Lab 2C: Basic Techniques. Dilute 10X TE Buffer to Make 1X TE Buffer Determine the Concentration of an Unknown DNA Sample Streak out bacteria colonies

Transcription

1 Demonstration of sterile technique. Lab 2 Basic Techniques Lab 2A: Lab 2B: Lab 2C: Dilute 10X TE Buffer to Make 1X TE Buffer Determine the Concentration of an Unknown DNA Sample Streak out bacteria colonies Buffers Many of the experiments performed in molecular biology and biochemistry require proteins to carry out a particular function, such as binding to DNA or cleaving a substrate. The activity of these proteins is often very dependent on the ph, salt concentrations, and temperature of the reaction mixture. In some cases a change of ph from 7.5 to 6.5 or a 10-degree change in the temperature may cause greater than a 1000-fold reduction in the protein's activity. It is therefore very important to understand the function of the proteins involved in each experiment and know their optimal conditions for activity. As little as 15 years ago, most enzymes used to manipulate DNA were difficult to obtain because they were purified only by biochemists in research laboratories. In addition, there were a very limited number of enzymes available. However, as more and more of the techniques in molecular biology and biochemistry have become commonplace, companies have stepped in and now provide these enzymes. Many of the techniques that we use in molecular biology and biochemistry are now provided by these companies in the form of kits that include all enzymes, reagents, buffers, protocols, and (frequently) controls for the experiment. These kits are often very helpful, as well as convenient, for carrying out standard procedures. However, it is easy to get very complacent about just following the instructions and not understanding what is actually involved at each step of the protocol. You should understand enough about the procedure to know the function of the kit's buffers, regardless of whether you have to personally make it. A buffer is not a magic potion but a chemical solution containing a specific mixture of salts, buffering agents, and sometimes reducing agents, detergents or cofactors, etc., in which each of the components has a purpose and is included to optimize the reaction. 2-1

2 Solutions Most of the solutions used in molecular biology and biochemistry are calculated on the basis of the molarity of the solute. Sometimes a solution will be made on the basis of weight percent, parts per million (ppm), or normality. I. Molarity: A solution based on the number of moles of solute in a given volume of solution. Molarity = moles of solute liter of solution Example 1: If 2.92 g of NaCl is dissolved in enough water to make 250 ml of solution, what is molarity of the NaCl? (The molecular weight of NaCl is 58.5 g/mole.) M = moles solute 2.92 liter of solution g X 1 mole NaCl = moles NaCl 58.5 g NaCl = moles liters = 0.2 M (or 200 mm) Example 2: Calculate how much NaCl to use to make 300 ml of a 450 mm solution. A 450 mm solution is 0.45 moles/ liter and 300 ml = 0.3 liter. (Remember to keep the units constant in your calculations forgetting to convert ml to liters or mg to g could throw your calculations off 1000 fold) moles X 0.3 liter X 58.5 g NaCl = 7.9 g NaCl liter moles NaCl Therefore, to make 300 ml of a 450 mm NaCl solution add 7.9 g NaCl to water. Once the NaCl is dissolved, bring the final volume up to 300 ml by adding water. Why? II. Percentage: A percentile of the total volume. There are two kinds of percentage solutions: Percentage by weight (w/v), for dissolving solids, and percentage by volume (v/v), for diluting concentrated % solutions. % by Weight (w/v) = grams of solute 100 ml of solution Example: How do you prepare a 50-ml solution of 20% (w/v) Sodium Dodecyl Sulfate (SDS)? 20 g X 50 ml = 10 g SDS 100 ml 2-2

3 Therefore, to make 50 ml of a 20% (w/v) SDS solution, dissolve 10 g SDS in water and bring the volume up to 50 ml with water. % by Volume (v/v) = volume of concentrate X concentrate % final volume of solution Example: How do you prepare a 200 ml solution of 70% Ethanol from 100% Ethanol? 70% =? X 100% 70% X 200 ml =? 200 ml 100%? = 140 ml of 100% Ethanol Therefore, to prepare a 200 ml solution of 70% (v/v) Ethanol, start with 140 ml of 100% Ethanol and add water (60 ml) to bring the final volume up to 200 ml. The X Factor In a molecular biology research lab, you will constantly need to make and use buffers. In order to save time and space, molecular biologists often make concentrated stocks of solutions to last over long periods of time. Such concentrated stocks take up less space. In addition, these stocks are easily diluted for use when necessary. In this and other labs, you will often deal with solutions that are labeled 10X, 5X, 100X, etc. It is important to understand what this X factor means. The X factor simply indicates that the solution is in a concentrated form that must usually be diluted to a 1X concentration for use. For example, a 5X concentrated solution must be diluted 5-fold, while a 100X concentrated solution must be diluted 100-fold. The dilutions are usually done using water. Dealing with the X factor eliminates the need to know the actual molar concentration of the various components within the solution. You simply need to add water to make a 1X solution. However, a good scientist should always understand the composition of his or her reagents. Example: Prepare 1 liter of 1X TBE buffer from a 10X TBE stock solution. Below is a useful formula for doing dilution calculations: V 1 C 1 = V 2 C 2 V 1 = volume of stock buffer =? V 1 C 1 = V 2 C 2 C 1 = concentration of stock buffer = 10X? (10X) = (1 liter) (1X) 2-3

4 V 2 = volume of dilute buffer = 1 liter C 2 = concentration of dilute buffer = 1X? = (1 liter) (1X) / (10X)? = 0.1 liter Therefore, to prepare 1 liter of 1X TBE from 10X TBE stock, you should add 100 ml of 10X TBE to 900 ml of water. (Note: Diluting a buffer or solution does not change its gram or molar amount only it s concentration. The example above illustrates how a small volume of a concentrated buffer (100 ml of 10X TBE) is equivalent to a large volume of its diluted form (1 liter of 1X TBE). Preparing Buffers It is common in molecular biology to have to prepare complex buffers by diluting two or more stock solutions. For example, if you want to prepare TE buffer (0.1 mm EDTA, 10 mm Tris), it is best to simply dilute stocks of concentrated EDTA and Tris solutions. Example: Prepare 100 ml of TE buffer from 0.5 M EDTA and 1 M Tris stocks. Use the formula from above (V 1 C 1 = V 2 C 2 ) and solve for each of the two components independently: EDTA V 1 = volume of stock buffer =? C 1 = concentration of stock buffer = 0.5 M V 2 = volume of dilute buffer = 100 ml C 2 = concentration of dilute buffer = 0.1 mm (?) = (100 ml) (0.1 mm) / (500 mm) (?) = 0.02 ml Tris V 1 = volume of stock buffer =? C 1 = concentration of stock buffer = 1 M V 2 = volume of dilute buffer = 100 ml C 2 = concentration of dilute buffer = 10 mm (?) = (100 ml) (10 mm) / (1000 mm) (?) = 1 ml Therefore, to prepare 100 ml of TE Buffer from 0.5 M EDTA and 1 M Tris stocks, you should add 0.2 ml of 0.5 M EDTA and 1 ml of 1 M Tris to ml of water. We strongly suggest that you do some of the buffer and dilution problems in the previous quizzes. Similar problems will be on quizzes and exams in this class! 2-4

5 Lab 2A: Dilute 10X TE Buffer to Make 1X TE Buffer (Each person in each group should make his/her own 1X TE) 1. Make up 25 ml of sterile 1X TE. You will need to make 25 ml of 1X TE from a 10X TE stock and sterile water. Prepare this in a sterile 50-ml tube. SAVE THIS SOLUTION. It will be used to dilute DNA in experiment #3. To solve this dilution problem ask yourself What volume of 10X TE stock will I need in order to make 25 ml of 1X TE? (See the example at the bottom of p. 2-3.) V 1 =? C 1 = 10X TE V 2 = 25 ml C 2 = 1X TE Answer the following questions. A course staff will check your answers. 1. How much 10X TE do you need to use? 2. What pipetor will you need to dispense the liquid? 3. How much sterile water do you need to use? 4. What pipetor will you need to dispense the liquid? 2-5

6 Lab 2B: Determine the Concentration of an Unknown DNA Sample by comparison Method A: Set up titration of DNA plus ethidium bromide and use the camera system to estimate the concentration. Many experiments in molecular biology involve manipulating small quantities of DNA to digest, clone, or sequence. In working with small quantities of DNA it is often important to know the concentration of the DNA samples in order to determine whether the experiment is practical and feasible. Although in most cases the exact concentration is not critical, it is often important to know the relative concentration of the DNA; i.e. whether you are working with 0.1, 1, 10 or even 100 micrograms (µg) of DNA. The following lab experiments are methods used to determine the concentration of DNA in a sample. This exercise is not only useful for learning how to measure the concentration of DNA, but it will also familiarize you with working with pipetmen and the camera set-up, which will be used often in the course for a variety of procedures. Each person in each group will be given a different sample of plasmid DNA of unknown concentration (Unknown DNA X or Unknown DNA Y) and a sample of plasmid DNA with a known concentration (the DNA Standard). Each person in each group will determine the concentration of their unknown sample by first serially diluting his/her DNA sample. Following the addition of ethidium bromide, the DNA dilutions will be visualized using a UV light source. The concentration of the Unknown DNA will also be measured by determining the amount of UV light that it absorbs. Ethidium bromide (EtBr) is a dye that intercalates between the DNA bases and emits fluorescence. The amount of fluorescence emitted is directly proportional to the amount of DNA in the sample (over a given range of DNA amount). The concentration of a sample of DNA can therefore be determined by comparing the fluorescence of sample with that of a DNA sample of known concentration. As little as 1-5 nanograms (ng) of DNA can be detected by this method. The presence of any protein contaminants in a presumably pure DNA solution does not interfere with the assay but there may be other contaminants in your DNA that may quench or possibly contribute to the fluorescence. Caution! Ethidium bromide, which is a potent mutagen, can intercalate into your DNA as well as your sample DNA. Gloves should be worn when working with solutions of EtBr. Prolonged exposure to UV light is also dangerous to your skin, and especially your eyes. To minimize exposure make sure the UV light box is shielded and that you wear protective eye goggles to efficiently block UV radiation. 2-6

7 Determine the concentration of DNA using ethidium bromide Using ethidium bromide to measure the concentration of your DNA is a good method when your DNA concentration is very low (<250 ng/µl) or if your sample is contaminated with proteins or phenol. In this experiment, each person will make a series of dilutions of a DNA sample of known concentration (the DNA standard). Another series of dilutions will be made using DNA samples of unknown concentration (one group partner gets Unknown DNA X and the other partner gets Unknown DNA Y). Ethidium bromide will then be added to all DNA dilutions, then the dilutions will be visualized under UV light. By comparing the fluorescence of the unknown DNA samples with that of the known DNA samples (the standards), it should be possible to estimate the concentration of DNA in the unknown sample. 1. Serially dilute your DNA sample (Unknown DNA X or Y) 5 times, by 5-fold dilutions. a) Label 5 microcentrifuge tubes with 1X, 2X, 3X, 4X, and 5X if you have the X sample or 1Y, 2Y, 3Y, 4Y and 5Y if you have the Y sample. b) Using a P-200, aliquot 40 µl of TE into each of 5 labeled microfuge tubes. Make sure that you have pipeted the correct volume into each tube by comparing it with standard tube with the blue dye. c) Use a P-20 to add 10 µl of the Unknown DNA (X or Y) to Tube #1 and vortex. (see the figure on the next page). You have now diluted the DNA 5 fold. d) Add 10 µl of the 1X (or Y) to Tube #2 and vortex. You have now diluted the DNA 25 fold. e) Add 10 µl of the 2X (or Y) to Tube #3 and vortex. You have now diluted the DNA 125 fold. f) Add 10 µl of the 3X (or Y) to Tube #4 and vortex. You have now diluted the DNA 625 fold. g) Add 10 µl of the 4X (or Y) to Tube #5 and vortex. You have now diluted the DNA 3125 fold. 2. Serially dilute your DNA Standard solution 5 times, by 5-fold dilutions. a) Label 5 microcentrifuge tubes with 1S, 2S, 3S, 4S, and 5S. b) Using a P-200, aliquot 40 µl of TE into each of 5 labeled microfuge tubes. c) Use a P-20 to add 10 µl of the DNA Standard ) to Tube #1S and vortex. (The concentration of the DNA Standard is 1 µg/µl). (See the figure on the next page). d) Add 10 µl of the 1S to Tube #2S and vortex. e) Add 10 µl of the 2S to Tube #3S and vortex. f) Add 10 µl of the 3S to Tube #4S and vortex. g) Add 10 µl of the 4S to Tube #5S and vortex. 3. Pipet 40 µl of TE buffer into another tube to serve as a blank control. You should now have 11 tubes in total 5 dilutions the Unknown DNA, 5 dilutions of the DNA Standard, and 1 with TE only. 2-7

8 Serial Dilutions for the EtBr Experiment Label 5 tubes for diluting the unknown DNA sample (X or Y). Add 40 µl of 1X TE to each tube Add 10 µl of the unknown DNA stock to the first tube. Transfer 10 µl of the sample in tube 1 to tube 2. 1 Unknown DNA 1 2 Cap tube 2 and vortex to mix. (This DNA Cap tube 1 and vortex to mix. (This DNA has now been diluted 5-fold from the has now been diluted 25-fold from the starting stock. ) starting stock. ) Transfer 10 µl of the sample in tube 3 to tube 4. Transfer 10 µl of the sample in tube 2 to tube Cap tube 4 and vortex to mix. 3 Cap tube 3 and vortex to mix. (This DNA has now been diluted 125-fold from the starting stock. ) Transfer 10 µl of the sample in tube 4 to tube 5. 4 Repeat this process for the known DNA standard. 5 Cap tube 5 and vortex to mix. 2-8

9 4. Prepare a 2 µg/ml solution of EtBr and spot 3 µl-drops of a diluted solution of EtBr onto a petri dish as shown below. a) Label two microfuge tubes ET-200 and ET-2 b) Add 98 ul of TE to the ET-200 tube and 198 µl TE to the ET-2 tube. c) Add 2 ul of 10 mg/ml EtBr stock to the ET-200 tube and vortex. This is a 1-50 dilution. d) Add 2 ul of ET-200 EtBr stock to the ET-2 tube and vortex. This is a dilution. (You have now serially diluted your 10 mg/ml EtBr stock to a final concentration of 2 µg/ml.) e) Place half of a clean 60-mm dish over the circular outline below. It will serve as a guide on where to place your drops. f) Add 3-µl drops of your 2 µg/ml EtBr solution such that there are two rows of 5 drops and one drop off to the side. (See the figure below.) Spots for DNA standards TE control Spots for unknown DNAs To each of the above spots of 2 µg/ml EtBr place an equal volume (3 µl) of one DNA sample dilution from the numbered tubes as shown and mix by pipeting up and down several times. To the remaining spot add 3 µl of TE as a negative control. This last spot is a zero DNA control and will indicate the background fluorescence of the assay. 6. Being careful not to disturb the spots, take the plate to the dark room and photograph the spots when exposed to UV light. It is best to photograph your plate by placing it upside down on the UV light box. The instructors or TAs will demonstrate how to use the camera set-up. You will use the camera set-up throughout the course, so take notes on how to use it!! 7. Estimate the concentration of your unknown sample by comparing the intensity of the spots with the spots of the DNA standards. Later you will compare and comment on the calculated values of each method. Tape your picture in your notebook neatly use transparent tape. And label your picture with a pen or thin marker. 2-9

10 Lab 2C: Streak out bacteria colonies In last week s lab you plated the bacteria library on selective media (LB-Kan) to select for cells that were transformed with a plasmid from the library. We therefore need to perform a second round of the screen to distinguish real mutants among the wild type cells. The best way to do this is to streak out the colonies for single cells and examine the differences in growth of these individual colonies. A streak out is exactly like it sounds. 1. Take a sterile toothpick and gently touch one of the colonies on the plate. A bacterial colony typically has around 10 6 cells. Touching the toothpick to the colony transfers a large number of the cells onto the toothpick. 2. Gently scrape the toothpick on a fresh plate. Normally this is just a short line (Streak 1 in Fig L2C-1). A large number of the cells will transfer from the toothpick to the plate. Often so many cells will transfer that a patch of cells will appear after the plate is incubated for a few days. This patch may contain a number of cells from different parents. Fig L2C.1 A streak out of the blue cell from the plate shows that there was a mixture of cells on the toothpick. 1, 2, and 3 indicate the order of the streaks. The streak out plate is shown after the cells have had a chance to grow ON. 3. Take a new sterile toothpick and drag it in perpendicular direction across the end of the first streak and then extend it a short distance (Streak 2 in Fig L2B-1). The toothpick will pick up some cells from the first streak and then spread them on the media as the toothpick is dragged across the plate. There may still be such a large number of cells that it forms a patch with a mixture of cells. 4. Take a new sterile toothpick and drag it in perpendicular direction across the end of second streak and then extend the streak in a zig-zag fashion across a section of the plate (Streak 3 in Fig L2B-1). The toothpick will pick up a few cells from the second streak and then spread them on the media as the toothpick is dragged down the plate. There will now be so few cells on the toothpick that the individual cells will deposited on the plate a disperse manner. When these cells grow they will form individual colonies instead of a patch of cells that came from different parents. All the cells in a colony will be generated from one cell and so they will be clonal. The example shows that the streak out contained a mixture of blue and white cells. However, 2-10 Fig L2C.2 Examples of good and bad streak outs. The streak outs on the left and in the middle are good because there are individual colonies. The one on the right is bad because it is a large patch with no individual colonies. This patch may be a mixture of colonies. Some of the colonies in the left and middle are smeared because they were picked to make overnight cultures.

11 individual colonies at the end of the streak out are clonally pure. Examples of good and bad streak outs are shown in Fig L2C.2. We are going to use the streak outs not only to make sure that the cells are clonally pure but to also rescreen the growth on Kan. Protocol for streak outs of colonies from the library plating. 1. Each student will be given 1 LB-Kan (two black lines on side) plate. a) Using a ruler and a fine point Sharpie draw two perpendicular lines on the back of each plate to divide it up into quarters (see Fig L2C.1). b) Label each quadrant with (a) Your Class day (e.g. T- Tuesday lab, W- Wednesday lab, H- Thursday lab) (b) Your Group Number (e.g. 1, 2, 3 etc) (c) Your initials (e.g. AV, JM, etc) (d) The numbers of the clone (e.g. 1, 2, 3 or 4) (e) The year (e.g..10) For example if you are a member of group 2 in the Thursday and your initials are AB, then you should label the four quadrants: H2AB1.10, H2AB2.10, H2AB3.10 and H2AB4.10. Do not write on the lids! Lids can be switched, causing strains to be mixed up. 2. Select a colony from the LB-Kan library transformation plate and streak it out onto the first quadrant on the plate. Remember that the plate is flipped so the quadrant will be the opposite side!!! a) As described above, use one toothpick to pick the colony from the transformation plate and make a short streak on one quadrant on the plate. c) Use a fresh toothpick to make a short diagonal streak across the previous streak on the plate. d) Use another fresh toothpick to make one streak across the second streak and then zig-zag the streak several times. 3. Repeat the streak out process picking 3 different colonies from the library plate that you made last week. You should mark the back of the library plate which colonies that you picked. 4. These plates will be incubated at 37 C for several days until the streak outs are well developed and single colonies are visible. 2-11