dna lab: restriction mapping - biolympiads · cell and molecular biology autumn 2005 4 lab #3 dna...

Cell and Molecular Biology Autumn 2005

1

Lab #3

DNA Mapping Lab

DNA Lab: Restriction mapping Readings in Alberts 2nd Ed (Ch 10: 328-330; 341-345); Becker 6th Ed (Ch18: 524-527; Ch 20: 633-640); 5th Ed (Ch16: 492-496; Ch 18: 608-610). Useful Web sites

Primer on mapping DNA: http://www.ncbi.nlm.nih.gov/About/primer/mapping.html Tutorial on restriction mapping:

http://www.carolina.com/biotech/plasmid_problems/plasmid_guide.asp

Within the 3 billion or so nucleotides that make up the human genome, identifying one particular

gene that is composed of only several thousand nucleotides is a challenging task. This task is simplified somewhat by creating maps of the DNA. Just like a road map, a DNA map uses DNA dependent landmarks to help determine where a gene resides in the genome. DNA can be mapped to varying degrees of resolution. Genetic maps are usually lower resolution maps, made by measuring the recombination frequency between different heritable traits. Genetic maps are used to associate a region of a chromosome with a particular trait; they also can be used to determine gene order along a chromosome. Physical (distance) maps are higher resolution maps based on DNA sequence, and are measured in numbers of base pairs (often in kilobases or megabases). The ultimate physical map of a piece of DNA is its complete sequence. However, if DNA sequence information is not available, a lower resolution map can still be generated using enzymes that cut DNA at specific sequences. These enzymes are called restriction enzymes. Restriction mapping is used to determine the distance between the sites where restriction enzymes cut and to order the different sites relative to each other. The cut sites serve as landmarks when keeping track of several pieces of DNA. Genetic and physical maps can be correlated to identify the DNA sequences encoding a particular trait. Beyond mapping, restriction enzymes also are useful in recombinant DNA technology. Two different DNA pieces cut by the same restriction enzyme can be readily joined back together in a test tube. Moreover, because the genetic code is universal, it is possible to join DNA from one organism to the DNA of another organism and produce a functional gene. Recall from the Fluorescence Microscopy lab where GFP was targeted to the mitochondria of tobacco cells. DNAs from four different organisms were incorporated into the tobacco cells. A short segment of the yeast coxIV gene was joined to the jellyfish GFP gene in a test tube; the fusion gene was then cloned into a transformation vector that contained sequences from bacteria and from a plant virus.

In this lab, you will be mapping some chloroplast genes isolated from the single-celled alga, Chlamydomonas. The chloroplast genome was broken up into smaller pieces called clones; your goal is to put together a restriction map for one of these clones. Based on the restriction enzyme pattern for each clone, it is possible to reconstruct the entire map of the chloroplast DNA. Please read through the background material so you will understand some common terms in molecular biology. Terms to understand:

Agarose gel electrophoresis Ethidium Bromide Multiple cloning site (MCS) Physical map Plasmid Restriction enzyme Restriction map Vector


2

Lab #3

DNA Mapping Lab

Background Restriction Endonucleases Restriction endonucleases are enzymes that cut DNA at specific sequences. For example, the enzyme EcoRI cuts DNA at the sequence GAATTC. The enzymes hydrolyze the phosphate backbone creating a nick in the DNA strand. Bacteria produce restriction enzymes as a defense against invading viral DNA. In E. coli, foreign DNA with the sequence GAATTC would be cut and inactivated. To protect its own DNA from the restriction enzyme, bacteria also produce DNA modifying enzymes. For each restriction endonuclease there is a corresponding modifying enzyme that blocks restriction activity on the host’s own DNA, generally by methylating the DNA at the recognition sequence. The protruding methyl group presumably prevents catalysis by interfering with the close molecular interaction between the restriction enzyme and its recognition site. For example, EcoRI methylase adds a methyl group to the second adenine residue within the EcoRI recognition site (GAATTC). The discovery of restriction enzymes is a wonderful lesson on the unexpected outcomes of basic research. The enzymes were discovered by scientists studying the infection of bacteria by viruses. As the scientists began to understand how bacteria protected themselves against viruses using these enzymes, the idea of using restriction enzymes for manipulating other DNAs was developed. Research on restriction enzymes led to a Nobel prize for Arber, Nathans, and Smith in 1978. Several hundred restriction enzymes have been isolated from prokaryotic organisms, and many are commercially available. Restriction enzyme names follow a standard nomenclature system: • The first letter is the initial letter of the genus name of the organism from which the enzyme is

isolated. • The second and third letters are the initial letters of the organism’s species name. • A fourth letter, if any, indicates a particular strain of organism. • Roman numerals indicate the sequence in which different endonucleases were isolated from a

particular organism and strain.

EcoRI HindIII E = genus Escherichia H = genus Hemophilus co = species coli in = species influenzae R = strain RY13 d = strain d I = first endonuclease isolated III = third endonuclease isolated

Each restriction endonuclease scans along a DNA molecule, stopping only when it recognizes a specific sequence of nucleotides. Most restriction enzymes recognize a four- or six-base pair (bp) sequence. At or near the recognition site, the enzyme catalyzes a hydrolysis reaction that breaks the phosphodiester linkage on each strand of the DNA helix. Two DNA fragments are produced, each with a phosphate at the 5' end and a hydroxyl at the 3’ end. Restriction enzymes cut both strands of the double helix. In some cases, the enzyme cuts at the midpoint of the recognition sequence. These enzymes leave DNA fragments with blunt-ends.

Example of blunt end digestion by EcoRV (recognition sequence GATATC) 5'- N-N-G-A-T-A-T-C-N-N-3' 5'-N-N-G-A-T A-T-C-N-N-3' 3'- N-N-C-T-A-T-A-G-N-N-5' 3'-N-N-C-T-A T-A-G-N-N-5' Other endonucleases cleave each strand off-center in the recognition site, at positions two-to-four nucleotides apart, creating fragments with exposed single-stranded ends. This leaves single-stranded “overhangs” on either the 5’ or 3’ ends of the DNA fragments. Single-stranded overhangs, also called “sticky” ends, are useful in making recombinant DNA molecules because they can hydrogen bond to each other efficiently. For example, EcoRI recognizes the six-base sequence GAATTC and it cuts leaving a 5’ overhang of four nucleotides (AATT).

Example of staggered (sticky) end digestion by EcoRI (recognition sequence GAATTC) 5'- N-N-G-A-A-T-T-C-N-N-3' 5'-N-N-G A-A-T-T-C-N-N-3'

+

+


3

Lab #3

DNA Mapping Lab

3'- N-N-C-T-T-A-A-G-N-N-5' 3'-N-N-C-T-T-A-A G-N-N-5' Regardless of the source of DNA - whether from prokaryote or eukaryote - EcoRI will always cut at the GAATTC recognition site and it will always produce fragments with 5'-AATT overhangs. When the enzyme is removed, however, the overhangs can hydrogen bond (anneal) to each because of complementary base-pairing. Any EcoRI fragment can anneal to any other EcoRI fragment. After annealing, the phosphate backbone of the two DNA strands can be repaired by the enzyme DNA ligase. These steps are the basis of recombinant DNA technology. For example, if an EcoRI fragment of a yeast gene is annealed and ligated to a jellyfish GFP gene EcoRI fragment, the resulting chimeric DNA would be a novel part yeast-part jellyfish gene.

Recombination of two EcoRI fragments 5'-N-G 5'-A-A-T-T-C-X-3' 5'-N-G A-A-T-T-C-X-3' 5'-N-G-A-A-T-T-C-X-3' 3'-N-C-T-T-A-A G-X-5' 3'-N-C-T-T-A-A G-X-5' 3'-N-C-T-T-A-A-G-X-5' Agarose Gel Electrophoresis Digestion of a large piece of DNA with a restriction enzyme will generate smaller DNA fragments, whose sizes can be determined by gel electrophoresis. Electrophoresis literally means, "to carry by electricity". In gel electrophoresis, molecules are separated based on their charge and their size. DNA, an organic acid, is a highly negatively charged molecule due to the phosphate backbone. In solution, hydrogen ions are liberated from the phosphate groups, leaving negatively-charged oxygen ions radiating along the outside of the DNA molecule. When placed within an electrical field, the negatively charged DNA molecules are attracted toward the positive pole and repelled from the negative pole. DNA migration takes place through a gel matrix that acts as a molecular sieve to sort restriction fragments by size. The two most common matrix materials are agarose, a highly purified form of agar, and polyacrylamide, a synthetic polymer. As DNA fragments move though the pores in the matrix toward the positive pole of the electrical field, the gel matrix (“the sieve”) impedes the movement of larger fragments. Fragment mobility is inversely proportional to molecular weight (number of base pairs). Smaller restriction fragments will migrate exponentially further from the origin compared to larger fragments. Optimal separation of DNA fragments can be achieved by adjusting the concentration of agarose in the gel. A relatively low concentration of matrix material produces a loose gel, which separates large fragments effectively. A high concentration produces a stiff gel with a tighter matrix that resolves small fragments. Best separation is obtained by running the gel at low voltage over a period of several hours. Agarose gels are most useful for separating DNA molecules from 200 bases to about 20 kb. Above 30-40 kb, it is difficult to resolve bands well because all molecules above that size are retarded nearly equally by the sieving action of the agarose. However, by applying pulses of current alternating in an orthogonal arrangement, large molecules can be resolved. The pulses make the DNA snake through the agarose "sieve". With pulse-field electrophoresis, it is possible to separate DNAs that range in size from 5 kb to 6 Mb (6 Mb = 6000kb). It is even possible to resolve intact chromosomes.

Staining DNA DNA is colorless thus the DNA fragments are not visible within the gel. To follow how far the DNA has migrated through the gel, a colored DNA loading dye is added to the DNA samples. For example, the dye bromphenol blue migrates at about the same rate as a DNA molecule of 300-400 bp. Loading dyes also contain sucrose or glycerol, which increase the sample density, and facilitates loading the gel.

yeast EcoRI fragment

GFP EcoRI fragment

Annealed EcoRI sites

Ligated EcoRI sites

note the ligated backbone

O- O- O- | | | O-P-O-R-O-P-O-R-O-P-O || || || O O O

_

+

origin


4

Lab #3

DNA Mapping Lab

Following electrophoresis, the DNA bands themselves are visualized using a fluorescent stain, ethidium bromide or SYBR Green, which becomes highly concentrated in regions where it binds to DNA fragments. When exposed to medium wavelength ultraviolet (UV) light, the DNA/ethidium bromide complex strongly absorbs UV light at 300 nm, exciting the ethidium molecule which re-emits visible light at about 590 nm. Thus, stained DNA appears as fluorescent orange-red bands in the gel.

It is important to understand that a band of DNA seen in a gel is not a single DNA molecule. Rather, the band is a collection of millions of DNA molecules, all of the same base pair length and thus having traveled at the same average speed. Gel electrophoresis can also be used to purify restriction fragments. Following electrophoresis and staining, the DNA band containing the fragment is cut out and eluted from the gel slice. After extracting the ethidium bromide stain, the DNA fragments can be joined to other DNA fragments to create recombinant DNA molecules. Cloning vectors Chromosomes are large DNA molecules, ranging from thousands of bases in viruses to hundreds of million bases in higher eukaryotes. Because intact chromosomes are too large to map, the large chromosomal DNA is first broken into smaller fragments. Each of the smaller fragments is then mapped and the overall map is pieced together from overlapping patterns in the smaller maps. In order to obtain enough DNA mass for mapping studies, the small DNA fragments are first cloned. Cloning is a method for replicating a piece of DNA many times, yielding large amounts of DNA. Cloning is frequently carried out using small circular DNAs called plasmids. Plasmids used for cloning are also called vectors since they carry the DNA. Two features of plasmid vectors that facilitate cloning are: • An origin of replication (ori), which allows the plasmid

DNA to be replicated in E. coli. • A multiple clone site (MCS) with sites for several

commonly used restriction enzymes. Sometimes this region is called a polylinker. Eleven different restriction sites are clustered next to each other in the ~100 bp region spanning from the EcoRI site to the HindIII site. Since the sites within the MCS are present only in the MCS (and no where else in the vector), DNA fragments can be inserted into these sites without affecting other regions of the vector.

FOR THIS LAB, an understanding of the multiple cloning site will be important in mapping the plasmids. The plasmids you will receive contain fragments of chloroplast DNA derived from the single-cell alga, Chlamydomonas. Cloning the plasmids was achieved in three steps:

EcoRIKpnI BamHI SalI PstI

HindIII

HindIIIEcoRI

pUC19

ori

ampR

gene

XbaI BspM1SmaI

SacI SphI

MCS

Combine maps from overlapping patterns

A B C A A B

C A B

A B C A

C A A


5

Lab #3

DNA Mapping Lab

(1) The chloroplast DNA was cleaved into smaller fragments using one of the enzymes present within the MCS of pUC19.

(2) The vector DNA was cut with the same enzyme, (3) Cut vector DNA and cut chloroplast DNA were mixed together and DNA ligase was added (to join

the vector and chloroplast DNAs). The ligated DNAs were propagated in E. coli.

Restriction Maps A restriction map is a diagrammatic representation of a DNA molecule showing the sites of cleavage by different restriction enzymes along with the distances (in kilobase pairs) between those sites. A restriction map for a circular piece of DNA (plasmid) is shown to the right. (E = EcoRI site; P = PstI site; B = BamHI site). If a complete DNA sequence is known, the sequence can be searched for specific enzyme recognition sites and a map can be drawn based on the distances between these sites. If sequence information is not available, however, a map can still be generated by comparing the pattern of DNA fragments produced when the DNA is cut singly versus in combination with different enzymes. This is what you will do in lab.

E

EB

P

1.5 kb

1.0 kb

1.8 kb

2.0 kb

6.3 kb

1. Break chloroplast DNA into small fragments with enzymeX 2. Digest vector DNA with enzymeX

X X X X Z

Y X

3. Ligate cut chloroplast DNA to cut vector DNA. Note that the vector band is constant (~ 3 kb) in each of the re-joined DNAs, however, the insert size varies.

Z

X

Y X

X

Z Y

X

Z

X

Y X

Y X

X

Z

Note how cutting at site X leaves two ends, each able to join to another piece of DNA cut with enzyme X.


6

Lab #3

DNA Mapping Lab

There are a few simple rules for mapping: a) Digests using a single enzyme show how many sites are present. For

example, in the digest pattern shown to the right, a linear DNA cut with EcoRI yields two fragments of 2 kb and 4 kb lengths, revealing that there is one EcoRI site in the piece of DNA. Cutting with HindIII (H) also results in two fragments of 1kb and 5 kb, so there is also one HindIII site in the DNA. The question then becomes, “Is the HindIII site in the 2 kb or the 4 kb EcoRI fragment?”

b) Cutting with two enzymes shows where the sites are relative to each other. The key to mapping is to identify the bands that change between the single digests and the double digests. The double digest produced three fragments - 1 kb, 2 kb and 3 kb long. A little bit of logic helps to order the sites relative to each other.

a. Because the 4 kb EcoRI band disappears in the H+E lane, yet the 2 kb band remains, then the HindIII site must lie in the 4 kb EcoRI band. Note that the 3 kb and 1 kb bands in the H+E lane add up to 4 kb.

b. The HindIII site must be 1 kb from the end of the DNA because digesting with HindIII alone yields a 1 kb fragment. If the HindIII site were 1 kb from the EcoRI site, then digestion with HindIII alone would be expected to produce two bands, both 3 kb long.

c. The same logic can be used to compare the HindIII alone versus the H+E lanes to show that the EcoRI site lies in the 5 kb HindIII fragment.

A map then can be drawn, as shown to the right. This example was a simple case showing sites for only two enzymes. Maps usually are constructed by looking at the digest patterns of several different enzymes.

Making a restriction map using several enzymes Below is a diagram showing the digestion pattern of a plasmid (circular DNA) cut with the enzymes EcoRI, BamHI and PstI. The digests are carried out with each enzyme alone and then with different combinations of the three enzymes.

B

6.3 kb

2 kb

B/E

1.5 kb

2.5 kb

E

3.8 kb

P/E

1.0 kb

P B/P

3.5 kb

B/P/E

2.8 kb

From the pattern it can be seen that BamHI and PstI each cut the DNA once, yielding a 6.3 kb DNA (this is the total size of the DNA). EcoRI alone, however, yields two bands of 3.8 and 2.5 kb. This reveals that there are two EcoRI sites in the DNA. Note that the total equals 6.3 kb (3.8+ 2.5). The sum total of the fragment sizes in every lane should always add up to the same value - the size of the uncut DNA.

LEGEND: B = BamHI B/E = BamHI + EcoRI E = EcoRI P/E=PstI + EcoRI P = PstI B/P = BamHI + PstI B/P/E = all three enzymes

E H+E H

5 4 3 2

1

E H

1 kb 5 kb

4 kb 2 kb


7

Lab #3

DNA Mapping Lab

The single digests, however, don’t tell us where the sites are placed relative to each other. This information comes from the double digests (B/P, B/E, E/P) and the triple digest (B/P/E). Notice in the B/E double digest that the 2.5 kb band seen in E alone is still present but the 3.8 kb EcoRI band is gone and two new bands appear of 2 kb and 1.8 kb (= 3.8 kb). Thus, there must be a BamHI site within the 3.8 kb EcoRI fragment. The BamHI site is 1.8 kb from one EcoRI site and 2 kb from the other EcoRI site. However, at his point you cannot tell which EcoRI site is closest to the BamHI site.

Orientation 1 Orientation 2

This ambiguity can be resolved by looking at other enzyme digest patterns and by mapping the BamHI site relative to these enzymes. For example, the P/E digest shows that the PstI site lies within the 2.5 kb EcoRI fragment and the P/B digest shows how far apart the BamHI and PstI sites are from each other. Taking into consideration the information from all of the double digests, one can construct a map with sites as shown to the right. Occasionally, ambiguities in the restriction map cannot be resolved using only a few enzymes, especially when the digests produce several small fragments. Different combinations of restriction enzymes may help to resolve the ambiguity. It is not uncommon to map at least four to five enzyme sites. Another way to simplify the mapping is to subclone fragments of the DNA into a vector and then to map the smaller fragments. REFERENCES: Micklos D. & Freyer, G. (1988). DNA Science: A First Course in Recombinant-DNA Technology pub. Cold Spring Harbor Laboratory.

E

EB

P

1.5 kb

1.0 kb

1.8 kb

2.0 kb

6.3 kb


8

Lab #3

DNA Mapping Lab

Hazards • WEAR GLOVES FOR ALL THE PROCEDURES. First, material from your hands can contaminate

the enzyme digest and potentially degrade the plasmid DNA. Second, ethidium bromide (EtBr) will be used to stain the DNA. This compound can intercalate between bases and it is a possible mutagen. Wear a lab coat and gloves when staining your gel. Clean up all EtBr spills immediately and notify the TA. Avoid contact with skin, mouth or eyes.

• The electrophoresis is carried out at high voltages. Take care when working with the gel boxes and power supplies to avoid electrocution.

• The UV transilluminators we will use have safety locks to shut off the UV lamps to prevent inadvertent exposure to UV light. Nevertheless, keep in mind that UV light can damage your eyes.

NITRILE GLOVES ARE AVAILABLE FOR INDIVIDUALS WITH LATEX ALLERGIES.

Protocol A) Restriction Analysis of Plasmid DNA Restriction enzymes must be treated with special care. Since they tend to be unstable at higher temperatures, all enzymes are stored at -20° C and kept in a bench top cooler (DYNACHILL™) during use. The cooler is more efficient than ice in keeping the enzymes cold. 1. Cast 0.9% Agarose Gel Before lab, your TA will prepare a solution of 0.9% agarose. The agarose is dissolved by boiling in 1X TBE (Tris/Borate/EDTA) running buffer. Since the gel needs to harden for about 30-40 minutes, begin with this step.

1. Make sure the gel tray is clean. Wipe it with a damp Kimwipe and then dry it with a Kimwipe.

2. At the ends of the gel casting tray are gates. Raise these gates, making sure that the bottom of the gate is flush with the bottom of the gel casting tray. Tighten down the screws to fix the gates in place. Note: If the gates protrude beneath the bottom of the tray, the tray will not sit flat on your bench. This will lead to a gel that is thicker on one side, which can result in aberrant DNA migration. Also, Do not over-tighten the screws, as they can easily be stripped!

3. Insert a 12-toothed comb into the casting tray.

4. Place the tray on the lab bench and check to make sure it sits flat. If not, adjust the gates. Do not cast the gel on the bench liner; it may not sit flat on the liner.

5. Obtain a flask of molten agarose from the 55°C water bath at the back of the room. CAUTION: the flask will be a little bit hot. Use an oven glove or rubber HotHand™ to hold the flask.

6. Carefully pour enough agarose solution into the casting tray to fill it just to the height of the ends of the casting tray - do not overfill the tray creating a curved meniscus. Do not move or jar the casting tray while the agarose is solidifying. As it gels, the agarose will change from clear to opaque.

7. Return the agarose to the 55°C waterbath (place a lead donut on the flask to prevent it from tipping over into the water).

2. Prepare Restriction Digest (WEAR GLOVES) 1. Label ten 1.5 ml microfuge tubes A through J; label the cap and the side of the vial. Also, label the

side of the tube with the enzymes to be used for each digest (see the table below).


9

Lab #3

DNA Mapping Lab

You will be pipetting 1 µl of enzyme into the reactions. It is critical that you pipet the enzymes correctly. If you use too much, you will run out of enzyme and excess glycerol may inhibit the enzymes. Too little enzyme and you will not cut the DNA completely. Test what 1 µl looks like by pipeting some water. You should be using a P-20 for setting up the reactions. Control the plunger carefully - make sure you are only going to the first stop when you are drawing up enzyme. The enzymes are kept in 50% glycerol, which makes the solution more viscous, so pipet the enzymes slowly. Check each time that there really is enzyme inside the tip and in a constant amount.

Rxn Enzyme H20 Rxn Buffer

DNA Bam Eco H3 Pst

1 A BamHI 13 µl 2 µl 4 µl 1 µl -- -- --

2 B B+E mix 12 µl 2 µl 4 µl 1 µl 1 µl -- --

3 C B+H mix 12 µl 2 µl 4 µl 1 µl -- 1 µl --

4 D B+P mix 12 µl 2 µl 4 µl 1 µl -- -- 1 µl

5 E EcoRI 13 µl 2 µl 4 µl -- 1 µl -- --

6 F E+H mix 12 µl 2 µl 4 µl -- 1 µl 1 µl --

7 G E+P mix 12 µl 2 µl 4 µl -- 1 µl -- 1 µl

8 H HindIII 13 µl 2 µl 4 µl -- -- 1 µl --

9 I H+P mix 12 µl 2 µl 4 µl -- -- 1 µl 1 µl

10 J Pst I 13 µl 2 µl 4 µl -- -- -- 1 µl

To each tube, you will add the reagents listed in the table above. Add them in the order listed (water first, then buffer, then DNA and finally add the enzymes). Cross off each reagent as you add it to the reaction mixture. Mistakes in this lab occur usually by adding reagents to the wrong tube or forgetting to add a reagent.

2. Add water to each tube [use the Molecular Biology grade (“MilliQ”) water that we supply to you in 1.5 ml microfuge tubes] – note, you can use the same tip for each tube.

3. Add reaction buffer to the water in each reaction tube. (Buffer must always be added before the enzymes.) You can use the same tip for each tube. Be sure to place the tip into the water when expelling the liquid. This will help the 2 µl drop come off the tip.

4. Add plasmid DNA to each tube. In this case, since the buffer is the same for all samples, you can use the same tip for each tube. If the buffer varied, it is best tot change tips.

5. Add enzyme, using a fresh tip for each addition of enzyme. RETURN the enzymes promptly to the DYNACHILL coolers. NOTE: Be careful not to cross contaminate the enzyme stock solutions. Change tips after each addition of enzyme!

6. Close the caps and gently flick the tubes to mix the reaction.

7. Spin the tubes for a 1-2 second pulse in the microfuge. Make sure your tubes are placed in a balanced configuration in the rotor. To pulse centrifuge, hold down the button between the time and speed dials (The button may be labeled “SHORT” on some microfuges).


10

Lab #3

DNA Mapping Lab

8. Place the reaction tubes in a heat block that has been equilibrated to 37°C and incubate the reaction for at least 45 minutes (if there is time, let it go for an hour – longer is better).

Setting up the gel in the gel box:

1. When the agarose is set, unscrew the ends of casting tray enough to allow the ends to drop, and then re-tighten the screws to keep the ends held down. Do not remove the ends - just loosen them enough so that they can be pushed down completely. If the gate is not completely pushed down, it will partially block the gel and the DNA migration may be affected.

2. Place the tray on the platform of the gel box, with the comb end of the gel near to the negative end (black) electrode. Remember DNA will move toward the positive electrode.

3. Fill the gel box with just enough 1X TBE buffer to cover the surface of the gel. The 1XTBE will be in a 10L carboy at the back of the room. The buffer level should only be a few millimeters higher than the surface of the gel. If you put too much buffer in the box, remove some of the buffer. Excess buffer will slow the DNA migration down.

5. Gently remove the comb, taking care not to rip the gel. The buffer solution helps lubricate the gel. It may help to remove the comb by starting to pull the comb on one side of the gel and then removing the rest of the comb at a slight angle.

6. Make certain that the wells left by the comb are completely submerged. If you notice “dimples” around the wells, slowly add buffer until they disappear.

7. Place the gel box on a sheet of dark paper; the wells are easier to see against a dark background. 3. Loading the gel 1. Obtain a tube of MW (molecular weight/size) standards from the TA. The size standards have

already been mixed with loading dye. You will use 15 µl of the MW std per lane.

2. Remove the microfuge tubes from the 37° C temperature block. Add 4 µl of the blue loading dye to each digest, using a separate tip each time. IMPORTANT: DO NOT add the MW std to your tubes.

3. Briefly, pulse-spin the tubes in the microfuge.

4. Load 15 µl of the digestion reaction (use the P20). Use a new tip each time. If you use a different loading order, be sure to record it. • Make sure there are no air spaces in the pipette tip. • Steady the pipetman over the well using two hands to keep the pipetman vertical. • Be careful not to punch the pipet tip through bottom of gel. • Gently depress the plunger slowly to expel sample into the appropriate well. If the tip is

centered over the well, the reaction solution will sink to the bottom of the well. • When expelling the liquid, only press down to the first stop. It is better to leave a little liquid

in the tip rather than pipetting a big bubble into the well.

Well Sample Well Sample 1 MW 7 Tube F – EcoRI+HindIII 2 Tube A – BamHI 8 Tube G – EcoRI+PstI 3 Tube B – BamHI+EcoRI 9 MW 4 Tube C – BamHI+HindIII 10 Tube H – HindIII 5 Tube D – BamHI+PstI 11 Tube I – HindIII+PstI 6 Tube E – EcoRI 12 Tube J – PstI


11

Lab #3

DNA Mapping Lab

4. Electrophoresis 1. Attach the top of the gel box. Try not to move the gel box too much or the samples may come out

of the wells. Connect the electrical leads to a power supply - anode to anode (red to red) and cathode to cathode (black to black). Make sure both electrodes are connected to one channel of the power supply.

2. To set Constant Voltage on the Hoefer power supply, turn the Voltage Adjust Knob fully counter clockwise and the Current Adjust Knob fully clockwise. On the BioRad power supplies, select the voltage button and change the voltage using the up and down arrow key pads.

3. Turn the power supply unit on, and then turn the Voltage Adjust knob until the power supply is set to 90 volts. If the current reading is zero, then have a TA verify your electrophoresis configuration. Lack of current is usually due to (a) improperly connected lid or electrodes, (b) or too little buffer in the gel box. Shortly after the current is applied, you should see the blue loading dye moving through the gel towards the positive side of the gel box (red electrode). Check you r gel ~ 5 minutes after starting it to make sure the sample is moving in the right direction.

4. Electrophorese for at least 60 minutes or until the blue dye has moved about three-quarters down the length of the gel. (Separation will have occurred when the blue dye has moved 4-8 cm. from the well; however, the longer the run the better, the resolution of the bands). Check with a TA before stopping the run.

5. Turn off the power supply and disconnect the leads.

Wear gloves, glasses, and lab coats

5. Staining 1. Carefully remove the tray from the gel box and slide the gel onto a piece of plastic wrap. Flip the

gel over on the plastic wrap so the bottom side of the gel is facing up.

2. WEAR GLOVES: Obtain a sheet of EtBr Instastain from the TA – the Instastain is coated with the mutagenic dye ethidium bromide so handle it carefully and treat your gloves as if they are contaminated with ethidium bromide when you are done.

3. Using a transfer pipet, pipet a few drops of 1XTBE buffer onto the top of the gel (use some of the buffer from your gel box).

4. Peel away the transparent plastic backing from the Instastain sheet. Discard the transparent backing in the EtBr waste bag in the hood.

5. Lay the Instastain sheet on top of the gel (PRINTED SIDE UP) and firmly run your fingers over the entire surface of the Instastain sheet. Do this several times.

6. Cover the Instastain sheet with plastic wrap and place an empty gel-casting tray on top of the gel (this will maintain contact between the gel and the Instastain sheet). Place a light weight on the tray (e.g. a dropper bottle or a small flask). Stain the gel for 10 min.

7. Remove the Instastain sheet and discard the sheet in the EtBr waste bag in the hood.

8. Carry the plastic wrap with the gel to the TA for photography.


12

Lab #3

DNA Mapping Lab

5. Photography Place the gel on the UV transilluminator and close the lid. Turn on the transilluminator and

examine the gel.

Use the Image Capture System to obtain a permanent record of your gel. Make sure the TA has checked your digest pattern before leaving.

Place the gel in the GEL WASTE BAG in the hood.

CLEANING UP

Rinse gel boxes, trays and combs with DISTILLED water. Leave the box at your bench to dry. The distilled water faucet is the curved neck faucet, usually on the left hand side of the sink.

Please place the gel comb in the tray - this reduces the chance of the comb being lost.

Discard pipet tip waste in the regular waste cans.

Wipe down your lab bench area, especially if you spilled any running buffer.

Please make sure your station has the following equipment/reagents: Gel Box Gel casting tray and comb – please place the comb in the tray so it is not discarded accidentally. Power supply Beaker for waste tips Microfuge tube rack


13

Lab #3

DNA Mapping Lab

RESTRICTION MAPPING LAB REPORT (30 PTS) Cover page.

Your name Your TA's name The name of your partner(s) The name/title of the Lab exercise The date you hand in the lab report

Results: (25 pts) Required figures/data

• Figure 1 (4 pts). Photo of your gel – gel should be annotated. • Figure 2 (4 pts). Calibration curve for your gel • Table 1 (3 pts). DNA size estimate of bands on your gel • Figure 3 (10 pts). Drawing of your final map (s). Include preliminary

maps showing the progress of your work in constructing the map. • (6 pts) Answers to the questions about vector size, insert size,

enzyme used for cloning the chloroplast DNA and relative orientation of MCS sites (see p. 19).

Discussion (3 pts): Qualitative Observations If you were not able to generate an unambiguous restriction map, you must also include a discussion. Why there was ambiguity and what you could do to resolve it. For example, consider:

Did you run the gel for a sufficient period of time? Did the gel run evenly? Are all bands sharp and resolvable (well separated from each other)? Did your digest have partially cut or uncut DNA molecules (i.e. did you put enzyme in the tube?)? Were your DNA fragment size estimates reliable?

RESTRICTION MAPPING ANALYSIS Include a printout of your gel (Figure 1). Annotate your gel thoroughly. Each well should be labeled - either with a number or with the enzymes used. If you number the wells, include a legend listing the enzymes used in each well. For example, lane 1. MW std; lane 2 BamHI. Label several of the bands in the size standard lane (e.g., mark the 0.5, 1, 2, 3, 5, 7 and 10 kb bands). The 1 kb plus ladder fragments can be identified by the pattern of bands. In particular, there are two closely spaced bands at 1.65 kb and 2kb that will help you orient yourself. Bands above 2 kb increase in 1 kb increments, up to 12 kb. We have software to help you in identifying the band sizes. In theory, the software will do it faster than by hand but there may be a bit of a learning curve as you become familiar with the commands. You are free to do the analysis either way - using the band matching software or by hand. You may also come in anytime before or after lab hours (1 – 5 pm) to use the software.


14

Lab #3

DNA Mapping Lab

Measuring using the BioRad Quantity One software. (1) Download the gel image file from the Hal Server

Click on the PUBLIC icon on the desktop (or select PUBLIC from Recent Servers under the Apple Menu). Open the folder for the course (20191 or 20181). Check with your TA if you cannot find your file. Download the file to your computer. You can also save your file to a USB memory stick (E.g. DiskonKey) or to a zip disk.

(2) Open the QuantityOne software and then open the gel file. Open the Band Analysis Quick Guide.

Transform Show/Hide overlay

Image Toolbar

Text overlay

Lane Toolbar

Band Toolbar

Band Analysis Quick Guide


15

Lab #3

DNA Mapping Lab

(3) Optimizing the signal

a. Click on the TRANSFORM button (#3 in the quick guide) b. If necessary invert the image (so you see black bands on a white background) c. Click on AUTO-SCALE. This should optimize the range of band intensities. You can

also change the range using the high and low bars or using the gamma slider. If you don’t like the changes, you can always reset the image to its original state.

(4) ANNOTATION: You may annotate your gel by hand or with the software. To annotate the file,

open the Text Overlay Toolbar.

Click on the ABC button. Label the lanes and mark a few of the 1 kb ladder sizes (0.5 kb, 1 kb, 2 kb). You can reposition the test by dragging it; you can also align the different labels relative to each other. Print a copy of the labeled gel. The Print command is found under the File Menu.

(5) FRAMING THE LANES: Click on Frame Lanes (#4 in the Quick Guide) or open the Lane toolbar and select Frame Lanes.

a. Set the frames for 12 wells. b. Drag the corners and sides of the

frame guides until the lines run down the center of each well. Position the top of the frame a little under the wells (otherwise the software will identify the well positions as bands). The frame guidelines do not have to be perfectly centered on the lanes. When the frame guides are lined up with the lanes, click OK.

These buttons align text labels

Drag at the corners to position the frame

Frame Lanes


16

Lab #3

DNA Mapping Lab

(6) DETECTING BANDS: Click on Detect Bands (# 8 in the Quick Guide) or open the Band Toolbar and select Detect Bands

a. Band identification

i. LOAD the identification settings file (CS band ID) or manually enter the settings (Sensitivity =2, lane width = 3 mm). Press the DETECT button at the bottom of the window. You can also lower the sensitivity using the arrow buttons by the sensitivity setting. Reduce the sensitivity until there are no or only a few false positives.

ii. Click on ACCEPT. b. Editing the band assignments

i. Smudges on the image may result in false band calls. To remove miscalls, click the Remove Band button. Then, position the pointer over each band and click on the band to remove it. Bands due to partial digestion products should also be removed.

ii. Remove any bands identified at the origin (well positions). iii. Missed bands can also be added to the analysis using the Add Band button.

Position the pointer over the band and click on the band to add it. iv. Sometimes, the line marking the band does not overlay the band properly. You

can adjust the line position using the Adjust Band button. Drag above or below the band and adjust the band position. A densitometric trace of the lane will be shown to help you align the marker line with the peak of the band intensity.

(7) ASSIGNING STANDARDS – click on the standards button (# 9 in the Quick Guide or from the Match Toolbar)

a. Load the 1kb PLUS ladder as the standard. This file contains all the band sizes in the

ladder. A new standard window will appear (see following page). b. This part can be tricky. Often the higher MW standard bands (bands over 8 kb) on the

gel are so close together that they are not clearly resolved from each other. The Detect Band routine may miss some of these closely spaced bands and the software will mis-assign band sizes. To avoid this problem, you can set the starting band from which standards are matched. Count up from the 2 kb band until you find the highest band that was identified as a band by the software (this will likely be the 7kb or 8kb band).

Detect Bands

Add a band

Remove a band

Adjusts the position of the assigned band

Match to standard


17

Lab #3

DNA Mapping Lab

Then, select that band type by clicking on the arrow by the name of the band in the “standard” window. Finally, go back to the gel image and click on that band in the gel. Now all bands lower than the selected starting band should be matched correctly. For example, if the 7 kb band was the highest band that was marked, then you would first click on the arrow by 7kb (band type 6) in the 1kb plus ladder set and then click on the 7 kb band on the gel image. Now that band should be assigned as the 7 kb band and all the bands below it will be automatically assigned.

c. Since you ran two standard lanes, assign the 1kb plus ladder to each standard lane. The software will use both standards to compensate for uneven DNA migration across the gel.

d. To look at the calibration curve, select the calibration curve button and then click on the lane with the standards. Use the semi-log point-to-point analysis; check the Show Size option. If you do not see a smooth curve, check the band assignment for the standards. Print out a copy of the calibration curve for one of the molecular weight standard lanes.

Assign standard to lane Calibration curve

Click on one of these “band types” first and then when you assign the standards lane, click on the corresponding band on the gel.

Use the 1.6 kb and 2 kb bands to help orient yourself on the ladder. This doublet is separated from the other bands by a gap.


18

Lab #3

DNA Mapping Lab

(8) Click on BAND ATTRIBUTES (# 10 in the Quick Guide). Select Band Numbers. Print a copy of the gel (under the File Menu, select Print).

(9) You can also view the gel with the size estimates for each band on the gel. Under BAND ATTRIBUTES, select base pairs. The size of the bands in kilobases will be displayed on the gel image. Because the lanes are closely spaced, however, it may be hard to read the MW values. You can zoom up to see the sizes better. Print out a copy of the image with the size overlay if you like.

(10) Generate a REPORT: Click on #13 in the Quick Guide (All lanes report). Select Base Pairs, Rf, and band number to be displayed using very small for all settings (font,etc.). The report can be printed directly or it can be exported as an Excel file (To save paper, export the report as an Excel file; in Excel, under Page Setup, set the worksheet to print on one page. Save the exported Excel report (just in case) – include your name in the file name. If you like, you can email the file to yourself. The report will print out the BasePair size and Rf values for each band. The report will list the band numbers as well as the assigned sizes.


19

Lab #3

DNA Mapping Lab

Making the map Use the estimated fragment sizes to piece together a restriction map of your plasmid.

(1) Total the size of all the bands in each lane. They should add up to almost the same value - the size of the uncut linear DNA. Because of the logarithmic scale, errors in measurement are greater for large bands than for small bands. Thus, size estimates for the total plasmid size are determined more accurately by adding up the fragment sizes in a lane with smaller bands than in a lane with a single large band. For your analysis, you only want to include the bands that arise from complete digestion of the plasmid. These bands will tend to be the darker bands on the gel. However, faint bands may appear due to partial digestion of the DNA or due to "star activity". Star activity arises when an enzyme cuts at a different sequence than its canonical cutting site (for example, EcoRI star activity arises when EcoRI cuts at GAAATC in addition to cutting at its normal site GAATTC). Star activity often arises when buffer conditions (salt concentrations, pH) are not optimal for the enzyme and the stringency for recognizing the specific sequence is relaxed. In the figure to the right, the star activity bands are marked with asterisks; note that the intensity of the star activity bands is low relative to the desired digest bands. There should be a stoichiometric relationship between band size and band intensity. Larger fragments bind more stain and should always stain more intensely than a band of lower MW. In the figure to the right, note that the top star activity band is much less intense than the band just below it, even though the star activity band is larger. However, do not ignore the small bands on the gel (those bands <1 kb). Very short DNA fragments will be faint because they pick up less stain. Sometimes these small fragments are necessary to generate a map that makes sense.

(2) Explain the way in which you developed your map. Please include preliminary drawings of the map as it was being developed. This will show the logic you used to construct the map. These maps will help you if your final map is wrong. The TA will still be able to give you credit if some of the logic is shown. Clearly distinguish the "temporary" maps from the final map(s). Double check your final map . Predict the fragments you would get from digests based on your map. If the predicted fragments do not match the gel data, then there must be an error – either in your map or in the interpretation of the gel data (e.g. wrong estimates of band sizes, including star activity or partial digest bands, ignoring small “real” bands).

(3) Show sites for all of the enzymes tested (EcoRI, BamHI, HindIII, & PstI) (4) Show distances between several of the fragments. (5) Based on the total size of the plasmid, what is the size of the inserted chloroplast DNA fragment

and what is the size of the vector (pUC19)? (6) Recall that the plasmids were made by cloning chloroplast DNA fragments into the Multiple

Cloning site (MCS) of pUC19. The same enzyme was used to cut the vector and the chloroplast DNA. Can you determine which enzyme was used? [Hint: look at the digestion patterns for the different plasmids – sample gels for all plasmids are on the lab web site].

(7) 1 pt extra credit: Show the order of the MCS sites (EcoRI, BamHI, HindIII, & PstI) relative to the insert DNA. You will not have enough information to show the relative orientation of all the sites but you should be able to place some of them relative to each other. A circular map of pUC19 is shown on page 4 of the handout.


20

Lab #3

DNA Mapping Lab

Measuring by hand

Construct a calibration curve 1. Draw a horizontal line across the gel just below the bottom of the

wells. Carefully, measure the distance from the horizontal line to the leading edge of each size standard (1 kb ladder) band.

distance (mm)

MW std size (kb)

36 10.1 40 7.1

Make a table (Table 1) with distance in one column and size in the other column.

45 5.1

2. Plot the distance migrated on the X-axis against the log (MW) of the band on the Y-axis. DNA migration in an electrical field is defined by the equation:

distance migrated~1/log molecular weight NOTE: If you use semi-log graph paper for this plot, you do not need to calculate the log (MW); simply plot the MW on the LOG scale axis.

3. Draw a smooth line through the points. Note, that the plot curves up near the high MW bands and it curves down near the low MW bands. Do NOT calculate a best-fit line (linear regression) for the curve. Because of the curvature, a best-fit line will under- or over- estimate small and large fragment sizes (values in the curved regions).


21

Lab #3

DNA Mapping Lab

4. Next, measure the distances for the digest bands; enter these values into a table. (See the note in the

“ Making the map” section about faint bands and star activity. 5. Once you have measured your bands, determine

their MW using the calibration curve. Locate on the X-axis the distance migrated by a fragment. Using a ruler, draw a vertical line from this point to its intersection with the calibration curve. Then, draw a horizontal line to the Y-axis and read off the MW on semi-log paper (or the LOG (MW) on regular linear graph paper).

6. Repeat this step for each DNA fragment, filling in the table with the estimated base pair sizes, in kilobases, for all of the DNA fragments.

LOG MW

DISTANCE (mm)


22

Lab #3

DNA Mapping Lab

Class example: We will generate a restriction map from the following data in class. Pre-lab exercise

• Use the semi-log paper (p. 23) to prepare a calibration curve from the labeled standards (left lane) • Estimate sizes for the bands in the EcoRI lane and in the E/B/P lane.

1.6

4.0


23

Lab #3

DNA Mapping Lab

dna lab: restriction mapping - biolympiads · cell and molecular biology autumn 2005 4 lab #3 dna...

Documents