Basic Lab Techniques Hhmi2

Princeton Molecular Biology Outreach Programs @ Springside School The goal of the experiments chosen for this Workshop is to illustrate the power of state-of-the-art molecular biological techniques. In an increasing number of newspaper and journal articles, websites, etc. about The Human Genome Project and other scientific advances, it should be clear that genes and genetics have become major topics of discussion in society today. All of genetics stems from the sequence of bases along the double helical DNA molecule. In addition to the obvious traits that are most frequently associated with genetics, such as eye and hair color, an increasing number of specific tests can detect precise sequences within regions of DNA molecules that have enormous implications for health and legal status. For example, accurate tests have been developed for the detection of numerous genetic disorders such as cystic fibrosis, sickle cell anemia and many others. In addition, analysis of the human genome has shown that different human genomes are 99.9% identical and that the divergent 0.1% is comprised mostly of point mutations known as single-nucleotide polymorphisms (SNPs). This discovery has led to the method of haplotype mapping, which correlates a specific pattern of SNPs in a particular chromosomal region with susceptibility to a certain genetic disorder, such as diabetes or age-related macular degeneration (AMD), the leading cause of blindness in people over 50. DNA analysis has also become very common in forensics. It is now possible to determine whether tissue samples as small as a single hair follicle or a smear of blood came from a certain individual or suspect, as you will learn during the Workshop. In addition, conservation groups now use forensics to combat illegal trade in protected animal species by identifying products that contain ingredients from rare species. Also, analysis of DNA obtained from a community of microbes (metagenomic DNA) is used to estimate the microbial diversity of natural environments of several niches in the human body. All this information requires a detailed understanding of DNA, gene structure, and some sophisticated technologies that have been developed only within the last 5 - 30 years. Yet, it is also critical to understand the limits of these technologies, which often determine the accuracy of the tests as well as their ethical implications. Some important questions are: Should an individual’s genetic information be private? How much should you know about your genetic makeup? How much should others be allowed to know about your genetic makeup? How should this information be used ethically? A thorough discussion of these ethical questions is obviously beyond the scope of this Workshop. However, the design of the experiments you will perform in the next will provide you with a basis for understanding the relationship between DNA structure, modern genetics, and several social and ethical concerns. In the laboratory, you should work with good laboratory technique at all times and keep your bench area clean and neat after each experiment by always: 1. Cleaning all lab equipment, including parts of your gel apparatus, and returning these to the proper places for storage. 2. Wiping your bench area using a paper towel and water. 3. Washing your hands when finished.

1

Basic Microchemical Techniques: Agarose Gel Electrophoresis & Micropipetting I. Introduction: The analysis of DNA, deoxyribonucleic acid, frequently requires determining the size of individual DNA molecules or even the nucleotide sequences of specific regions within DNA molecules. Under most conditions, DNA has a relatively uniform acidic or negative charge caused by the phosphate groups on the outer surface of the molecule. When placed in an electric field in a process known as electrophoresis, negatively charged DNA molecules will migrate toward the positive electrode. In a semisolid matrix like an agarose gel, larger DNA molecules will be retarded by the sieving properties of the matrix, so they move more slowly and migrate a shorter distance in a given time than smaller DNA molecules. Today, you will first pour an agarose gel and then use this gel for electrophoresis of several different DNA samples. After staining and photographing the resulting pattern of bands in the gel, you will use your photograph to analyze these DNA samples by determining the sizes of unknown DNA molecules. For this analysis, you will use two samples of DNA standards that contain molecules of known sizes to plot a calibration or standard curve. Some of the samples you will analyze have been digested with restriction enzymes, which recognize, bind to and cleave DNA at specific nucleotide sequences. Approximately 300 different restriction enzymes are currently commercially available, and these enzymes are very powerful tools currently available for analyzing DNA, as you have learned from your reading and will demonstrate to yourselves during this Workshop. In addition to electrophoresis, most of the procedures in this Workshop and also in research laboratories employ microchemical techniques; i.e., using very small amounts of reagents such as DNA and enzymes. Many reactions require the addition of only one microliter (1 µl, one one-millionth of a liter) of some components, so it is essential that you become familiar with measuring small volumes accurately and reproducibly. In addition, because many basic procedures in molecular biology use bacterial strains, particularly Escherichia coli (E. coli) a superstar in molecular biology, you will also become familiar with some basic techniques for working with microbes.

http://www.fao.org/docrep/005/AC802E/ac802e05.htm

2

GEL ELECTROPHORESIS & MICROPIPETTING I. Micropipetting: Micropipettors are precisely calibrated > instruments that combine the functions of a pipette and a pump. Disposable plastic tips make micropipettors reusable. To measure the small volumes used in molecular biology accurately, you will use the P20 and P200 Gilson adjustable micropipettors.

The P20 is used to transfer volumes between 1 and 20 µl, and the P200 is used for volumes between 20 and 200µl. It is preferable to use the P20 for volumes less than 20 µl, because the P20 measures small volumes more accurately than the P200.

The pushbutton at the top of the micropipettor has two working positions in addition to the fully upright position. The first position is used for picking up liquid, and the second position is used to > The P20 should NEVER be eject or deliver liquid from the tip. used for volumes greater than 20 µl nor the P200 for volumes greater than 200 µl because this will damage the pipettes.

1. To reach the first position, push the button down using relatively > If using your index finger is uncomfortable, your thumb can light pressure with your index finger until you meet some be used instead. resistance. 2. Reaching the second position requires significantly more pressure from your finger, so push the button down with more force to reach this position. 3. Change the volume setting of the scale on the front of the P20 to > The two black numerals represent integer microliters, read “1-0-0” from top to bottom using the black gears near the and the red numeral represents top of the pipettor. What volume does this represent? tenths of microliters. The reading “1-0-0” indicates 10 µl on the P20 as shown below:

1 0 0

Black Numerals Red Numeral

4. Everyone should practice pipetting using your P20 micropipettor, > The white tip has graduations at 10, 50 and 100 µl, with the mark the white graduated tips and the red liquid as described next. nearest the point of the tip being the 10 µl mark.

5. Put a white graduated tip firmly onto the end of the pipettor by > Using the pipettor to pick up tips is preferable to picking up a tip gently pushing the barrel of the pipettor into a tip in the rack.

with your hand to avoid contaminating the tip with microbes on your hand. Then use your fingers to push the tip gently but firmly onto the barrel of the pipette without touching the end of the tip.

3

6. Depress the button at the top of the pipettor until you reach the first > Be sure not to push the button down to the second stop position stop position and feel considerable resistance.

when picking up liquid or your measurement will be grossly incorrect.

7. Keep the button at this position and insert the tip into the red > Do not insert the tip too far into the solution or extra drops will solution. cling to the tip and make your measurement inaccurate.

8. With the tip still in the red liquid, SLOWLY release the pressure of > If you release the button too quickly, air bubbles could enter your finger and allow the button to return to a full upright position. the tip, and the volume of liquid The red solution will enter the tip. picked up will not be correct. Check the graduations on the tip to determine whether the volume you have picked up is correct.

9. Place the bottom of the tip just at or just above the meniscus of the solution to which you are transferring (or near the bottom of a tube if the tube is empty). 10. Depress the button all the way to the second stop position until all the red liquid is expelled. 11. Use new graduated tips to transfer 5 µl and 1 µl of the red > Observe the amounts of liquid in the tip for these measurements and solution to the 1.5 ml microtube. 12. Continue practicing with the red solution and a microtube until you can pipette accurately and easily.

try to remember how full the tip appears for these volumes. If you know approximately how much liquid should be in the tip, you will recognize when your pipettor misfunctions while you are pipetting instead of after an experiment fails.

13. Always use a new tip for each transfer because residual liquid on or in the tip from a previous transfer could contaminate the new solution or cause inaccurate measurements. 14. Micropipettors are very precise instruments that are expensive to purchase (more than $280 each), repair and recalibrate. There are three maneuvers that you should NEVER try with these pipettors: a. Never measure a volume greater than the maximum volume of a micropipettor, the volume indicated on the pushbutton. > Liquid will clog the pipette and

b. Never use the micropipettor without a tip.

ruin parts.

c. Never lay down a pipettor with a tip containing liquid > The liquid could run back into the pipette and damage it. attached.

4

Micropipetting Small Volumes: Each person should use the white > These exercises were adapted from Laboratory #1 of DNA graduated tips to perform the exercises described below. Science, pp. 327 – 328.

1. Use a black marker to label three clear 1.5 ml microtubes with the > The microtubes are in the whitish rack at your bench. letter “A”, “B”, or “C”. 2. Set your P20 micropipettor to 5 µl and add 5 µl of Solution I (blue) > Be sure you use a new tip each time you pipette. Use the chart to each microtube, as shown in the chart below. below as a checklist to ensure that you add the correct amounts to the reaction tubes.

Tube

Solution I (Blue)

Solution II (Red)

A B C

5 µl 5 µl 5 µl

4 µl 4 µl 4 µl

Solution III Solution IV (Green) (Yellow) 1 µl 1 µl 1 µl

Total Volume

--5 µl

3. Use new tips to add 4 µl of Solution II (red) to each reaction tube. 4. Use new tips again to add the indicated volumes of Solutions III (green) and IV (yellow) to the reaction tubes. 5. Close the tops of the tubes and mix the reagents by flicking each tube with your fingers. 6. To bring the contents to the bottom of each tube, place your tubes and your partner’s tubes in a microfuge and pulse spin them for a few seconds as described to the right. 7. Remove the tubes from the microfuge. 8. Label tube “A” with your initials and place it in the rack on the able at the front of the lab.

> Be sure there is a tube across the

rotor from each of your tubes for balance. To operate the microfuge, screw or press the top of the rotor on, close the lid, and spin the tubes for three seconds. Centrifuging for short times is accomplished easily by holding down the button on the front panel of the microfuge for the required time and then releasing it to terminate the spin.

9. Check the volumes in tubes “B” and “C” yourself as described to > Check the volumes by setting your pipette to the total volume that the right. 10. If you have pipetted accurately, the tip should be completely filled with liquid; there should be no air space at the end of the tip, and no liquid should be left in the tube. Be sure to check the volumes carefully because it is essential that you pipette accurately.

11. Discard tubes “B” and “C before proceeding.

5

should be present in the first tube. Then put a new tip on your pipettor, depress the pushbutton to the first stop, put the tip to the bottom of the tube and release the button slowly so all the liquid in the tube enters the tip. The tip should be completely full if the volume is correct. Repeat this for your other tubes.

Micropipetting Large Volumes: Each person will each now practice pipetting using his/her P200 pipettor. 1. Use a black marker to label three new (1.5 ml) microtubes “D”, > These tubes are in the whitish rack at your bench. “E”, or “F”. 2. Pipette the volumes shown in the chart below into the appropriate > Use a new tip for each transfer. tubes: Tube

Solution I

Solution II

Solution III

Solution IV

D E F

20 µl 20 µl 20 µl

40 µl 40 µl 40 µl

50 µl 50 µl 50 µl

30 µl 30 µl 90 µl

Total Volume

3. Close the tops of the tubes and mix the reagents by flicking each tube with your finger. 4. Spin the contents to the bottoms of the tubes for 3 sec in the microfuge. 5. Mark tube “D” with your initials and place it in a rack on the front table to be checked for accuracy. 6. Next, calculate what the volumes of tubes “E” and “F” should be and check the volumes in these tubes yourself to assess your accuracy. 7. If your volumes are not correct, be sure to ask a member of the lab staff for assistance. 8. Discard tubes “E” and “F” before proceeding.

waynesword.palomar.edu/lmexer3b.htm

6

III. Pouring an Agarose Gel: Agarose is a purified form of agar, a complex polysaccharide derived from seaweed. When boiled in Electrophoresis is the process by buffer, agarose dissolves and then solidifies upon cooling much like > which molecules are sorted by size Jell-O. Agarose gels for electrophoresis are prepared in a pH- iin a semi-solid matrix like agarose iin an electric field. buffered solution, which conducts an electric current through the gel The phosphate groups on the outside and provides the electric field through which the DNA molecules move. oof DNA give it a uniformly nnegative charge, so DNA migrates tttoward the positive electrode.

1. Prepare a 1% agarose gel (gel bed volume = 50 ml) by adding .5 g agarose powder to 49.5 ml of 1X electrophoresis buffer. 2. Wearing gloves and working with your partner, place a black dam in > The dams are asymmetric. The side of each dam that forms a right angle each of the two slots at the ends of the gel bed. with the bottom of the dam should f fface the gel bed as shown below to tthe left.

3. Obtain a flask of molten agarose from the water bath. 4. Use the orange Hot Hand mitts available beside the waterbath to hold the flask.

> Molten o1.0% agarose can be kept iin a a65 C water bath so it will not solidify.

5. Close the cap of the flask and swirl the flask to mix the agarose. 6. Use a transfer pipette to seal the ends of the gel bed as shown below > Although the gel apparatus does not lbreak often, it is best to be certain with a narrow bead of agarose before pouring your gel. Dam

Seal with Agarose

no leaks ever occur by sealing the gel bed. Do not overfill or underfill the gel bbed or your gel will be too thick or ttoo thin. Surface tension will make the gel appear thicker than it is. Observe the gel level closely while ppouring.

Dam

Gel Bed

7. After waiting one minute (min) while the beads solidify, swirl the > oPosition the comb, which is aasymmetric, so its teeth are nearest flask again and pour melted agarose into the gel bed. tthe black electrode.

8. Put a comb with 8 teeth in the slot at the end of the bed nearest the > Note that the gel box has three slots iin which a comb can be placed and negative (black) electrode.

that below each slot is a red stripe on the bottom of the gel bed. After tthe comb has been removed, these rred stripes will enable you to v visualize the wells easily.

9. Allow your gel to solidify at room temperature for 20 – 30 min.. 7

> When finished, it should look ocludy and be firm to the touch.

Gel Electrophoresis Pt. 2 >

1. Carefully remove the black dams.

Rinse the dams well at the sink with water from the white plastic faucets. Put them in the slots at the back of the gel box for storage.

2. Use the TAE (Tris Acetate EDTA) electrophoresis buffer that you prepared by diluting the concentrated 50X buffer to make a 1X solution. 3. Pour enough buffer into the gel chamber to completely cover your gel. 4. Let the buffer sit over your gel for a few minutes while each lab partner practices the gel loading process described next. 5. First, prepare the practice loading solution as described in the sidebar to the right. To make this solution, you will use loading dye, which contains a dye so samples are visible and a dense reagent so samples sink to the bottoms of wells when loaded. If there are drops of dye on the sides of the tube (clear tube, purple dot), spin the tube for a few seconds (sec) in the microfuge before making the practice loading solution. **some practice solutions are pre-mixed and ready to use. 6. Locate the practice gel.

Prepare approximately 400 ml per electrophoresis chamber and 100 ml per gel (this will give you some extra) 10 ml conc. buffer + 490 ml distilled water Practice loading solution: Pipette 180 µl H2O (clear tube) into a 1.5 ml microtube and add 20 µl loading dye (clear tube with purple dot). Touch the pipette tip containing the dye to the meniscus of the water before expelling the dye. Close the tube top and flick the tube vigorously to mix.

7. Pick up 20 µl of the practice solution with your P20 micropipettor. 8. Place the pipette tip just inside the top of a well or rest the tip against the upper edge of a well in the mock gel. 9. Push the button of the pipettor down slowly to carefully dispense the dye solution, allowing it to sink to the bottom of the well.

Do not actually put the tip into the well because you could puncture the well and lose the sample.

10. Load one aliquot of practice loading solution, but do not load the rest unless you feel confident that you can do this successfully.

Depressing the button of the pipettor too quickly could cause the sample to squirt out of the well.

11. When all samples are loaded, close the lid of the gel box and > begin electrophoresis as described below.

8

Each power supply can run two gels.

12. a. Check that the power switch is off and the rheostat is turned to the lowest setting. b. Attach the electrical leads, taking care not to jostle the box.

> Connect the leads to the same

colored plugs on the power supply, red to red and black to black. The red lead should be at the opposite end of the gel box from the wells of your gel.

c. Next, turn on the power and adjust the voltage to 100 – 110 V by turning the rheostat clockwise. d. Small bubbles will start to rise from the electrodes if current is > If the dye does not move, or if it flowing, and you will see the dye start to move out of the wells does not move toward the red electrode at the far end of your gel, within a minute or two. immediately turn the rheostat down and the power off.

13. Continue the electrophoresis for 30 – 50 min or until the dark blue dye has traveled to a point midway between the second and third red stripes, about 2.5 cm from the end of the gel.

14. Turn the rheostat counterclockwise to the lowest setting, turn off the power and unplug the leads from the supply. >

III. Staining and Photographing Your Gel: When electrophoresis of your gel ends, proceed to stain your gel with Methylene Blue Plus as described next. To visualize the DNA in your gel, you will stain the gel in a methylene blue based stain, Fast Blast (BioRad), or use SYBR stain (pre-measured into the gel when preparing the gel – 1ul SYBR stain per 10 ul of gel)

9

In our hands, Methylene Blue Plus and FastBast are about half as sensitive as ethidium bromide but has the great advantage of not being mutagenic so you can use it in your classrooms. SYBR stain is a great alternative, having almost the sensitivity f ethidium bromide combined with the safety of methylene blue based stains.

> Put on gloves before proceeding. 1. Before you stain your gel, each lab pair will be assigned a number. > To avoid confusion, please use the

staining and destaining trays with the number you have been assigned for your gels during the Workshop.

2.Lift the gel chamber out of the electrophoresis box and carry the chamber to the nearest sink. 3. Use your fingers to hold your gel against the gel bed and carefully pour the electrophoresis buffer into the sink. 5. Pour enough Methylene Blue Plus solution into a yellow staining tray to fill it half full. 6. Transfer your gel from the gel bed to the tray by lifting the red- > Rock the tray carefully a few times to ensure the gel is striped gel bed out of the chamber and slowly sliding the gel from submerged in the stain. the bed into the stain. 7. Note the number of the tray you are using. 8. In 30 min, when staining is complete, fill a white/clear destaining > As Edvotek recommends, staining for 30 min seems to work best. tray about half full of deionized water from the white faucets at Once the gel is well stained, DNA the sinks. bands become visible after only a 9. Use a spatula to remove your gel from the stain and transfer it to your destaining tray containing the water.

short 15 - 20 min destaining period. If staining is decreased to less than 30 min, a longer period of destaining is necessary for bands to become visible.

10. Take your gel in the destain tray to the sink. 11. Hold the gel in the tray with your fingers. 12. Pour off the destain water and rinse the gel several times with > Rinsing several times removes excess stain. fresh deionized water. 13. Return the tray to the hood and put a twisted paper towel in the > The towel absorbs stain and speeds up the destaining process. tray around the gel (not on top of it). 14. Let the gel destain for 10 - 30 min, changing the water a couple of times until the background is lighter and the DNA bands are readily visible. 15. While your gel is destaining, pour the stain back into the bottle in > Methylene Blue Plus can be reused several times. DNA bands the hood.

stained with Methylene Blue Plus will remain visible for several days when gels are stored at 4o C.

10

16. Wearing gloves, carry your gel in the destaining tray to the light > Take two pictures of your gel, as described next, so each partner has box. a copy.

17. Use a spatula to lift your gel from the tray, tipping the spatula slightly so excess water drains back into the tray. 18. Transfer your gel to the surface of the light box to observe and meausre your bands. You can also use the ProScope camera to capture an image of your gel and measure using Logger Pro. 26. Return your gel to the destaining tray and add some fresh water. 27. Put the tray in the refrigerator overnight inside a plastic Baggie.

http://www.edvotek.com

www.vernier.com 11

> Because the visibility of DNA bands stained with Methylene Blue Plus often improves with prolonged destaining, you will let your gel destain overnight tonight and photograph it again tomorrow morning.