Cell & Molecular Biology Laboratory Molecular biology module (PDF
BIOB12H3S: Cell & Molecular Biology Laboratory Molecular biology module (PDF #5) Introduction and Terminology
In the molecular biology module, we wish to acquaint you with some of the most basic (but some of the most powerful) techniques in molecular biology, and help you to understand how clones are generated and how they can be characterized and used to examine gene expression. To begin to understand how powerful cloning tools can be, one must have some background in several areas. These concern recombinant DNA techniques, host cell transformation and selection of recombinant clones. We will discuss these aspects and you will perform some of them in the lab. Recombinant DNA technology is a battery of techniques, which continues to grow as clever scientists discover new methods. One of the most used techniques of molecular biology is the ability to propagate individual DNA molecules. We refer to this as ‘cloning’, for in essence, a clone of host cells is generated which contains only one type of recombinant DNA molecule. Here are some terms that are commonly used in molecular biology laboratories: 1. Restriction enzymes: Enzymes isolated from prokaryotes which recognize specific DNA sequences and act as endonucleases to break the phosphodiester backbone of DNA at specific sites. The type II restriction enzymes recognize palindromic sequences. Palindromes are regions of two fold symmetry and the nucleotide sequence is identical on both strands if read from the same direction. For example, the double stranded DNA below contains the palindrome AAGCTT. 5’ CCTGTCAGAATTGTAAGCTTCAATGGCGCATATCG3’ 3’ GGACAGTCTTAACATTCGAAGTTACCGCGTATAGC5’ 2. Library: A collection of clones representing either the entire genome of an organism or the genes that are expressed as mRNA (see below) • cDNA library: the target DNA is generated by making a DNA copy of mRNA from the organism (or tissue or developmental stage) of interest. • genomic library: the target DNA is generated (usually) by restriction enzyme digestion of genomic DNA. 3. Vector: A plasmid or a bacteriophage molecule that has at least three distinct properties: • an origin of DNA replication such that the vector can replicate autonomously. • single restriction enzyme sites for the introduction of the target DNA. • a selectable marker gene (usually one conferring antibiotic resistance).
1
4. Host: A bacterium (or sometimes a yeast strain), which is used to propagate individual DNA molecules. 5. Target or insert DNA: The foreign DNA fragment inserted into a vector. 6. Multiple cloning site: A region of a vector which contains several unique restriction enzyme recognition sites which may be used to clone target sequences. When incubated with a specific endonuclease, the enzyme will make a double stranded cut at its site in the MCS, resulting in a linear plasmid. The gene of interest is also digested using the same endonuclease. When the gene and linear vector are mixed together, the vector will reform a loop with the gene of interest in its MCQ. Note that the total size of the vector will thus increase. 7. Antibiotic-‐resistance genes: A sequence of bases in the plasmid that encodes a protein that makes the bacterial cell resistance to a select antibiotic. The antibiotic resistance gene is critical for maximizing the efficiency of DNA cloning. When bacteria are transformed with the modified plasmids, all bacteria will not take up the plasmid. The bacteria that fail to do so are of no use to us and should be killed to maximize the growth of the bacteria that did take up the vector. After transformation, the bacterial cells are exposed to this antibiotic. Only the cells that have taken up the vectors will survive, and these can then be cultured to clone our gene of interest. Some helpful reference material on recombinant DNA and cloning: Look in the Karp textbook (used for BIO B10/B11) at figures in chapter 18. (the formation of a recombinant plasmid produced by ligating a restriction fragment to a plasmid vector; the production of a library of human DNA fragments (genomic library); colony filter hybridization; a cDNA library). Molecular biology module labs: In this module, you will grow bacteria and then make them competent to take up foreign DNA (4A). You will transform them with an unknown plasmid DNA, and plate them on selective medium. Later, (4B) you will prepare plasmid DNA from these bacteria and use restriction enzymes to generate a restriction map of the clone (5A). By comparing the banding pattern to that of the known restriction map, you should be able to identify which unknown you were given (5B). Lab 4A: Preparation of competent cells, transformation with unknown plasmid DNA. Lab 4B: Preparation of plasmid DNA from cells. Lab 5A: Restriction enzyme digestion, gel electrophoresis analysis of fragments from digestion. Lab 5B: Analysis of data and restriction mapping. Plasmids used: (see Appendix 1 on page 22) pKNAT1 pOpaque 2 pLIM9a pKNAT3 pM1-‐9 2
Lab 4A: Preparation of competent host cells for transformation
E. coli is the host cell used in most molecular biology laboratories. A substantial amount of what we know about basic molecular processes is the result of research on E. coli. Plasmids and bacteriophages, which infect E. coli have been studied for over 60 years, and within the past 25 years a variety of different methods have been employed to make bacteria competent (capable of taking up DNA). One of the simplest techniques centres on the use of CaCl2. The calcium ions are thought to serve two purposes. First, they interact with the negatively charged phosphates in the phospholipid molecules of the E. coli cell membrane. At low temperature, this creates a relatively rigid membrane structure that has the propensity to form minute pores where the positive charges of the calcium shield the negative charges of the phospholipids. Since DNA also has a phosphate backbone, it interacts with the CaCl2 and forms a complex that precipitates onto the cells. This is the second role that calcium plays. After a short time, a heat shock is given which is thought to generate transient holes in the membrane and permit the plasmid DNA to ‘rush in’. Now the plasmid is inside the cell. If the plasmid confers some new and advantageous property to the bacterium, these cells may then be able to grow under conditions that would not ordinarily allow for growth. In the classical procedure, bacteria are grown to mid log phase, harvested, and resuspended in a small amount of sterile, ice-‐cold 50mM CaCl2. They are left on ice for about 20 minutes, harvested, and resuspended in fresh CaCl2. The cells are then left overnight in the refrigerator to ‘season’, which generally increases transformation efficiency by 5 to 10 fold. Such high efficiency competent cells are necessary when there is only a small amount of DNA available or one wants to obtain as many clones as possible (for example in constructing a library). Under optimal conditions such cells may give rise to over 107 transformants per microgram of supercoiled DNA. In most labs, libraries are carefully generated, and eventually useful clones are selected from the library. Consider a genomic library in bacteriophage lambda (λ). The genomic clone selected may contain as much as 23kbp of insert DNA, yet very little of this may be relevant to the ultimate goal (for example, one might want the promoter of a gene which could account for only about 1kbp of the total). Thus, it is of interest to take the long λ clone and make smaller subclones from it. For this purpose, there is generally a lot of DNA available, and its complexity is low (that is, there are many copies of a small number of different fragments present). In practice, this means that the transformation efficiency does not need to be high for typical subcloning purposes. There is a simple procedure that can be carried out in less than an hour beginning with a bacterial stock on a solid medium. In the lab, you will prepare competent E. coli by this method, and transform with one of the plasmids diagrammed in Appendix 1. You will then need to identify which plasmid your group was given after molecular analyses.
3
Transformation of E. coli with plasmid DNA
This experiment must be performed under sterile conditions. Use sterile tubes, pipette tips and solutions, and wear gloves when handling the tubes. Each pair will need: Other reagents & equipment XL-‐1 blue culture Inoculating loop sterile 10mM Tris, 50mM CaCl2 solution Bunsen burner 50ng/ul solution of plasmid unknown P1000, P20 micropipettors LB broth sterile pipette tips (200/1000ul) 2 LB plates ice bucket with crushed ice 2 LB + Ampicillin plates sterile 1.5 ml (Epi) tubes Epi tube rack beaker with 95% ethanol cell spreader Sharpie marker 37°C and 42°C water bath Procedure: 1. Wearing gloves, take 2 sterile Epi tubes from the stock beaker, close them and label them on the top: * “+” =experimental, contains plasmid DNA, or * “-“ =control, does not contain plasmid DNA 2. Using sterile technique and a sterile 1ml blue tip, take 1ml of bacteria from the XL-‐1 blue culture and place in each of the two sterile Epi tubes. Spin in a microfuge for 30 seconds to pellet the bacteria. Take off the supernatant with a Pasteur pipette. Turn tube upside down on a KimWipe and tap to remove all liquid. Immediately return tube to ice bucket. 3. Use another sterile pipette tip and add 250µl of the sterile, cold CaCl2 solution into the tube marked “+”. * Keep the tube cold. You might try leaving the tube in the ice (if it is packed ice and the tube won’t sink into it). * Gently mix the bacteria by pipetting with a P1000 to obtain a homogeneous suspension. 4. Repeat step 3 for the tube marked “-“. Now you have an experimental and a control sample ready. 5. Using a P20 and a sterile tip, take 10µl of the unknown plasmid DNA and introduce it directly into the cell suspension in the tube marked “+”. Mix gently by tapping lightly with your finger. 6. Leave the samples on ice for 15 minutes. 7. While the samples are incubating, use a Sharpie marker to write your name on each of the plates. For each of the LB plates, write “+” or “-‐“ (one of each) and for each of the LB + ampicillin plates, write “+” or “-‐“ (one of each). You will plate equal amounts of each sample on each of the two types of plates. (Total number of plates = 4). 8. Now for the heat shock. It is imperative that the cells are shocked. Take your ice bucket to the 42°C water bath, and plunge the tube into the water such that the cell suspension is covered. Do not shake or tap the tube, just hold it in place for
4
90 seconds (and no longer!). Return the tube to the ice bucket for one minute and do not shake or tap the tube. 9. Using a sterile tip, add 250µl of LB to each tube. Place in the 37°C water bath for 30 minutes. 10. Collect your samples, and spread 100µl of each onto the appropriate plates (LB or LB + ampicillin which are marked + or -‐). 11. Take your plates to the 37°C incubator and stack them upside down. Why is it important to do this? (record your answer in your notebook). 12. Clean up your lab bench and dispose of materials properly. **Proper disposal: All materials which have bacteria in/on them (pipette tips, paper towels, etc.) are to be placed in a AUTOCLAVE bag to be sterilized. The technician will take the plates out of the incubator tomorrow and start liquid cultures from an isolated colony. In the next lab period, your group will receive a liquid culture of bacteria harboring the unknown plasmid. For lab 4B of this module, you will harvest the bacteria and purify plasmid DNA from them.
Plasmid DNA conformations
The result of SSB and DSB on supercoiled DNA. This forms the basis of analyzing DNA fragments with SDS-‐ PAGE. Adapted from http://www.biochemsoctrans.org/bst/037/0893/bst0370893f01.htm
Plasmids are typically small molecules, usually not more than 10kbp in length. Replication in most E. coli strains generates molecules that are supercoiled. Supercoiling refers to the topology of the molecule, and is described by a solenoid or coiled coil. This superhelical tension gives the molecule a smaller surface area than a circular, non-‐supercoiled or linear molecule. Superhelical tension can be illustrated by taking a rubber band and, while holding one end immobile, twisting the other end around. The rubber band quickly adopts a supercoil, which makes it shorter. More twisting of the free end will produce additional supercoils and the rubber band will assume a very compact conformation. Plasmids are found in E. coli as supercoiled molecules, but during the purification process, nicking can occur. Nicking can occur by physical forces or can be due to the action of nucleases (endonuclease activity), with the result being the relaxation of superhelical density
5
(for one end of the double helix is now free to rotate) and the formation of open circular DNA.
Formation of an open circle due to a SSB
Supercoiled: Is very compact and rapidly moves through the gel matrix
-
Open circle: Has greatest surface area and is least flexible. Linear: can easily pass through gel matrix but may become tangled.
+
The 3 forms of DNA travel different distances during gel electrophoresis
Another form of DNA that you will encounter in this laboratory is linear DNA. A double stranded break will generate linear DNA from either supercoiled or open circular forms, and is often generated by restriction enzyme digestion (where the restriction enzyme cuts the target DNA only once). If there are two sites in the target molecule, linear DNA is generated, but it is in the form of two (linear) fragments. If these three forms are available you can see the effect of the conformation on the migration of the molecules in gel electrophoresis. Keep in 6
mind that these different forms are all the same size, but their conformation dictates how effectively they can move through a gel matrix. After you have purified your unknown plasmid DNA, you will subject it to restriction enzyme digestion and will run the products of digestion on a gel. Cleavage with a restriction enzyme (once or multiple times) will render all fragments linear and the migration of these fragments through the gel can be directly compared to linear size standards (usually purchased commercially, but can be made in the lab if you know the nucleotide sequence of an appropriate molecule).
Plasmid DNA purification
The cells are harvested and re-‐suspended in Solution 1, which contains a buffer (Tris) to stabilize the pH. This buffer also contains EDTA, which chelates cations. The effect of this is to deprive endogenous (or exogenously introduced) nucleases of the cations they require for activity and thus prevent the degradation of the plasmid DNA. Glucose is present to stabilize the osmotic environment of the cells and prevent premature lysis. Solution 2 is then added. It is a strongly basic solution (0.2 N NaOH) and so denatures macromolecules and disrupts hydrogen bonding between the DNA strands. It also contains SDS, a detergent, which also denatures proteins and solubilizes lipids. The effect of these two reagents is to lyse the cell and denature nucleic acids and proteins. Solution 3 is a concentrated potassium acetate salt solution at low pH. Upon addition, this has a two-‐fold effect. First, the low pH neutralizes the basic pH introduced by Solution 2. Second, the potassium salt forms a complex with SDS and the molecules to which it is bound. This results in a precipitate containing most of the membranes and chromosomal DNA of the cell. The plasmid DNA, in part due to the fact that it is supercoiled, can rapidly renature and it remains soluble.
Centrifugation removes most of the precipitate. Finally, the addition of ethanol dehydrates the plasmid DNA, and it can be collected as a precipitate, washed with 70% ethanol to remove excess salt, and resuspended in an aqueous buffer for further analysis. 7
Lab 4B: Purification of Plasmid DNA
In this lab you will purify plasmid DNA from bacteria. A few words of caution first: 1. Remember to dispose of bacteriological waste properly, by putting it in the BIOHAZARD bag. An EXCEPTION are glass Pasteur pipettes-‐these go in a special ‘sharps’ container that is separately autoclaved-‐your TA will advise you. 2. You haven’t yet used the microcentrifuge. Your TA will talk to you about it, but there are two rules: be sure the caps of your tubes are on securely, and be sure that the rotor is balanced (i.e. there is equal weight distribution. An Epi tube with the appropriate amount of water is a good balance tube).
Reagents Needed Solution 1: 50mM Tris pH8.0, 10mM EDTA, 50mM glucose Solution 2: 1% SDS, 0.2N NaOH Solution 3: 5M potassium acetate/acetic acid 70% and 95% ethanol 1X TE (10mM Tris pH7.5, 1mm EDTA) E. coli culture harboring unknown plasmid Supplies and Apparatus Needed 1.5ml microfuge (Epi) tubes Sharpie marker Micropipettors Epi tube rack Pasteur pipettes Disposable gloves Waste beaker Ice
Microfuge Ice bucket Vortex
Procedure: 1. You will work in 2 groups per bench. 2. Position a waste beaker, and place two paper towels on your bench surface. Using the Sharpie marker, label the tops of 3 Epi tubes with some label that identifies you. 3. Put on a pair of disposable gloves and wear them throughout the experiment. 4. Take out approximately 1ml of your bacterial culture with a pasteur pipette and dispense into the first labelled Epi tube. 5. Close the cap and place in a microfuge. Be sure that your partner has his/her tube in a position to balance the rotor. Spin for 30 seconds. 6. Take the tube out of the rotor, open the cap and pour the medium (the supernatant) into the waste beaker. Tap the tube on the paper towels several times to dislodge the last drops. 7. Place the tube in the ice bucket with the cap open. You should have a small brownish pellet. Note: it is essential that you keep the sample cold during the next several steps. There are times when you need to take it out of the ice bucket, but do your manipulations quickly and return the sample to the ice to chill. 8. Using a micropipettor set for 100µl, take out 100µl of Solution 1 and add it to the tube. Close the cap. Vortex the tube vigorously for 5 seconds and return to ice. Try to position the tube at a 45° angle to the vortexer cup to get the best mixing
8
action. This will likely not dislodge all the bacteria, but remember it is important to keep them cold. Hold it tightly near the top (so your hand doesn’t transfer heat to the sample). Repeat the vortex step until the suspension is uniform. 9. Now add 200µl of Solution 2. Mix by gentle inversion (DO NOT VORTEX!!!). You should see the solution clear. Return it to ice and wait 5 minutes. 10. Add 150µl of Solution 3, recap the tube. Now vortex vigorously for 10 seconds. You should see lots of white stringy material. Return to the ice for 5 minutes. 11. Working quickly, wipe the outside of the tube with a KimWipe™, and place it in the microfuge. Be sure the rotor is balanced. Spin for 3 minutes. During this time open the second of your Epi tubes and get your micropipettor (P1000) or a pasteur pipette ready to transfer the supernatant to the new tube. 12. The sample will have a large pellet along one of the walls of the tube and some of the precipitated material may be floating on the top of the sample. Avoid this precipitate and draw off the clear liquid into the second Epi tube. 13. Estimate the volume (it should be about 0.5ml). Using a P1000 micropipette, add approximately 2 volumes (1ml) of 95% ethanol (liquid should come up to the 1.5ml mark). Vortex 10 seconds. Let sit on the bench for 10 minutes. 14. Place the tube in the microfuge and balance it. Spin for 3 minutes. As in step 5, pour the supernatant into the waste beaker and tap the tube on the paper towel. 15. Using a new pasteur pipette, add approximately 1ml of 70% ethanol. Vortex for 10 seconds and spin for 3 minutes. Quickly decant the supernatant and tap tube on the paper towels. Let the DNA pellet dry. Note: this is important! 16. When the pellet is dry, resuspend it in 25µl of 1XTE. Label the tube side with your name and the date and “ML” (minilysate). Place it in the refrigerator in the box marked for your lab section. 17. Now there is a mess to clean up. The pasteur pipettes are to be placed (not thrown) into the decontamination solution. Carefully place any sharps and liquids in the appropriate waste bins. This leaves the rest of the solid waste (Epi, paper towels, etc). Be sure there are no sharps in it. Now take a wet paper towel and wipe down the bench area where you were working. Your gloves can be thrown in the garbage. 18. Be sure to wash your hands well with soap before you leave.
9
Lab 5A: Restriction Enzyme Digestion & Agarose Gel electrophoresis
In today’s lab, you will take set up restriction enzyme digests on aliquots of your plasmid DNA and separate the products of digestion by agarose gel electrophoresis. Restriction enzymes are found in prokaryotes and have greatly assisted molecular biologists by permitting the reproducible cleavage of DNA at defined sites. Hundreds of restriction enzymes can be purchased from commercial sources. The type II restriction enzymes recognize short, palindromic sequences and make two single stranded cuts at the target site. These may generate 5’ overhangs, 3’ overhangs or blunt ends. Examples of each of these are shown below. Eco RI (5’ overhang) Pst I (3’ overhang) Sca I (blunt ends) 5’-‐GAATTC-‐3’ 5’-‐CTGCAG-‐3’ 5’-‐AGTACT-‐3’ 3’-‐CTTAAG-‐5’ 3’-‐GACGTC-‐5’ 3’-‐TCATGA-‐5’ 5’-‐G 5’-‐CTGCA-‐3’ 5’-‐AGT 3’-‐CTTAAG-‐5’ 3’-‐G 3’-‐TCA (Note: 3’ halves are not shown) Restriction enzymes are commonly employed for DNA gel blots (Southern blots) to reproducibly fragment large chromosomal DNA molecules, and they are important in molecular cloning schemes to prepare both the vector and the target DNA molecules for ligation. Often after the subcloning of a fragment from a larger DNA molecule, restriction enzymes are used to determine if the proper fragment has been cloned. In some cases, large DNA fragments are subjected to restriction enzyme mapping to determine the linear order of restriction sites. This is useful in determining the orientation of the cloned fragment in the vector and the information can be used to decide on the subcloning strategy (i.e., which enzymes might be useful for generating small pieces for subcloning). In this laboratory exercise, you will subject aliquots of the DNA (you extracted in lab 4B) to restriction digests. You will also use electrophoresis to separate the DNA fragments by size, in order to get information to permit you to identify which plasmid you were given.
10
Electrophoresis
For most purposes, agarose gel electrophoresis is the method used to separate DNA fragments. Agarose is a neutral polysaccharide isolated from certain seaweeds, which when cooled forms a solid matrix that will carry an electrical current. DNA molecules have a phosphate backbone and thus carry a negative charge. During electrophoresis they move towards the positive electrode. When a gel is cast, the agarose is dissolved in the appropriate buffer [usually 0.5X TBE (Tris, boric acid, EDTA)] by heating (in a microwave oven), and after it cools to approximately 50-‐ 60°C, ethidium bromide is added and the gel is poured into a mold. A comb is inserted which creates ‘wells’. When the comb is removed, these wells are observed as small rectangular troughs. Agarose Gel DNA fragments with wells separated by size (-‐) (+) (-‐) (+) Buffer Chamber Small DNA fragments can easily move through the gel, but larger molecules encounter more resistance and therefore migrate more slowly. In most cases, one also runs a molecular size standard on the gel (we will use the 1kb ladder which is commercially purchased). The migration of these standard fragments through the gel can be measured and a graph can be constructed which plots the distance migrated versus the logarithm of the molecular weight. By measuring the distance migrated and correlating this to the plot of the standards, one can determine the antilog and hence the molecular size of the restriction fragments. In order to do this, you need to know one ‘constant’, which is that the molecular weight of 1basepair (bp) is approximately 660g/mole. Therefore a fragment of 4000bp: 4000 x 660 = 2.64 x 106g/mole. Of course when you plot the migration of the restriction fragments to get the logarithm value and take the antilogarithm, you will need to divide by 660 in order to get the length of the fragment. After you run your gel, you will visualize the fragments by taking your gel to the UV light box. Incorporated into the gel is a dye called ethidium bromide. Ethidium bromide is a small molecule that intercalates between the DNA bases and fluoresces orange upon excitation with ultraviolet (UV) light. This gives the DNA fragments an orange appearance against a black background.
11
There are two safety notes associated with this procedure: 1. Ethidium bromide is a mutagen and possibly a carcinogen. The TA will handle the chemical but you are required to wear gloves when handling the gel. The gel must be disposed of in the bag marked “ethidium bromide waste”. 2. The second caution relates to the use of the UV light source. The light box emits light in the UV range (260-‐360nm), and this causes damage to the retina of the eye. To avoid this, we will take three precautions. * First, you are required to have your lab partner or the TA with you when viewing the gel as a ‘buddy system’ to prevent exposure. * Second, before you turn on the light box, be sure the protective plastic cover is in place. This will block most of the harmful UV rays. * Lastly, you are required to wear a full face UV shield. This is to protect not only your eyes but also your entire face from UV irradiation.
Procedures: Notes on the restriction enzyme digest Restriction enzymes need the proper buffer in order to work. The buffer usually is 20mM Tris at pH 7.5-‐8.5, and has MgCl2 (divalent cations are required for activity). It may contain NaCl. Commerically supplied enzymes are shipped with a 10X buffer stock. You should also be aware that the plasmid miniprep technique results in a DNA preparation contaminated by E. coli RNA, so you need to include some RNAse in your digests. Note that some of the volumes you will use are very small and you need to be aware of what a microlitre looks like in the micropipettor tip. It is difficult to measure 0.5ul, but if you can see any liquid at all in the tip, that will be enough, as enzymes are very efficient. Logistics of this lab In order to be most efficient, arrange your experiment as follows: 1. Set up the restriction digests 2. Cast the gel (your TA will likely help you do this): make sure to take careful notes about how to do this in your notebook. 3. Wait 50 minutes and during this time set up the buffer tank and get the marker DNA ready. During this time you can get some practice in loading a gel with some marker dye. 4. Load your gel and perform electrophoresis 5. View gel, take photos (TA will assist) 6. Clean up your apparatus with DIW and clean the benches Each group will be required to build a restriction digest, using the enzymes EcoRI, Pst I, Hind III, and Xho I. Decide amongst yourselves which digest each person will do. In the interest of time and because we don’t have enough micropipettes for each person to have one, do the following. 12
1. Each person labels a reaction (Epi) tube. Build the reaction in the order given below (e.g. DNA is first). 2. Everyone adds their DNA, so no time is lost in adjusting the micropipette. 3. Then everyone adds the water, etc. A typical reaction contains: 5.0ul of DNA IMPORTANT!! 5.0ul of DIW Add the reagents in this order, using a 5.0ul of 4X buffer with RNAse different tip for each component. Be sure 5.0ul of diluted enzyme not to contaminate the 4X buffer, 20.0ul total volume or the restriction enzyme with your DNA. Place your tube in 37°C for 60 minutes. Then add 5ul of loading dye [contains glycerol to make the solution denser than water so that it will sink into the well; and tracking dyes to monitor progress of electrophoresis]. Centrifuge the tube and then prepare to load the gel.
Loading the gel
CAUTION: Wear gloves while handling the gel While your digest is proceeding, obtain a tube containing an aliquot of the 1kb ladder. When you are ready, fill the electrophoresis tank with 0.5X TBE. With gloves on, gently take the comb out of your gel with one smooth motion (after the gel has set up for 60 minutes), take the tape off of the ends (and place in the ethidium bromide waste) and place the tray in the buffer. Be sure there is enough buffer to cover the gel. (If there is too much buffer, pour some off before adding the gel tray). To load the gel, first note that your digest and the loading dye accounts for about 25µl, which is too much to load with a P20 micropipettor. There are two choices: (1) Load half the volume twice, or (2) use a P200. One other thing to be careful of is not to ‘stab’ the bottom of the well with the pipette tip. It is obviously important to know ‘which lane is which’, that is, to remember the order of the digests loaded on the gel. A simple way to do this is to load the marker lane first, then load the digests in alphabetical order. Each gel should also have one uncut plasmid lane so you can tell if your restriction digests actually worked. For example, 1 10-‐ lane gel could be loaded with 1 lane of marker, 1 lane of uncut DNA and 8 Restriction digests (4 from each group). Hint: In order to effectively load the gel, you need to have a steady hand. Most people cannot hold the micropipettor still enough to do this, but there is an easy way to ensure good loading. If you are right-‐handed, place your left hand on the gel tank and extend your index finger over the gel. Balance the barrel of the micropipettor on your extended finger to steady it and slowly introduce the liquid
13
into the well. While your samples are digesting, you can practice your loading technique on some gels the TA has cast, so when the time comes, you will be familiar with the loading technique.
Running the gel
CAUTION: Wear gloves while handling the gel
After the gel is loaded, attach the top with the electrical leads. Be sure that the DNA is traveling toward the positive pole (the electrodes are color coded; be sure the DNA runs toward the red pole and be sure that the appropriate electrical lead is plugged into the red panel on the power supply). Turn on the power supply and run at 100 volts. It should take about an hour for the bromophenol blue tracking dye to reach the end. Your TA will assist you in taking a photograph (this is your data for today). When you are finished with the gel, dispose of it in the ethidium bromide waste.
14
Lab 5B: How do I determine the identity of my unknown?
Your goal in this lab is to examine the restriction digests from Lab 5A and determine the identity of your unknown plasmid DNA. First, you need to look at the plasmid maps in Appendix 1 (page 22). They show that the inserts for all of the plasmids vary between 1.2 and 2.2kbp in length. Obviously they are different molecules and thus have different restriction maps. Take a look at the maps and see if you can cut with a single restriction enzyme, which will give you a diagnostic fragment. For example, suppose one of the plasmids has an Eco RI site about 800bp from the multiple cloning site, which also has an Eco RI site. If this plasmid is your unknown, digestion with Eco RI would give rise to a 0.8kpb fragment on a gel. Think about what patterns you would get if you were to digest with different restriction enzymes. What if you did a double digest (used two restriction enzymes in the same reaction)? In some cases a particular plasmid may not contain a particular restriction enzyme site. Thus it will remain uncut and migrate differently than linearized molecules or molecules which have been cleaved at more than one site (see page 5).
How can I determine the sizes of the fragments I see?
Scientists always run molecular size standards when performing restriction mapping. You can make your own by performing the appropriate restriction enzyme digest on a plasmid of known sequence (like a vector for example), or you can purchase size standards commercially. The marker we will use is called the ‘1kb-‐ plus’ ladder (see diagram below). Note that the intensity of the 1.6kbp band is double that of the others, and this provides a means for you to know “which band is which”. You will run 5µl of this marker on your gel and compare the sizes of the marker to those of your digests. Your TA will take a photograph of the gel and give you a photocopy. Plot the distance migrated verses the logarithm of the molecular weight (you can use semi-‐log paper to convert values to log) and determine the size of your fragments by reading off the graph.
15
Plasmid Maps and their interpretation
Plasmid maps can be drawn in several ways, with the two most common ones shown below. They can be sparse in information (for example, if little is known about the insert), or they can be very dense, with many restriction sites, the boundaries of genes (if any exist), and other information convenient to the investigator. Here are two ways that a plasmid can be drawn: 1. As a circle with insert in an arc, where it is difficult to measure distances.
16
2. With the insert displayed as a line above the plasmid, so that measurements are readily made. Using graph paper makes it easier to draw it to scale.
Restriction mapping strategy
Restriction mapping is generally used to determine whether sites for particular restriction enzymes exist and then to determine the linear order of sites. A few complications can arise. For example, if an enzyme cuts more than once, how do you determine where the sites occur? Regardless of whether an enzyme cuts once, twice, or multiple times, one thing that must be done is to determine where some sites exist (anchor points), and then try to determine where the unknown sites lie in relation to these anchor points. Often restriction sites comprising part of the multiple cloning site (MCS) can be used as anchor points. Example Question Consider the plasmid map above. A 5kbp Bam H1/ Hind III fragment (insert) is cloned into a 3kbp vector. If the plasmid is cut with Eco R1 and two fragments of 2.0kbp and 6.0kbp are generated as a result, where are the Eco R1 sites? Solution: Since there is only one Eco RI site in the vector (in the MCS) and the vector is 3kb in length, it is not possible for the 2.0kb fragment to be generated by the vector. Instead, the EcoRI site must be located in the insert, 2.0kb from the site in the vector. Thus, you have established that an EcoRI site is 2kb from one end.
17
Use of a second anchor point and the value of double digests
This information can now be used as a second anchor site to find where unknown sites lie in the target fragment. For example, you know that when you cut with EcoRI, you will get 2kb and 6kb fragments. What happens if you cut with Spe I? Suppose that you get an 8kbp linear fragment and that you know that there is no Spe I site in the vector. In order to place this site on the map, one of the things you could do is to perform a double digest with EcoRI and Spe I. Suppose that you get fragments of 0.5kbp, 2.0kbp, and 5.5kbp. Right away this tells you that the Spe I site lies near one of the ends of the 6kbp EcoRI fragment. But which end? The answer is: you know that there is no Spe I site in the vector and the vector comprises half of the 6kbp fragment. Therefore, the Spe I site must be located at the other end of the 6kbp fragment, 0.5kbp away from the EcoRI site, as shown below. EcoRI 6kb 2kb 5.5kb 0.5kb 2kb Spe I EcoRI A similar strategy can be used for other enzymes, but be wary! There may be more than one possible position for a site, and it may not be possible to determine the exact location based on the double digest. Several double digests with different combinations of restriction enzymes may be necessary to identify restriction sites.
Restriction Mapping Example
Suppose that you have cloned a 12kbp Eco RI/Bam HI fragment into a cloning vector. Other analyses have indicated that the gene you are interested in cloning lies within this fragment. Now you need to determine it’s restriction map so that you can subclone (make smaller clones) fragments and begin molecular analyses. You already know the entire sequence of the cloning vector, so you know where certain restriction sites exist in this sequence. Because you have cloned the 12kbp fragment in a particular direction (Eco RI to Bam HI), you also know the orientation of the insert with respect to the known sites in the vector molecule. Now you perform single and double digests, separate the fragments by electrophoresis, and calculate how large they are. From this information you can deduce the relative order of restriction sites within the insert. 18
Figure 1: The vector and insert
The data: Alu I: 2200, 4790, 8000 Xmn I: 4770, 10225 Pvu I: 5900, 9100 Sac I/Xmn I: 250, 500, 1500, 4000, 8725 Hind III: 2500, 12500 Alu I/Bam HI:1800, 2200, 3000, 8000 Sac I: 2000, 4000, 9000 Bam HI: 15000 Pvu I/Bam HI: 1110, 5900, 8000 The solution: I will give you an example of how to deduce where the Pvu I site is within the insert, and then I will give you the solution to the map on the following page. You should work through the data and convince yourself that you know how to do this exercise. Pvu I cuts the plasmid vector at a position 1612bp away from the site designated zero. The multiple cloning site is between 500-‐530, with the Eco
19
RI site at 515 and the Bam HI site at 525. Cutting with Pvu I gives two fragments (5900 and 9100bp). Thus, as this is a circular molecule, and since there is one site in the vector there must be one site in the insert. However, there are two possibilities. The site could be 5900bp away from the vector site in the clockwise direction, or it could be in the counterclockwise direction. It is impossible to determine from this information which possibility is correct. Thus, one must employ a double digest to decide. The Pvu I/Bam HI double digest is informative. This digest gives rise to fragments of 110, 5900, and 8000bp. Note that the 5900bp Pvu I fragment does not get cleaved by Bam HI, whereas the 9100 gets cleaved into 1100 and 8000bp fragments. Since you know that the only Bam HI site is the one used for cloning, and you can measure the distance from the vector Pvu I site to the Bam HI site in the multiple cloning site as 1087bp, this indicates that the Pvu I site in the insert is located 9100bp away from the vector Pvu I site in the counter-‐clockwise direction. In other words it is 8000bp away from the Bam HI cloning site. Take a look at the solution to the problem on the next page.
How can I get some practice at this?
1. There are several websites of other courses, which have tutorials. Try these: http://faculty.plattsburgh.edu/donald.slish/RestMap/RestMapTutorial.html http://faculty.plattsburgh.edu/donald.slish/Restmap.html 2. The best way to practice as well as appreciate the pitfalls of generating problems is to make up the problem yourself, beginning with the answer. On a graph sheet, draw an insert DNA and position some sites A, B, and C. If you want to increase the difficulty, put two A sites in. Connect this linear map to the backbone of a common vector (see page 23 and appendix 2). You might consider placing a site for A, B, and/or C in the vector, either at an end of the insert (as part of a multiple cloning site), or elsewhere. Now do your measuring and generate the data for each enzyme or enzyme pair. Now forget you know where the sites are and try to solve the problem just from the data. Often there are ambiguities and often you realize that the problem cannot be solved without giving more data (e.g. a double digest with an additional enzyme pair). By troubleshooting your own problem, you get some practice with “molecular logic” and appreciate how to approach a mapping problem.
20
Solution to problem on page 19:
21
22
23
In the molecular biology module, we wish to acquaint you with some of the most basic (but some of the most powerful) techniques in molecular biology, and help you to understand how clones are generated and how they can be characterized and used to examine gene expression. To begin to understand how powerful cloning tools can be, one must have some background in several areas. These concern recombinant DNA techniques, host cell transformation and selection of recombinant clones. We will discuss these aspects and you will perform some of them in the lab. Recombinant DNA technology is a battery of techniques, which continues to grow as clever scientists discover new methods. One of the most used techniques of molecular biology is the ability to propagate individual DNA molecules. We refer to this as ‘cloning’, for in essence, a clone of host cells is generated which contains only one type of recombinant DNA molecule. Here are some terms that are commonly used in molecular biology laboratories: 1. Restriction enzymes: Enzymes isolated from prokaryotes which recognize specific DNA sequences and act as endonucleases to break the phosphodiester backbone of DNA at specific sites. The type II restriction enzymes recognize palindromic sequences. Palindromes are regions of two fold symmetry and the nucleotide sequence is identical on both strands if read from the same direction. For example, the double stranded DNA below contains the palindrome AAGCTT. 5’ CCTGTCAGAATTGTAAGCTTCAATGGCGCATATCG3’ 3’ GGACAGTCTTAACATTCGAAGTTACCGCGTATAGC5’ 2. Library: A collection of clones representing either the entire genome of an organism or the genes that are expressed as mRNA (see below) • cDNA library: the target DNA is generated by making a DNA copy of mRNA from the organism (or tissue or developmental stage) of interest. • genomic library: the target DNA is generated (usually) by restriction enzyme digestion of genomic DNA. 3. Vector: A plasmid or a bacteriophage molecule that has at least three distinct properties: • an origin of DNA replication such that the vector can replicate autonomously. • single restriction enzyme sites for the introduction of the target DNA. • a selectable marker gene (usually one conferring antibiotic resistance).
1
4. Host: A bacterium (or sometimes a yeast strain), which is used to propagate individual DNA molecules. 5. Target or insert DNA: The foreign DNA fragment inserted into a vector. 6. Multiple cloning site: A region of a vector which contains several unique restriction enzyme recognition sites which may be used to clone target sequences. When incubated with a specific endonuclease, the enzyme will make a double stranded cut at its site in the MCS, resulting in a linear plasmid. The gene of interest is also digested using the same endonuclease. When the gene and linear vector are mixed together, the vector will reform a loop with the gene of interest in its MCQ. Note that the total size of the vector will thus increase. 7. Antibiotic-‐resistance genes: A sequence of bases in the plasmid that encodes a protein that makes the bacterial cell resistance to a select antibiotic. The antibiotic resistance gene is critical for maximizing the efficiency of DNA cloning. When bacteria are transformed with the modified plasmids, all bacteria will not take up the plasmid. The bacteria that fail to do so are of no use to us and should be killed to maximize the growth of the bacteria that did take up the vector. After transformation, the bacterial cells are exposed to this antibiotic. Only the cells that have taken up the vectors will survive, and these can then be cultured to clone our gene of interest. Some helpful reference material on recombinant DNA and cloning: Look in the Karp textbook (used for BIO B10/B11) at figures in chapter 18. (the formation of a recombinant plasmid produced by ligating a restriction fragment to a plasmid vector; the production of a library of human DNA fragments (genomic library); colony filter hybridization; a cDNA library). Molecular biology module labs: In this module, you will grow bacteria and then make them competent to take up foreign DNA (4A). You will transform them with an unknown plasmid DNA, and plate them on selective medium. Later, (4B) you will prepare plasmid DNA from these bacteria and use restriction enzymes to generate a restriction map of the clone (5A). By comparing the banding pattern to that of the known restriction map, you should be able to identify which unknown you were given (5B). Lab 4A: Preparation of competent cells, transformation with unknown plasmid DNA. Lab 4B: Preparation of plasmid DNA from cells. Lab 5A: Restriction enzyme digestion, gel electrophoresis analysis of fragments from digestion. Lab 5B: Analysis of data and restriction mapping. Plasmids used: (see Appendix 1 on page 22) pKNAT1 pOpaque 2 pLIM9a pKNAT3 pM1-‐9 2
Lab 4A: Preparation of competent host cells for transformation
E. coli is the host cell used in most molecular biology laboratories. A substantial amount of what we know about basic molecular processes is the result of research on E. coli. Plasmids and bacteriophages, which infect E. coli have been studied for over 60 years, and within the past 25 years a variety of different methods have been employed to make bacteria competent (capable of taking up DNA). One of the simplest techniques centres on the use of CaCl2. The calcium ions are thought to serve two purposes. First, they interact with the negatively charged phosphates in the phospholipid molecules of the E. coli cell membrane. At low temperature, this creates a relatively rigid membrane structure that has the propensity to form minute pores where the positive charges of the calcium shield the negative charges of the phospholipids. Since DNA also has a phosphate backbone, it interacts with the CaCl2 and forms a complex that precipitates onto the cells. This is the second role that calcium plays. After a short time, a heat shock is given which is thought to generate transient holes in the membrane and permit the plasmid DNA to ‘rush in’. Now the plasmid is inside the cell. If the plasmid confers some new and advantageous property to the bacterium, these cells may then be able to grow under conditions that would not ordinarily allow for growth. In the classical procedure, bacteria are grown to mid log phase, harvested, and resuspended in a small amount of sterile, ice-‐cold 50mM CaCl2. They are left on ice for about 20 minutes, harvested, and resuspended in fresh CaCl2. The cells are then left overnight in the refrigerator to ‘season’, which generally increases transformation efficiency by 5 to 10 fold. Such high efficiency competent cells are necessary when there is only a small amount of DNA available or one wants to obtain as many clones as possible (for example in constructing a library). Under optimal conditions such cells may give rise to over 107 transformants per microgram of supercoiled DNA. In most labs, libraries are carefully generated, and eventually useful clones are selected from the library. Consider a genomic library in bacteriophage lambda (λ). The genomic clone selected may contain as much as 23kbp of insert DNA, yet very little of this may be relevant to the ultimate goal (for example, one might want the promoter of a gene which could account for only about 1kbp of the total). Thus, it is of interest to take the long λ clone and make smaller subclones from it. For this purpose, there is generally a lot of DNA available, and its complexity is low (that is, there are many copies of a small number of different fragments present). In practice, this means that the transformation efficiency does not need to be high for typical subcloning purposes. There is a simple procedure that can be carried out in less than an hour beginning with a bacterial stock on a solid medium. In the lab, you will prepare competent E. coli by this method, and transform with one of the plasmids diagrammed in Appendix 1. You will then need to identify which plasmid your group was given after molecular analyses.
3
Transformation of E. coli with plasmid DNA
This experiment must be performed under sterile conditions. Use sterile tubes, pipette tips and solutions, and wear gloves when handling the tubes. Each pair will need: Other reagents & equipment XL-‐1 blue culture Inoculating loop sterile 10mM Tris, 50mM CaCl2 solution Bunsen burner 50ng/ul solution of plasmid unknown P1000, P20 micropipettors LB broth sterile pipette tips (200/1000ul) 2 LB plates ice bucket with crushed ice 2 LB + Ampicillin plates sterile 1.5 ml (Epi) tubes Epi tube rack beaker with 95% ethanol cell spreader Sharpie marker 37°C and 42°C water bath Procedure: 1. Wearing gloves, take 2 sterile Epi tubes from the stock beaker, close them and label them on the top: * “+” =experimental, contains plasmid DNA, or * “-“ =control, does not contain plasmid DNA 2. Using sterile technique and a sterile 1ml blue tip, take 1ml of bacteria from the XL-‐1 blue culture and place in each of the two sterile Epi tubes. Spin in a microfuge for 30 seconds to pellet the bacteria. Take off the supernatant with a Pasteur pipette. Turn tube upside down on a KimWipe and tap to remove all liquid. Immediately return tube to ice bucket. 3. Use another sterile pipette tip and add 250µl of the sterile, cold CaCl2 solution into the tube marked “+”. * Keep the tube cold. You might try leaving the tube in the ice (if it is packed ice and the tube won’t sink into it). * Gently mix the bacteria by pipetting with a P1000 to obtain a homogeneous suspension. 4. Repeat step 3 for the tube marked “-“. Now you have an experimental and a control sample ready. 5. Using a P20 and a sterile tip, take 10µl of the unknown plasmid DNA and introduce it directly into the cell suspension in the tube marked “+”. Mix gently by tapping lightly with your finger. 6. Leave the samples on ice for 15 minutes. 7. While the samples are incubating, use a Sharpie marker to write your name on each of the plates. For each of the LB plates, write “+” or “-‐“ (one of each) and for each of the LB + ampicillin plates, write “+” or “-‐“ (one of each). You will plate equal amounts of each sample on each of the two types of plates. (Total number of plates = 4). 8. Now for the heat shock. It is imperative that the cells are shocked. Take your ice bucket to the 42°C water bath, and plunge the tube into the water such that the cell suspension is covered. Do not shake or tap the tube, just hold it in place for
4
90 seconds (and no longer!). Return the tube to the ice bucket for one minute and do not shake or tap the tube. 9. Using a sterile tip, add 250µl of LB to each tube. Place in the 37°C water bath for 30 minutes. 10. Collect your samples, and spread 100µl of each onto the appropriate plates (LB or LB + ampicillin which are marked + or -‐). 11. Take your plates to the 37°C incubator and stack them upside down. Why is it important to do this? (record your answer in your notebook). 12. Clean up your lab bench and dispose of materials properly. **Proper disposal: All materials which have bacteria in/on them (pipette tips, paper towels, etc.) are to be placed in a AUTOCLAVE bag to be sterilized. The technician will take the plates out of the incubator tomorrow and start liquid cultures from an isolated colony. In the next lab period, your group will receive a liquid culture of bacteria harboring the unknown plasmid. For lab 4B of this module, you will harvest the bacteria and purify plasmid DNA from them.
Plasmid DNA conformations
The result of SSB and DSB on supercoiled DNA. This forms the basis of analyzing DNA fragments with SDS-‐ PAGE. Adapted from http://www.biochemsoctrans.org/bst/037/0893/bst0370893f01.htm
Plasmids are typically small molecules, usually not more than 10kbp in length. Replication in most E. coli strains generates molecules that are supercoiled. Supercoiling refers to the topology of the molecule, and is described by a solenoid or coiled coil. This superhelical tension gives the molecule a smaller surface area than a circular, non-‐supercoiled or linear molecule. Superhelical tension can be illustrated by taking a rubber band and, while holding one end immobile, twisting the other end around. The rubber band quickly adopts a supercoil, which makes it shorter. More twisting of the free end will produce additional supercoils and the rubber band will assume a very compact conformation. Plasmids are found in E. coli as supercoiled molecules, but during the purification process, nicking can occur. Nicking can occur by physical forces or can be due to the action of nucleases (endonuclease activity), with the result being the relaxation of superhelical density
5
(for one end of the double helix is now free to rotate) and the formation of open circular DNA.
Formation of an open circle due to a SSB
Supercoiled: Is very compact and rapidly moves through the gel matrix
-
Open circle: Has greatest surface area and is least flexible. Linear: can easily pass through gel matrix but may become tangled.
+
The 3 forms of DNA travel different distances during gel electrophoresis
Another form of DNA that you will encounter in this laboratory is linear DNA. A double stranded break will generate linear DNA from either supercoiled or open circular forms, and is often generated by restriction enzyme digestion (where the restriction enzyme cuts the target DNA only once). If there are two sites in the target molecule, linear DNA is generated, but it is in the form of two (linear) fragments. If these three forms are available you can see the effect of the conformation on the migration of the molecules in gel electrophoresis. Keep in 6
mind that these different forms are all the same size, but their conformation dictates how effectively they can move through a gel matrix. After you have purified your unknown plasmid DNA, you will subject it to restriction enzyme digestion and will run the products of digestion on a gel. Cleavage with a restriction enzyme (once or multiple times) will render all fragments linear and the migration of these fragments through the gel can be directly compared to linear size standards (usually purchased commercially, but can be made in the lab if you know the nucleotide sequence of an appropriate molecule).
Plasmid DNA purification
The cells are harvested and re-‐suspended in Solution 1, which contains a buffer (Tris) to stabilize the pH. This buffer also contains EDTA, which chelates cations. The effect of this is to deprive endogenous (or exogenously introduced) nucleases of the cations they require for activity and thus prevent the degradation of the plasmid DNA. Glucose is present to stabilize the osmotic environment of the cells and prevent premature lysis. Solution 2 is then added. It is a strongly basic solution (0.2 N NaOH) and so denatures macromolecules and disrupts hydrogen bonding between the DNA strands. It also contains SDS, a detergent, which also denatures proteins and solubilizes lipids. The effect of these two reagents is to lyse the cell and denature nucleic acids and proteins. Solution 3 is a concentrated potassium acetate salt solution at low pH. Upon addition, this has a two-‐fold effect. First, the low pH neutralizes the basic pH introduced by Solution 2. Second, the potassium salt forms a complex with SDS and the molecules to which it is bound. This results in a precipitate containing most of the membranes and chromosomal DNA of the cell. The plasmid DNA, in part due to the fact that it is supercoiled, can rapidly renature and it remains soluble.
Centrifugation removes most of the precipitate. Finally, the addition of ethanol dehydrates the plasmid DNA, and it can be collected as a precipitate, washed with 70% ethanol to remove excess salt, and resuspended in an aqueous buffer for further analysis. 7
Lab 4B: Purification of Plasmid DNA
In this lab you will purify plasmid DNA from bacteria. A few words of caution first: 1. Remember to dispose of bacteriological waste properly, by putting it in the BIOHAZARD bag. An EXCEPTION are glass Pasteur pipettes-‐these go in a special ‘sharps’ container that is separately autoclaved-‐your TA will advise you. 2. You haven’t yet used the microcentrifuge. Your TA will talk to you about it, but there are two rules: be sure the caps of your tubes are on securely, and be sure that the rotor is balanced (i.e. there is equal weight distribution. An Epi tube with the appropriate amount of water is a good balance tube).
Reagents Needed Solution 1: 50mM Tris pH8.0, 10mM EDTA, 50mM glucose Solution 2: 1% SDS, 0.2N NaOH Solution 3: 5M potassium acetate/acetic acid 70% and 95% ethanol 1X TE (10mM Tris pH7.5, 1mm EDTA) E. coli culture harboring unknown plasmid Supplies and Apparatus Needed 1.5ml microfuge (Epi) tubes Sharpie marker Micropipettors Epi tube rack Pasteur pipettes Disposable gloves Waste beaker Ice
Microfuge Ice bucket Vortex
Procedure: 1. You will work in 2 groups per bench. 2. Position a waste beaker, and place two paper towels on your bench surface. Using the Sharpie marker, label the tops of 3 Epi tubes with some label that identifies you. 3. Put on a pair of disposable gloves and wear them throughout the experiment. 4. Take out approximately 1ml of your bacterial culture with a pasteur pipette and dispense into the first labelled Epi tube. 5. Close the cap and place in a microfuge. Be sure that your partner has his/her tube in a position to balance the rotor. Spin for 30 seconds. 6. Take the tube out of the rotor, open the cap and pour the medium (the supernatant) into the waste beaker. Tap the tube on the paper towels several times to dislodge the last drops. 7. Place the tube in the ice bucket with the cap open. You should have a small brownish pellet. Note: it is essential that you keep the sample cold during the next several steps. There are times when you need to take it out of the ice bucket, but do your manipulations quickly and return the sample to the ice to chill. 8. Using a micropipettor set for 100µl, take out 100µl of Solution 1 and add it to the tube. Close the cap. Vortex the tube vigorously for 5 seconds and return to ice. Try to position the tube at a 45° angle to the vortexer cup to get the best mixing
8
action. This will likely not dislodge all the bacteria, but remember it is important to keep them cold. Hold it tightly near the top (so your hand doesn’t transfer heat to the sample). Repeat the vortex step until the suspension is uniform. 9. Now add 200µl of Solution 2. Mix by gentle inversion (DO NOT VORTEX!!!). You should see the solution clear. Return it to ice and wait 5 minutes. 10. Add 150µl of Solution 3, recap the tube. Now vortex vigorously for 10 seconds. You should see lots of white stringy material. Return to the ice for 5 minutes. 11. Working quickly, wipe the outside of the tube with a KimWipe™, and place it in the microfuge. Be sure the rotor is balanced. Spin for 3 minutes. During this time open the second of your Epi tubes and get your micropipettor (P1000) or a pasteur pipette ready to transfer the supernatant to the new tube. 12. The sample will have a large pellet along one of the walls of the tube and some of the precipitated material may be floating on the top of the sample. Avoid this precipitate and draw off the clear liquid into the second Epi tube. 13. Estimate the volume (it should be about 0.5ml). Using a P1000 micropipette, add approximately 2 volumes (1ml) of 95% ethanol (liquid should come up to the 1.5ml mark). Vortex 10 seconds. Let sit on the bench for 10 minutes. 14. Place the tube in the microfuge and balance it. Spin for 3 minutes. As in step 5, pour the supernatant into the waste beaker and tap the tube on the paper towel. 15. Using a new pasteur pipette, add approximately 1ml of 70% ethanol. Vortex for 10 seconds and spin for 3 minutes. Quickly decant the supernatant and tap tube on the paper towels. Let the DNA pellet dry. Note: this is important! 16. When the pellet is dry, resuspend it in 25µl of 1XTE. Label the tube side with your name and the date and “ML” (minilysate). Place it in the refrigerator in the box marked for your lab section. 17. Now there is a mess to clean up. The pasteur pipettes are to be placed (not thrown) into the decontamination solution. Carefully place any sharps and liquids in the appropriate waste bins. This leaves the rest of the solid waste (Epi, paper towels, etc). Be sure there are no sharps in it. Now take a wet paper towel and wipe down the bench area where you were working. Your gloves can be thrown in the garbage. 18. Be sure to wash your hands well with soap before you leave.
9
Lab 5A: Restriction Enzyme Digestion & Agarose Gel electrophoresis
In today’s lab, you will take set up restriction enzyme digests on aliquots of your plasmid DNA and separate the products of digestion by agarose gel electrophoresis. Restriction enzymes are found in prokaryotes and have greatly assisted molecular biologists by permitting the reproducible cleavage of DNA at defined sites. Hundreds of restriction enzymes can be purchased from commercial sources. The type II restriction enzymes recognize short, palindromic sequences and make two single stranded cuts at the target site. These may generate 5’ overhangs, 3’ overhangs or blunt ends. Examples of each of these are shown below. Eco RI (5’ overhang) Pst I (3’ overhang) Sca I (blunt ends) 5’-‐GAATTC-‐3’ 5’-‐CTGCAG-‐3’ 5’-‐AGTACT-‐3’ 3’-‐CTTAAG-‐5’ 3’-‐GACGTC-‐5’ 3’-‐TCATGA-‐5’ 5’-‐G 5’-‐CTGCA-‐3’ 5’-‐AGT 3’-‐CTTAAG-‐5’ 3’-‐G 3’-‐TCA (Note: 3’ halves are not shown) Restriction enzymes are commonly employed for DNA gel blots (Southern blots) to reproducibly fragment large chromosomal DNA molecules, and they are important in molecular cloning schemes to prepare both the vector and the target DNA molecules for ligation. Often after the subcloning of a fragment from a larger DNA molecule, restriction enzymes are used to determine if the proper fragment has been cloned. In some cases, large DNA fragments are subjected to restriction enzyme mapping to determine the linear order of restriction sites. This is useful in determining the orientation of the cloned fragment in the vector and the information can be used to decide on the subcloning strategy (i.e., which enzymes might be useful for generating small pieces for subcloning). In this laboratory exercise, you will subject aliquots of the DNA (you extracted in lab 4B) to restriction digests. You will also use electrophoresis to separate the DNA fragments by size, in order to get information to permit you to identify which plasmid you were given.
10
Electrophoresis
For most purposes, agarose gel electrophoresis is the method used to separate DNA fragments. Agarose is a neutral polysaccharide isolated from certain seaweeds, which when cooled forms a solid matrix that will carry an electrical current. DNA molecules have a phosphate backbone and thus carry a negative charge. During electrophoresis they move towards the positive electrode. When a gel is cast, the agarose is dissolved in the appropriate buffer [usually 0.5X TBE (Tris, boric acid, EDTA)] by heating (in a microwave oven), and after it cools to approximately 50-‐ 60°C, ethidium bromide is added and the gel is poured into a mold. A comb is inserted which creates ‘wells’. When the comb is removed, these wells are observed as small rectangular troughs. Agarose Gel DNA fragments with wells separated by size (-‐) (+) (-‐) (+) Buffer Chamber Small DNA fragments can easily move through the gel, but larger molecules encounter more resistance and therefore migrate more slowly. In most cases, one also runs a molecular size standard on the gel (we will use the 1kb ladder which is commercially purchased). The migration of these standard fragments through the gel can be measured and a graph can be constructed which plots the distance migrated versus the logarithm of the molecular weight. By measuring the distance migrated and correlating this to the plot of the standards, one can determine the antilog and hence the molecular size of the restriction fragments. In order to do this, you need to know one ‘constant’, which is that the molecular weight of 1basepair (bp) is approximately 660g/mole. Therefore a fragment of 4000bp: 4000 x 660 = 2.64 x 106g/mole. Of course when you plot the migration of the restriction fragments to get the logarithm value and take the antilogarithm, you will need to divide by 660 in order to get the length of the fragment. After you run your gel, you will visualize the fragments by taking your gel to the UV light box. Incorporated into the gel is a dye called ethidium bromide. Ethidium bromide is a small molecule that intercalates between the DNA bases and fluoresces orange upon excitation with ultraviolet (UV) light. This gives the DNA fragments an orange appearance against a black background.
11
There are two safety notes associated with this procedure: 1. Ethidium bromide is a mutagen and possibly a carcinogen. The TA will handle the chemical but you are required to wear gloves when handling the gel. The gel must be disposed of in the bag marked “ethidium bromide waste”. 2. The second caution relates to the use of the UV light source. The light box emits light in the UV range (260-‐360nm), and this causes damage to the retina of the eye. To avoid this, we will take three precautions. * First, you are required to have your lab partner or the TA with you when viewing the gel as a ‘buddy system’ to prevent exposure. * Second, before you turn on the light box, be sure the protective plastic cover is in place. This will block most of the harmful UV rays. * Lastly, you are required to wear a full face UV shield. This is to protect not only your eyes but also your entire face from UV irradiation.
Procedures: Notes on the restriction enzyme digest Restriction enzymes need the proper buffer in order to work. The buffer usually is 20mM Tris at pH 7.5-‐8.5, and has MgCl2 (divalent cations are required for activity). It may contain NaCl. Commerically supplied enzymes are shipped with a 10X buffer stock. You should also be aware that the plasmid miniprep technique results in a DNA preparation contaminated by E. coli RNA, so you need to include some RNAse in your digests. Note that some of the volumes you will use are very small and you need to be aware of what a microlitre looks like in the micropipettor tip. It is difficult to measure 0.5ul, but if you can see any liquid at all in the tip, that will be enough, as enzymes are very efficient. Logistics of this lab In order to be most efficient, arrange your experiment as follows: 1. Set up the restriction digests 2. Cast the gel (your TA will likely help you do this): make sure to take careful notes about how to do this in your notebook. 3. Wait 50 minutes and during this time set up the buffer tank and get the marker DNA ready. During this time you can get some practice in loading a gel with some marker dye. 4. Load your gel and perform electrophoresis 5. View gel, take photos (TA will assist) 6. Clean up your apparatus with DIW and clean the benches Each group will be required to build a restriction digest, using the enzymes EcoRI, Pst I, Hind III, and Xho I. Decide amongst yourselves which digest each person will do. In the interest of time and because we don’t have enough micropipettes for each person to have one, do the following. 12
1. Each person labels a reaction (Epi) tube. Build the reaction in the order given below (e.g. DNA is first). 2. Everyone adds their DNA, so no time is lost in adjusting the micropipette. 3. Then everyone adds the water, etc. A typical reaction contains: 5.0ul of DNA IMPORTANT!! 5.0ul of DIW Add the reagents in this order, using a 5.0ul of 4X buffer with RNAse different tip for each component. Be sure 5.0ul of diluted enzyme not to contaminate the 4X buffer, 20.0ul total volume or the restriction enzyme with your DNA. Place your tube in 37°C for 60 minutes. Then add 5ul of loading dye [contains glycerol to make the solution denser than water so that it will sink into the well; and tracking dyes to monitor progress of electrophoresis]. Centrifuge the tube and then prepare to load the gel.
Loading the gel
CAUTION: Wear gloves while handling the gel While your digest is proceeding, obtain a tube containing an aliquot of the 1kb ladder. When you are ready, fill the electrophoresis tank with 0.5X TBE. With gloves on, gently take the comb out of your gel with one smooth motion (after the gel has set up for 60 minutes), take the tape off of the ends (and place in the ethidium bromide waste) and place the tray in the buffer. Be sure there is enough buffer to cover the gel. (If there is too much buffer, pour some off before adding the gel tray). To load the gel, first note that your digest and the loading dye accounts for about 25µl, which is too much to load with a P20 micropipettor. There are two choices: (1) Load half the volume twice, or (2) use a P200. One other thing to be careful of is not to ‘stab’ the bottom of the well with the pipette tip. It is obviously important to know ‘which lane is which’, that is, to remember the order of the digests loaded on the gel. A simple way to do this is to load the marker lane first, then load the digests in alphabetical order. Each gel should also have one uncut plasmid lane so you can tell if your restriction digests actually worked. For example, 1 10-‐ lane gel could be loaded with 1 lane of marker, 1 lane of uncut DNA and 8 Restriction digests (4 from each group). Hint: In order to effectively load the gel, you need to have a steady hand. Most people cannot hold the micropipettor still enough to do this, but there is an easy way to ensure good loading. If you are right-‐handed, place your left hand on the gel tank and extend your index finger over the gel. Balance the barrel of the micropipettor on your extended finger to steady it and slowly introduce the liquid
13
into the well. While your samples are digesting, you can practice your loading technique on some gels the TA has cast, so when the time comes, you will be familiar with the loading technique.
Running the gel
CAUTION: Wear gloves while handling the gel
After the gel is loaded, attach the top with the electrical leads. Be sure that the DNA is traveling toward the positive pole (the electrodes are color coded; be sure the DNA runs toward the red pole and be sure that the appropriate electrical lead is plugged into the red panel on the power supply). Turn on the power supply and run at 100 volts. It should take about an hour for the bromophenol blue tracking dye to reach the end. Your TA will assist you in taking a photograph (this is your data for today). When you are finished with the gel, dispose of it in the ethidium bromide waste.
14
Lab 5B: How do I determine the identity of my unknown?
Your goal in this lab is to examine the restriction digests from Lab 5A and determine the identity of your unknown plasmid DNA. First, you need to look at the plasmid maps in Appendix 1 (page 22). They show that the inserts for all of the plasmids vary between 1.2 and 2.2kbp in length. Obviously they are different molecules and thus have different restriction maps. Take a look at the maps and see if you can cut with a single restriction enzyme, which will give you a diagnostic fragment. For example, suppose one of the plasmids has an Eco RI site about 800bp from the multiple cloning site, which also has an Eco RI site. If this plasmid is your unknown, digestion with Eco RI would give rise to a 0.8kpb fragment on a gel. Think about what patterns you would get if you were to digest with different restriction enzymes. What if you did a double digest (used two restriction enzymes in the same reaction)? In some cases a particular plasmid may not contain a particular restriction enzyme site. Thus it will remain uncut and migrate differently than linearized molecules or molecules which have been cleaved at more than one site (see page 5).
How can I determine the sizes of the fragments I see?
Scientists always run molecular size standards when performing restriction mapping. You can make your own by performing the appropriate restriction enzyme digest on a plasmid of known sequence (like a vector for example), or you can purchase size standards commercially. The marker we will use is called the ‘1kb-‐ plus’ ladder (see diagram below). Note that the intensity of the 1.6kbp band is double that of the others, and this provides a means for you to know “which band is which”. You will run 5µl of this marker on your gel and compare the sizes of the marker to those of your digests. Your TA will take a photograph of the gel and give you a photocopy. Plot the distance migrated verses the logarithm of the molecular weight (you can use semi-‐log paper to convert values to log) and determine the size of your fragments by reading off the graph.
15
Plasmid Maps and their interpretation
Plasmid maps can be drawn in several ways, with the two most common ones shown below. They can be sparse in information (for example, if little is known about the insert), or they can be very dense, with many restriction sites, the boundaries of genes (if any exist), and other information convenient to the investigator. Here are two ways that a plasmid can be drawn: 1. As a circle with insert in an arc, where it is difficult to measure distances.
16
2. With the insert displayed as a line above the plasmid, so that measurements are readily made. Using graph paper makes it easier to draw it to scale.
Restriction mapping strategy
Restriction mapping is generally used to determine whether sites for particular restriction enzymes exist and then to determine the linear order of sites. A few complications can arise. For example, if an enzyme cuts more than once, how do you determine where the sites occur? Regardless of whether an enzyme cuts once, twice, or multiple times, one thing that must be done is to determine where some sites exist (anchor points), and then try to determine where the unknown sites lie in relation to these anchor points. Often restriction sites comprising part of the multiple cloning site (MCS) can be used as anchor points. Example Question Consider the plasmid map above. A 5kbp Bam H1/ Hind III fragment (insert) is cloned into a 3kbp vector. If the plasmid is cut with Eco R1 and two fragments of 2.0kbp and 6.0kbp are generated as a result, where are the Eco R1 sites? Solution: Since there is only one Eco RI site in the vector (in the MCS) and the vector is 3kb in length, it is not possible for the 2.0kb fragment to be generated by the vector. Instead, the EcoRI site must be located in the insert, 2.0kb from the site in the vector. Thus, you have established that an EcoRI site is 2kb from one end.
17
Use of a second anchor point and the value of double digests
This information can now be used as a second anchor site to find where unknown sites lie in the target fragment. For example, you know that when you cut with EcoRI, you will get 2kb and 6kb fragments. What happens if you cut with Spe I? Suppose that you get an 8kbp linear fragment and that you know that there is no Spe I site in the vector. In order to place this site on the map, one of the things you could do is to perform a double digest with EcoRI and Spe I. Suppose that you get fragments of 0.5kbp, 2.0kbp, and 5.5kbp. Right away this tells you that the Spe I site lies near one of the ends of the 6kbp EcoRI fragment. But which end? The answer is: you know that there is no Spe I site in the vector and the vector comprises half of the 6kbp fragment. Therefore, the Spe I site must be located at the other end of the 6kbp fragment, 0.5kbp away from the EcoRI site, as shown below. EcoRI 6kb 2kb 5.5kb 0.5kb 2kb Spe I EcoRI A similar strategy can be used for other enzymes, but be wary! There may be more than one possible position for a site, and it may not be possible to determine the exact location based on the double digest. Several double digests with different combinations of restriction enzymes may be necessary to identify restriction sites.
Restriction Mapping Example
Suppose that you have cloned a 12kbp Eco RI/Bam HI fragment into a cloning vector. Other analyses have indicated that the gene you are interested in cloning lies within this fragment. Now you need to determine it’s restriction map so that you can subclone (make smaller clones) fragments and begin molecular analyses. You already know the entire sequence of the cloning vector, so you know where certain restriction sites exist in this sequence. Because you have cloned the 12kbp fragment in a particular direction (Eco RI to Bam HI), you also know the orientation of the insert with respect to the known sites in the vector molecule. Now you perform single and double digests, separate the fragments by electrophoresis, and calculate how large they are. From this information you can deduce the relative order of restriction sites within the insert. 18
Figure 1: The vector and insert
The data: Alu I: 2200, 4790, 8000 Xmn I: 4770, 10225 Pvu I: 5900, 9100 Sac I/Xmn I: 250, 500, 1500, 4000, 8725 Hind III: 2500, 12500 Alu I/Bam HI:1800, 2200, 3000, 8000 Sac I: 2000, 4000, 9000 Bam HI: 15000 Pvu I/Bam HI: 1110, 5900, 8000 The solution: I will give you an example of how to deduce where the Pvu I site is within the insert, and then I will give you the solution to the map on the following page. You should work through the data and convince yourself that you know how to do this exercise. Pvu I cuts the plasmid vector at a position 1612bp away from the site designated zero. The multiple cloning site is between 500-‐530, with the Eco
19
RI site at 515 and the Bam HI site at 525. Cutting with Pvu I gives two fragments (5900 and 9100bp). Thus, as this is a circular molecule, and since there is one site in the vector there must be one site in the insert. However, there are two possibilities. The site could be 5900bp away from the vector site in the clockwise direction, or it could be in the counterclockwise direction. It is impossible to determine from this information which possibility is correct. Thus, one must employ a double digest to decide. The Pvu I/Bam HI double digest is informative. This digest gives rise to fragments of 110, 5900, and 8000bp. Note that the 5900bp Pvu I fragment does not get cleaved by Bam HI, whereas the 9100 gets cleaved into 1100 and 8000bp fragments. Since you know that the only Bam HI site is the one used for cloning, and you can measure the distance from the vector Pvu I site to the Bam HI site in the multiple cloning site as 1087bp, this indicates that the Pvu I site in the insert is located 9100bp away from the vector Pvu I site in the counter-‐clockwise direction. In other words it is 8000bp away from the Bam HI cloning site. Take a look at the solution to the problem on the next page.
How can I get some practice at this?
1. There are several websites of other courses, which have tutorials. Try these: http://faculty.plattsburgh.edu/donald.slish/RestMap/RestMapTutorial.html http://faculty.plattsburgh.edu/donald.slish/Restmap.html 2. The best way to practice as well as appreciate the pitfalls of generating problems is to make up the problem yourself, beginning with the answer. On a graph sheet, draw an insert DNA and position some sites A, B, and C. If you want to increase the difficulty, put two A sites in. Connect this linear map to the backbone of a common vector (see page 23 and appendix 2). You might consider placing a site for A, B, and/or C in the vector, either at an end of the insert (as part of a multiple cloning site), or elsewhere. Now do your measuring and generate the data for each enzyme or enzyme pair. Now forget you know where the sites are and try to solve the problem just from the data. Often there are ambiguities and often you realize that the problem cannot be solved without giving more data (e.g. a double digest with an additional enzyme pair). By troubleshooting your own problem, you get some practice with “molecular logic” and appreciate how to approach a mapping problem.
20
Solution to problem on page 19:
21
22
23
