How to Pipette
Pipettes in the lab are available in sizes of 0.5-10 μL, 1-20 μL, 20-200 μL, and 200-1000 μL.
After selecting a pipette with the appropriate volume range for a given application, be sure to sterilize it by applying ethanol to a Kimwipe and rubbing the moist Kimwipe along the length of the pipette.
One can use the dial to set the desired volume to be taken up by the pipette. Note that the uppermost digit usually corresponds to the highest digit of the highest volume in the range. For example, the highest digit on a 20-200 μL pipette indicates hundreds of microliters; setting this digit to “2” using the dial should then indicate 200 μL.
Choose a box of tips that is compatible with the pipette you have selected. In general, the clear tips can be used by the smallest pipettes, the yellow tips can be used by the 20-200 μL pipette, and the blue tips can be used by the 200-1000 μL pipette.
There are two stops on each pipette:
- When collecting liquid, be sure to go down to the first stop only
- The level of the first stop depends upon the volume that has been set using the dial
- When dispensing liquid that has already been collected, push down all the way until the second stop
While collecting liquid, attempt to hold the pipette as vertically as possible
While dispending a liquid, it may help to do so by touching the pipette tip along the side of a tube to make use of surface tension. One should also be sure to dispense slowly unless otherwise directed by a specific protocol.
When a homogeneous mixture is preferred, one can also mix a solution by placing the pipette tip inside the solution and pipetting up and down.
After dispensing liquid, one should usually eject the tip of the pipette into a proper receptacle. Reuse of tips is only merited when the tip touches a single solution during its use.
According to sterile technique, should the pipette tip touch anything other than the liquid one is trying to manipulate, one should eject the tip and replace it with a new one.
In this particular lab, mixed 2 μL blue solution with 5 μL clear solution in an eppendorf tube. Set dial to 7 μL and, upon collecting the mixture, discovered that there was air at the tip end of the pipette tip,. This finding indicates either that the mixture was actually less than 7 μL or that the specific pipette used was improperly calibrated.
How to Make Solutions
Note: always use distilled water if water is required in the protocol
Basic strategy: Fill container halfway with water, add the predetermined amount of salt, and then fill to the necessary volume to achieve a particular concentration of salt solution.
- Potassium Chloride (KCl)
- Magnesium Chloride (MgCl2)
- Sodium Chloride (NaCl)
When weighing on the balance, be sure to close both doors on the side of the balance before pressing “tare”
Chemicals used during this lab:
- KCl: Molecular weight = 74.56 g/mol, Fisher Scientific, CAS 7447-40-7
- MgCl2: Molecular weight = 203.31 g/mol, Fisher Scientific, CAS 7791-18-6
- NaCl: Molecular weight = 74.56 g/mol, Fisher Scientific, CAS 7647-14-5
So, for the preparation of a 0.5 M solution of MgCl2 in 200 mL:
For MgCl2, the molecular weight is 203.31 g/mol, thus:
One should add 20.33 g of MgCl2 to 200 mL of water in order to create a solution with a concentration of 0.5M.
When the solution has been made, it should be labeled with the following information:
- identity of the solution, including concentration
- date of preparation
- name or initials of preparer
How to Make LB Agar Plates
To grow, bacteria need glucose, biotin, and salts, amongst other things
A protocol for making LB Agar plates does already exist as a handout. However, this is another version with unique annotations.
(In the iGEM lab) Weigh out 20 g Bactoagar and 20 g LB mix. Alternatively, one may use 15 g Bactoagar and 25 g LB. DO NOT USE Agarose instead of Bactoagar.
There is a large gray bin on a cart in the lab. The practice of the team has been to fill this bin about one quarter full of water so as to place the flask inside the water during autoclaving. However, some people feel that this is not necessary.
When going to the autoclave, be sure to bring autoclave tape and tin foil as well as a 2 L flask for mixing the ingredients. The 2 L flask ensures that there is extra volume to allow the liquid to boil.
(In the MDL office) Fill a large graduated cylinder with about 800 mL of distilled water. Add the LB to this water, making sure to mix it in slowly so as to allow it to dissolve. A spatula may be useful for adding the LB a more controlled fashion. Bring the final volume up to 1 L with additional distilled water. Transfer this solution into the 2 L flask, while also adding the Bactoagar. It might be helpful to alternate between pouring the liquid LB/water solution and adding the Bactoagar to allow for good mixing between the two. Swirl the flask until all of the solid has been dissolved.
When ready to place the final mixture into the autoclave, cover the top of the flask with tin foil and tape the bottom of the tin foil to the flask using autoclave tape. This tape should produce either black letters or black stripes to indicate successful autoclaving. Tape that has such black markings indicates that the container or substance is considered sterile.
Place in autoclave for about 20 minutes on the “sterile” setting using slow exhaust. Refer to the guide for autoclaving for more detailed instructions on using the autoclave.
Immediately after autoclaving is complete, avoid touching the flask for protracted periods of time. It goes without saying that this mixture should still be quite hot.
(In the iGEM lab) Drop a magnetic stir bar into the mixture and place the flask on a hotplate that can generate magnetic stirring. The tin foil covering should remain on the flask during this time. One should monitor the temperature using a thermometer. At around 50°C, the mixture should be ready for pouring into plates.
If you desire to make LB/AMP plates, you should add 100 mg of ampicillin (ampicillin trihydrate, which is stored in the door of the fridge) to the stirring mixture and allow it to dissolve. Ampicillin is heat-sensitive, so it makes sense to add the ampicillin close to the 50°C mark rather than when the mixture is hotter.
Clear a level space on the lab bench that will fit about 40 plates, unstacked. The plates should be labeled according to the contents (e.g. either “LB agar” or “LB agar + AMP”), the date of their preparation, and the name or initials of the preparer. Make sure to distinguish whether the plates have ampicillin or not. Labeling can be done while waiting for the mixture to cool on the hotplate, which can take some time.
When the agar mixture is ready, pour to a depth of about 3 mm into the bottom of each plate. The bottom of a plate is the smaller disk of the two composing the plate, while the lid is the larger one. At the very least, make sure that the entire surface of the bottom is uniformly covered in agar mixture. While pouring, lift each lid individually just high enough to allow pouring. This is generally done by tilting the lid while lifting so that one side is higher than the other. Do not allow anything to touch the inside of the lid.
Once the mixture has been poured, leave the plates sitting to allow it to congeal. This should take a roughly a couple of hours. After confirmation of agar solidification, one should stack up the plates and pull the bag in which they came over this stack, enclosing them in that formation. One should then tape the open end of the bag with regular tape. On this tape, one should again write the identity of the plates and the date of their preparation. This bag can then be placed in the fridge.