Chin-Sang Lab Manual


General Lab Rules

      All solutions, after autoclaving or filter sterilizing, must be labeled with tape and should also contain the date.  Even flasks of ddH2O and other liquid media must be labeled (even by WHMIS standards) so that we are certain what is inside.  It is not enough that you know, particularly if we are trying to throw out old solutions to free up glassware, everything must be labeled.

     All poured plates should be labeled with the appropriate colour code, except worm plates.  All antibiotic plates should be bagged and stored at 4 degrees within the next 24 hours of pouring to ensure their effectiveness.  All drop-out plates (-trp etc.) should be bagged and refrigerated in the next day or two, once dry, to avoid contamination.   Bags from when plates are prepared, should be kept, particularly the large bags, for storing plates later.  Large volumes of small worm plate bags can be thrown out, but some should be left for the storing of small drop out media plates.

     Worm plates, in general, are prepared by work study students, or technicians, as needed, at least 3L a week.   It is also the technicians responsibility to ensure that all components of worm plates (plates, solutions, chemicals) are prepared for 3 litre aliquots, and that they are well stocked.  If any unusual use of worm or other plates is needed, the person requiring them should prepare them themselves.  Anyone doing a mutagenesis or large worm or yeast screens, should prepare plates for such use, the exception being post-docs, who can make special requests for plates from the technicians.  If you have any questions about the proper pouring of plates, or the use of the PourBoy III, please ask the technicians for help.

     A PCR sign-up sheet will be provided during busy periods.  Please indicate the approximate time, your name, and if you need two machines please indicate this as well.

     The spectrophotometer, PCR machines, weigh scales, ph meter, and the PourBoy III, should be turned off, or on standby when not in use.  Try not to turn the spectrophotometer on and off without a rest between, and keep a "zero" or "blank"  parafilmed for later use and turn the machine off, if you won’t be using it for a few hours.  The pH meter must be rinsed with ddH2O before use, and the vent opened.  The vent must be closed after use, the meter rinsed again with ddH2O, and stored in the ph7 (yellow) buffer to keep from drying out.

     The technicians are responsible for very general lab solutions (2XTY, M9, Freezing Solution), other more specific solutions should be made by individuals as needed.  This is important not only in the full understanding of your protocol and project, but it also helps with trouble shooting, if your own copy is contaminated or deficient.

     The presence of many fungi and contaminants in the lab air means that sterile technique should be employed when opening any stock solution at a bench, even if it isn’t a rich media.  Try to keep the bottle near a flame while opening, use sterile tips, and try to flame the opening and lid before closing.  Rather than be too concerned with flaming, be sure to minimize the time the lid is actually off the bottle, the less time it is off the less the chance a contaminant will fall in.

Queen’s plasmids

      Once a gel confirms a new clone, two of the best isolates should be reprepped immediately from the left over culture using new Qiagen columns.  They should also be assigned a pIC number, followed by an isolate number or letter.  The Qiagen preps should then be put into the pIC stock boxes, and all other isolates should be stored in the back up boxes or your own personal box.  Sheets of the pIC clones should be printed, with the full description of the cloning process, and a concise map of the relevant details drawn roughly beneath.  If for any reason you notice a clone in the stock with very little DNA present, retransform it, use a new Qiagen column, do a diagnostic digest, and return the new prep to the stock.  All of the details, which plate you will need, which enzymes to digest, will all be found on the queen’s plasmid data base, and ideally, in the book as well.  Do not for any reason use the last microlitre of stock plasmid, retransform it immediately. 

 IC Strains, CGC and other lab strains

       All strains constructed  within the lab should be prepared for freezing and as frozen given an IC name from the queen’s strains database. Alleles isolated in our lab are given a qu designation. For C. elegans nomenclature see here.   Tony is responsible for the organization of the IC database, but not necessarily the actual freezing of the strains.  If you are generous enough to do the freezing and naming yourself, please ensure that Tony is aware that there is an IC strain in the –80 which will need organizing in the proper box.   All CGC and other lab strains are the responsibility of Jeff, from freezing to final organization.

     To freeze a strain it is best to have two plates full of newly hatched L1 larvae (it is usually optimal for freezing when a few eggs are still visible and the L1 larvae are distributed throughout the plate).  Chunking two similar chunks to two plates with a large bacterial field will usually produce this result.   You must monitor the plates for a few days afterward as different strains will grow at different rates, so that the optimal stage isn’t missed.  When ready, label your cryogen tubes first (you will need four tubes for two plates of each strain).  Add 2mls of M9 to each of the two plates, and consolidate the 2mls from the first plate to the second washing all of the worms onto one plate.  Set your p1000 to 800ul now, (this will make up for liquids lost in the agar) and wash 800ul of worms into 4 separate cryogen tubes.  Once all of the tubes are full for all of the strains get the freezing solution from the 4 degree, ensuring MgSO4 has been added.  Add an equal volume freezing solution to the M9/worm mixture (800ul) and cap tightly.  Quickly collect the tubes into Styrofoam, bind with elastic, invert a few times to mix, and put into the –80 as quickly as possible as the freezing solution is not optimal for the worms to sit around in.  Once the worms have frozen in a day or two, make sure that they are placed in the proper box and that the Styrofoam is removed from the –80 to make room for future freezings.

  Dishwashing

      Everyone should be responsible for the washing of any glassware they use for making solutions, for growing cells, or of containers they no longer need.  Any flasks containing live culture, or contaminated solutions, must be bleached before disposal in the sink.  All glassware should be washed with Alconox, rinsed three times with hot water, and then rinsed three times with the distilled tap water.  This removes all traces of contaminants as well as the hard water from the regular tap.  The autoclaving of washed dishware is the responsibility of the technicians.  Bottles are to be autoclaved with loose caps and a piece of autoclave tape from cap to bottle.  Flasks are to be covered in foil and a piece of autoclave tape for autoclaving.  All dry goods are normally autoclaved 40 minutes, with 15 minutes drying time (cycle 4, in the 2ndfloor autoclave).

Miniprep columns

      New columns should be used for preparing stock plasmids, purifying final Yeast 2 Hybrid plasmids, and PCR purifications and gel extractions.  Please Note: You must use the Qiagen solution when using the Qiagen columns. Do not use our homemade solutions on new columns.  Immediately after use, these must be washed twice with ddH20 and once with 95% ethanol HPLC grade.  These are then to be put into the CLEAN boxes of our glass milk column stock.

     All old columns, after use in the glass milk protocol, should be put into a DIRTY box and not mixed with the cleaned columns.  These will need to dry and be blasted with air before they too can be cleaned twice with ddH2O and once with 95% ethanol.  These are then put into the CLEAN boxes behind the manifolds.  If during the cleaning process any older columns do not empty as the others, they should be thrown in the garbage right away, as they are not suitable anymore for preparing plasmids.

     The manifolds should be checked regularly that they are not overly full of liquid, especially after cleaning many columns.  Spill over into the vacumn flasks will mean unnecessarily cleaning, more often, unless the manifolds are emptied before they do this.

     No columns should be thrown away unless they have been reused, and are no longer capable of allowing liquid to pass through.

 Culture Tubes

      All old culture tubes should be stored on the tops of shelves until enough have been collected to make up at least one rack.  The liquid from the used tubes should then be poured into a flask with bleach, and the glass disposed of in the glass container.  The lids are to be gathered into a tub for washing.  New racks of culture tubes are prepared from brand new culture tubes, and topped with washed lids or new lids.  These must be autoclaved before use, on dry cycle, 40 min., 15 min. dry.  These are then stored on the lower shelf of the 2nd bench.  Only autoclaved tubes should be placed there.

 Gel boxes

      A litre of 10x TBE should always be prepared and ready for when the carboy empties.  Dilute the litre of 10x TBE with nine litres of distilled tap water, being sure not to lose count.  Gels are typically made in 100ml aliquots, a 1% gel being 1g of agarose in 100mls of TBE.  After microwaving  (55 seconds) and the gel has cooled slightly, 4ul (10mg/ml) of EtBr can be added.  The gel is typically ready to run after 15 minutes in the 4 degree.  Rinse combs under the distilled water tap after pulling them from the gel and replace in the comb box.  Once finished, first time gels can be put back into flasks for diagnostic gels.  If you are running a gel to gel purify DNA, you must use new agarose and TBE in the gel and in the running buffer.  Once done with gels, if you are not reconstituting them, throw them in the bin in the fume hood to dry.  Used running buffer is temporarily collected in a beaker near the gel boxes and should not be confused with the unused running buffer.  Once the used running buffer beaker is full, it can be emptied into the large beaker in the fume hood.  Once the large beaker in the fume hood is full, bleach should be added, leave it to sit for an hour or so, and then this can be poured down the sink with running water.  Gels that have been dried in the fume hood can be deposited in the ethidium bromide disposal basket.

 Microscopes and slides

Microscopes: No person will be allowed to use the Axioplan 2 or Axiovert unless personally trained by  Ian or by someone  Ian delegates to train.  Tony is our OIT funded resident microscopy maintenance person and will make sue the microscopes are clean and covered when not in use.  Report any problems to Ian immediately. 

Notes: make sure your are using the right immersion oil. Do not mix oils and DO NOT  use the halocarbon oil (only for injections) on the Axioplan 2.   Gently wipe off excess oil the objective after each use and check surrounding area and stage for any oil drips.  Try and do all your low power (20X objective) dry lens work first and then switch over to high power oil objective lenses. When you are using the mercury arc bulb please ask others in the lab if they need it on before turning the bulb off. If you turn the bulb off pleas let it cool down ( at least 1/2 hour) before turning the bulb on again.

Slides:  When we are using many slides (such as a screen) pleas re-use the slides. After use soak the the slide and agar/agarose pad with hot water and soap. Let sit submerged in water for a couple of days and clean with soap and water and rinse with distilled water. You can dry the slides in the blue slide storage racks.

Pipettes

We have a variety of Pipettes in the lab. Usually each person will have their own set but some of you will have to share.  Please ask to use other people's pipettes.  The pipettes can last a very long time if they are used and cared for properly.  Always use a tip on a pipette and use the proper size tip. Do not force the tip on the Pipette barrel--just a slight push and tap.  Most of the pipettes have two position on the plunger the second position is to eject the last drop of fluid. Make sure when you draw up you liquid you: 1) Watch the tip go into the solution. 2) Make sure the thumb plunger is not pushed in all the way to the second position-just to the first position. If you push the plunger all the way down it will give you an inaccurate measurement . 3) Release the plunger slowly and watch the liquid move up into the tip.  Failure to watch the liquid move in the tip usually results in liquid going into the barrel. 4) Watch the liquid release from the pipette tip. Don't assume that if you press on the plunger the liquid goes in, especially for small volumes. Push out the last drops by depressing the plunger to the second position.

If you get liquid in the barrel clean it as soon as possible or bring the pipette to Ian.  If your suspect you calibration is off please bring the pipettes to Ian. To get a rough calibration you can pipette water onto the microbalance. 1.0 ml should equal 1.0 gram.

For Repair call PreAcc Inc. 416-736-0176  service@preacc.com 

Pleas see this site on Pipette Education

Pipette Education

Original Replacement Parts

Spare parts manufactured by Gilson are products of intensive research and stringent quality control, guaranteed to deliver value and assure reliability and precision over the long run. Conversely, counterfeiters cut corners on technical studies and quality control to slash costs. Appearances can be deceptive, these look-alike parts are often poorly finished, made to a poorer standard of low quality materials offering less durability as well as less thermal and chemical resistance. Their coarse design cannot be compared with the real thing. They find a privileged sales outlet with unscrupulous multi-brand service centers. Tip-holders, tip-ejectors, pistons, seals and O-rings are the preferred counterfeit parts.

 Stringent quality control, guaranteed to deliver value and assure reliability and precision over the long run.

  To view the complete “Counterfeit Parts and Pipettes” document from Gilson, please click here.

 Chemical Compatibility Chart (PDF)

 View the Chart

 Pipette Decontamination

 View the PDF

 The Proper Way to Load Pipette Tips

See the proper way to load your tips onto both your single channel and multi-channel pipettes.

Load Your Pipette Tip Properly

Symptoms of Improper Use and Maintenance Solutions

Pipetman® pipettes are easy to maintain. They can be easily dismantled and cleaned right in your own laboratory. Original Gilson replacement parts are always available from Mandel. Expert servicing is available from recalibration to a complete overhaul. Gilson pipettes will ensure the highest possible accuracy and precision, year after year.

Here are a few symptoms of improper use and solutions to keep maintenance costs down.

  1. Leaking at the Base of the Tip 
  2. Worn Piston 
  3. Jamming of the Operation Rod 
  4. Poor, inconsistent Results 
  5. Corrosion or Staining 

Caring for your Pipette

View or print the Gilson Pipetman® Two-minute Inspection

Problem

1. Leaking at the Base of the Tip

Solution

Use genuine Gilson tips

Not only do Gilson tips offer the finest quality but they are also the only tips designed specifically for Pipetman. A special sealing ring assures a perfect fit which is the only way to obtain results which systematically conform with Pipetman specifications.

Check the tip holder

Check the tip holder for scratches, wear, cracks or dirt at the point where the tip fits on. Dirty tip holders should be cleaned in a weak cleansing solution. Those with stubborn stains or scratches should be replaced. Check for blockages in the end of the tip holder. Removal of dirt or other foreign matter should be restricted to washing or lightly tapping on a bench. Never attempt to dig out blockages with a sharp tool.

Problem

2. Worn Piston 

Solution

Make sure the pipette is being used correctly

A worn piston can sometimes be temporarily corrected by replacing the seal and the 'O' ring. However, more often this indicates that the piston has become severely corroded due to improper use of the instrument and should be replaced. A piston should only be replaced by a qualified Gilson representative.

Problem

3. Jamming of the Operation Rod

Solution

Careful handling

When the push-button is depressed, the action should be smooth. If this is not the case, the operating rod maybe damaged or bent (this can occur when a pipette is dropped). The rod needs to be replaced and the pipette should be recalibrated. A rod should only be replaced by a qualified Gilson representative.

Problem

4. Poor, Inconsistent Results

Solution

Change the piston seal and the 'O' ring

Worn piston seals and 'O' rings can cause leaking and lead to inaccuracies. A good seal should move freely about the piston with a slight resistance to sliding. It should never drop freely under its own weight. Pipetman seals will withstand a minimum of 100,000 operations. But if they show any signs of wear or collected dirt particles, both the seal and 'O' ring should be changed.

Problem

5. Corrosion or Staining

Solution

Never leave pipettes on their side with liquid still in the tip

When not in use, pipettes should be placed vertically on a stable pipette stand such as the Gilson Carrousel™ or a Single™ wall-hanger. Otherwise liquid will run back into the tip holder and possibly damage the piston.

Operate with care

Like all precision instruments, Pipetman needs to be operated with care. If the push-button is allowed to spring back too quickly, the pistons will be splashed causing possible corrosion.


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