MMBB255 Week 9 Experiments 1 Tuesday's Procedures: A. Cloning: Isolation of plasmid DNA from the Escherichia coli transformants via the alkaline lysis miniprep procedure. Work in Pairs 1. Collect the bacterial cells that contain the plasmid DNA. Vortex your cultures and transfer 1 to 1.5 ml (just pour it) from each of your four overnight cultures of E. coli into four labeled eppi tubes. 2. Centrifuge (balance the tubes) at 12,000 rpm (revolutions per minute) for 5 min. Remove the tubes and gently pour off the supernatant back into the starting culture test tube. Save the cell pellet! Make sure to remove as much of the supernatant as possible (the medium can contaminate the DNA). 3. Add 100 l of Solution I to each of your cell pellets. Suspend the cell pellet in solution a by pipetting up and down or by vortexing. Make sure you have an even suspension. It is very important to get rid of all clumps in order to increase yield. 4. Add 100 l of Solution II. Work with one tube at a time. As soon as you have added solution II, cap the tube and invert back and forth 10 times to gently but evenly mix the contents. Ice for 5 minutes only. The solution should become more clear and viscous indicating that cells have lysed. Viscosity is caused by the abundance of nucleic acids. 5. Add 100 l of Solution III. Work with one tube at a time, vortex briefly to incorporate the solution into the viscous liquid. A white precipitate should form; this is the denatured protein and chromosomal DNA and will look like heated egg white. 6. Centrifuge for 10 minutes at max RPM. This will pellet the proteins/chromosomal DNA. 7. Pour or pipette the supernatant containing your DNA into a clean eppi tube. Be careful not to get any of the white precipitate. Leave some supernatant behind if need be. 8. Precipitate the DNA with alcohol. Isopropanol (0.7 volumes) or 95% ethanol (2 volumes) can be used to precipitate DNA. We will use ethanol. The approximate volume of liquid containing your DNA (the supernatant from step 7) is 300 l. To this add 600 l or 0.6 ml ethanol (2 x 300 l). Adjust volumes as needed. Mix by inverting several times gently and leave at room temperature for 5 min. The DNA will precipitate under these conditions. 9. Centrifuge. To collect the precipitated DNA, we will centrifuge the DNA. Place your tubes in the microfuge, making sure to balance the tubes in the rotor. Centrifuge at maximum rpm for 15 minutes. Open the cap and drain the liquid. Tap the edge of the tube on a paper towel to wick out the excess liquid. Look for a small white pellet. This is your plasmid DNA. 10. Carefully rinse the DNA pellet with1 ml ice cold 70% ethanol. Remove the ethanol carefully and air dry the tubes for 10 min. If the pellet comes up then centrifuge at max RPM for 10 min. 11. Suspend the DNA in TE. Add 25 l of Tris-EDTA buffer (TE, pH 8.0). Label your tubes and place them in a microtube rack. Your TA's will add 3 l of react 2 buffer and 1 l of HinDIII enzyme, incubate at 37 C for 1 hr., and store them at 4 C. You will examine your DNA via electrophoresis in a 0.7% <a href="/keyword/agarose-gel/" >agarose gel</a> during the next lab period. What is going on? Solution I: 50 mM Tris pH8, 10 mM EDTA pH8, and 100 g/ml RNase A. This is like the first solution in the chromosomal prep. A Buffer (Tris) plus the EDTA, which chelates divalent metals thereby destabilizing the membrane and inhibiting DNases. RNase to eliminate RNA molecules. Solution II: 0.2 M NaOH and 1% SDS. SDS (sodium dodecyl sulfate same as sodium lauryl sulfate) is a detergent that disrupts the membranes (thus lyses) and denatures proteins while the NaOH (very important difference from the chromosomal DNA isolation) denatures the large chromosomal DNA (makes it ss and linearizes it) and proteins (like the cell wall). The plasmid DNA is smaller and is more resistant to full denaturing in the limited exposure to the NaOH (it becomes ss but remains circular). Solution III: 3 M KAc, potassium acetate, (actually 3 M K and 5 M acetate since the pH is adjusted with glacial acetic acid) at about pH 5. Renatures circular DNA (ss to ds), precipitates ss linear DNA, proteins, and SDS (as KDS is not soluble in water). It also adds enough salt to allow the plasmid DNA to precipitate once ethanol is added (by masking the negatively charged DNA). MMBB255 Week 9 Experiments 2 Thursday's Procedures: A. Cloning: Analyzing the DNA via <a href="/keyword/agarose-gel/" >agarose gel</a> electrophoresis. Work in Pairs. Background: Agarose is a wells for positive highly purified form of agar and negative DNA (red lead) (black lead) gels made from this material can be used to separate DNA. The 0.7% <a href="/keyword/agarose-gel/" >agarose gel</a> purpose of this experiment is to demonstrate this electrophoretic TAE TAE buffer procedure using the plasmid buffer DNA isolated last Tuesday. We will use a 0.7% <a href="/keyword/agarose-gel/" >agarose gel</a> that side view of <a href="/keyword/agarose-gel/" >agarose gel</a> electrophoretic setup can resolve DNA in the range of 0.5 to 10 kbp. Using other wells for DNA agarose percentages will separate different DNA size to power supply ranges. Because DNA is a to power supply negatively charged molecule, it TAE TAE direction of migration buffer will migrate toward the positive buffer electrode, the red anode, in an electric field. For optimal top view of <a href="/keyword/agarose-gel/" >agarose gel</a> electrophoretic setup separation no more than a constant voltage of 5 V/cm (as measured from electrode to electrode) should be run. An aliquot of your DNA mixed with a loading buffer called &quot;blue juice&quot; will be placed in a well at one end of an <a href="/keyword/agarose-gel/" >agarose gel</a> that is surrounded by a tris-acetateEDTA (TAE) or a tris-borate-EDTA (TBE) running buffer (to allow current and to buffer the acid and base formations at each electrode). If you were to run the gel in just water you would not get any current and if you use concentrated running buffer then the current would be very high and a lot of heat would be generated possibly melting the gel and denaturing the DNA. The blue juice (loading buffer) that you will use contains 30% glycerol (to increase the density of the sample), 0.25% bromophenol blue (a blue tracking dye that migrates like a 300 bp DNA piece), and 0.25% xylene cyanol FF (another blue tracking dye that migrates like a 4 Kbp DNA piece). Once the gel is loaded with samples current is applied as shown in the diagram above. 1. Pouring the <a href="/keyword/agarose-gel/" >agarose gel</a> . Pre-poured 0.7% <a href="/keyword/agarose-gel/" >agarose gel</a> s will be prepared for you. Gel material is prepared by dissolving 0.7 g of agarose in 100 ml TAE or TBE buffer (0.7%); always use the same running buffer that will be used to run the gel in. This amount is usually enough for three thick mini-gels. Prior to pouring, the agarose is heated to boiling in the microwave for several minutes to completely dissolve the agarose. The liquid agarose is then cooled to about 55 C, ethidium bromide added (0.5 g/ml), and poured into the gel mold containing combs that form wells in the agarose into which you will later add your DNA. When the combs are added, it is important to make sure they do not touch the bottom of the mold so the samples do not leak out. Make sure the gel is completely set up (about 30 min. at room temperature) and then gently remove the combs. 2. Loading DNA into the <a href="/keyword/agarose-gel/" >agarose gel</a> . At the gel station, you will find the 0.7% <a href="/keyword/agarose-gel/" >agarose gel</a> in the gel box covered with 1 mm of running buffer, a P-20 pipettor, sterile tips for the pipettor, and a size standard (known DNA fragment sizes) - phage (lambda) DNA that has been cut with the restriction enzyme HindIII. 2 l blue juice Parafilm i. Prepare your DNA (see diagram) + 8 l of your strip- clean a. Place a piece of parafilm, clean side DNA sample side up, on your workbench. DNA: #1 #2 #3 #4 b. Set a P-20 pipettor to 2 l and place four (or the number of DNA samples that you have) 2 l spots of blue juice onto the parafilm. c. Remove 8 l from your first plasmid DNA miniprep and mix with one spot of blue juice. Repeat with each DNA sample, each into a different blue juice spot. Make sure you label the samples. ii. Load the gel a. Your instructor will demonstrate how to load samples into the wells. She/he will demonstrate by using the HindIII-digested DNA, which will serve as a size standard for your samples, and uncut plasmid. b. Load one DNA sample per well. Be careful not to insert your tip to far into the well as it may tear the well. Make sure you record which gel and lanes you placed your samples in. MMBB255 Week 9 Experiments 3 3. Running the <a href="/keyword/agarose-gel/" >agarose gel</a> at 5 V/cm for about 1.5 hr. Attach the electrodes to the gel box and the power supply. Connect the leads so that they are black to black and red to red connections. It may help to remember the phrase &quot;run to red&quot;. The DNA and tracking dye will move toward the red (positive-anode) pole and ethidium bromide toward the black (negative-cathode) pole. Adjust the power supply to 80 volts (16 cm distance between electrodes in this case); this may be increased, but watch out for heat melting the agarose and note that your resulting separation will not be pretty. The current should be about 25-30 mA per gel. If multiple gels are connected to the same power supply, this number will increase. We will run the gels at 120 V to decrease the time involved to about 45 minutes. If you see no bubbles forming at the electrodes then there is a bad connection somewhere (or the power supply is not turned on) and you should check that all connections have been made properly; ask for assistance. 4. Viewing the DNA. After 1.5 hr or until the leading dye front (bromophenol blue) has reached 3/4 of the gel, turn off the power supply and disconnect the leads (you can run the dye front to the end of the gel if you are not concerned about DNA less than 400 bps). The gel contains ethidium bromide (EtBr), a chemical that fluoresces when it intercalates into a double-stranded DNA (or RNA) molecule; it is also positively charged so the excess EtBr will migrate opposite of the DNA. The gel is still on the casting tray, so carefully pull the tray/agarose out of the running buffer with your gloved (Etbr is a carcinogen) hands onto a paper towel. Transport the tray/gel to the gel doc system down the hall for photographing the gel. There you will transfer the gel off the plastic tray onto a transilluminator (a device that shines ultraviolet light from underneath a sample placed on it) and take a picture. Your instructor will help you photograph the gel and store the resulting image in your notebook (there is a place for it). There are examples of documented DNA gel pictures below. The gels will look something like this: A B 1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 Gel A: Lane 1: (lambda) HindIII-digested DNA. ( DNA digested with Hi...