol (or salt; but not less than 66% or 0.3M, respectively) so that you can collect all of the precipitate in the
same tube in one pellet. Careful: If you wish to precipitate RNA that has been purified from an ion exchange chromatography column or the equivalent, remember that the RNA is already in high salt. Therefore, you usually do not need to add any salt (or at least not much); simply add ethanol to the sample.
If you use the maximum precipitation conditions, 1.0M salt and 3 volumes of ethanol, some salt is also likely to precipitate out of solution. Therefore your pellet will look very white, and it may also come loose from the tube as you are pouring off the supernatant. Therefore, be very careful when removing the supernatant: pour it off slowly, and hold the tube so that the pellet is on the top side of the tube as you pour. Then if the pellet breaks loose, you will have a chance to stop pouring and save it. If the pellet does break loose, then you will need to repeat the
centrifugation. This problem with salt coming out of solution is one reason to maintain a high concentration of RNA or DNA in your sample: then you can use less salt and less ethanol to precipitate the NA.