Mastery of micropipetting and sterile technique is essential for reliable success in molecular biology. Take these exercises and our discussions about pipetting seriously. Micropipettors A micropipettor* is essentially a precision pump fitted with a disposable tip. The volume of air space in the barrel is adjusted by screwing the plunger in or out of the piston, and the volume is displayed on a digital readout. Depressing the plunger displaces the specified volume of air from the piston; releasing the plunger creates a vacuum which draws an equal volume of fluid into the tip. The fluid in the tip is expelled by depressing the plunger again, but to a further, second 'blow- out' stop. * The nickname is originally derived from a different pipettor brand ('Pipetman'), but still widely used by molecular molecular biologists for all micropipettors. Take the following precautions when using the pipettor: Never rotate the volume adjustor beyond the upper or lower range indicated on the pipettor. Never invert or lay down a pipettor with a filled tip (fluid can run into the barrel). Never let the plunger snap back after withdrawing or expelling fluid (could damage the piston or cause fluid to enter the barrel, and contact the piston aka fountaining ). Never reuse a tip that has been used to measure a different reagent, or that has entered a tube containing a different reagent. When in doubt, use a fresh tip. Pipet- Lite (LTS) Pipettors: Pipet- Lite (LTS) is a brand of micro- pipettor used in the laboratory. Pipet- Lite is an air- displacement pipette which uses a magnetic assist and is slightly different than the Gilson pipets used in other labs. Become familiar with this pipet and it s components. Setting Volume: Turn the volume lock counter- clockwise to unlock the pipet. The position shown at left below so the volume setting mechanism is unlocked and free to turn. With the mechanism unlocked, then rotate the plunger button to change volume counter- clockwise to increase, and clockwise to decrease volume. When setting the desired volume, first turn the knob slowly clockwise until the desired volume is displayed. Always dial down to the desired volume. Lock the pipet by turning the volume lock clockwise (see figure for locked position) 1
Operation: Press the plunger button tot the first stop, and hold it in this position. The magnetic latch will help you sense and hold this position. Holding the pipet vertically, place the tip into the sample at the proper depth (see tip emersion chart) and relax your thumb pressure on the plunger. Do not let go of the plunger button, or the piston may snap up quickly, resulting in inaccurate measurement. Pause briefly to ensure the full volume of sample is drawn into the tip. Withdraw the tip from the sample. If any liquid remains on the outside surface of the tip, wipe it carefully with a lint- free tissue, taking care NOT to touch the tip orifice. Touch the tip end against the side wall of the receiving vessel and press the plunger slowly, past the first stop. Wait one to two seconds. Still holding the plunger down, with draw the tip. Then release the plunger. Watch: (http://www.mt.com/us/en/home/supportive_content/specific_overviews/product- organizations/pipe/rainin- pipette- technique- videos.html) Lab Experiment 1 Measurements, Micropipetting, and Sterile Technique Part1 Micropipetting Reagents Tubes of colored Solutions I, II, III & IV Sterile LB broth I. Practice with a small- volume pipettor This exercise simulates setting up a set of typical small molecular biology reactions such as a small restriction enzyme digest, using a 10 µl pipettor. 1. Label three 1.5 ml microtubes on the lid with a permanent marker: A, B & C. 2. Use the matrix below as a checklist while adding solutions to each reaction tube. Use the techniques discussed above for pipetting. Tube Soln. I Soln. II Soln. III Soln. IV A 4 µl 5 µl 1 µl B 4 µl 5 µl 1 µl C 4 µl 4 µl 1 µl 1 µl 3. Set the micropipettor to 4 µl, and add Solution I to each reaction tube. 4. Use a fresh tip to add the appropriate volume of Solution II at a clean spot on reactions tubes A, B, and C. Eject used tips into a small waste tip beaker. 5. Use fresh tip to add the indicated volumes of Solutions III and IV to the appropriate tubes. 6. Close the tops. Pool and mix the reagents by placing the tubes in a balanced configuration in a microcentrifuge and running for a few seconds. 2
7. Each tube should have 10 µl of reagents at the bottom. To check your pipetting technique, set the pipettor to 10 µl and carefully draw the solution into a new pipet tip. a. Is the tip barely filled? (large undermeasurement) or b. Does a small volume of fluid remain in the tube? (overmeasurement) c. Is there a small air space left in the tip? (undermeasurement). Measure the actual volume by rotating the volume adjustment knob downward until the fluid reaches the tip, and read the indicated volume. 9. If one or more measurements is inaccurate, repeat the exercise and check results again. II. Practice with Medium- and Large- volume Micropipettors This exercise simulates the pipetting done in, for example, a bacterial transformation or plasmid DNA preparation for which 50 1000 µl pipettor is used. It is much easier to mismeasure with a large volume pipettor especially the P1000 type. It is also easy to draw fluid into the barrel of the pipettor accidentally. The plunger must be released slowly when pipetting. 1. Label two 1.5 ml microtubes E and F. 2. Use the matrix below as a checklist while adding solutions to each reaction tube. Use the techniques discussed above for pipetting. Add the volumes up to see if the tubes will be balanced for centrifuging when you have completed pipetting. Use the pipettor for which the range best matches the volume to be pipetted. Tube E F Soln. I 45 µl 30 µl Soln. II 155 µl 170 µl Soln. III 250 µl 375 µl Soln. IV 550 µl 425 µl 3. Pipet the volumes as before, using a fresh tip when appropriate (i.e., you may use the same tip for the first pipetting of the same reagent into clean tubes. After that, you must use a new, clean tip for each pipetting). 4. Briefly spin down all the reagents in your microcentrifuge. 5. Set your P1000 to the appropriate volume to measure the total you determined for each reaction tube. Slowly draw the fluid into the pipet tip. Ask the same questions (a, b, c) in exercise I. 7. above, and repeat measurements if any of your pipetting was inaccurate. 3
III. Practice Sterile Use of a 10 ml Serological Pipet The key to successful sterile technique at the bench is to work quickly and efficiently. Before beginning, clear off the lab bench and arrange tubes, pipets and culture media within easy reach. You may want to simulate the sequence of events of taking off and replacing caps without setting them down before you proceed. If you believe at any time that you may have compromised sterility, assume that you have and replace any plastic or media with reliably clean items. As a general rule of thumb, use sterile technique if live bacteria are needed at the end of the manipulation (e.g., general culturing, transformation). Sterility is not required if bacteria are being destroyed in the experiment (e.g., after an overnight culture, isolating bacteria for DNA isolation). Cleanliness, of course, is always desirable. When in doubt, use sterile technique. 1. Loosen caps on any tube or media container (sterile culture tube, container of sterile LB broth) so they can be easily lifted off. 2. Open the end of the pipet sleeve and attach the pipet bulb. Squeeze the bulb so that it is prepared to suck. Remove the sleeve from the pipet. 3. Lift the lid from the LB broth container (do not set down), insert the pipet and withdraw 5 ml of broth into the pipet. 4. Replace the LB broth container lid loosely. 5. Remove the lid from the culture tube (do not set down), and transfer the LB broth to the tube. Replace the lid, with the lid on the fingers. 6. Place the tube in a rack in the 37 C incubator to test its sterility (you may do this also with your LB broth container). The broth should be clear tomorrow if sterility was maintained. Check and record in your lab book. Place your tube (after one day of 37oC) into the refrigerator for later use. 4
Basic Techniques Bacterial Culture I. Isolation of Individual Colonies by Streaking Good bacteriological technique dictates that any culture is begun with a single well- isolated bacterial colony, which represents a clone the progeny of a single bacterium. This way one is reasonably certain that the culture grown contains a single, isogenic entity (all the same genotype), and not more than one kind of bacteria or other microbe. This exercise teaches you how to make a bacterial culture plate that has such isolated bacterial colonies. This exercise also demonstrates the phenomenon of antibiotic resistance by comparing the growth of two different strains of bacteria one with an ampicillin resistance gene on a plasmid, and one without on culture plates with or without the antibiotic ampicillin. Cultures and Bacterial Plates DH5a streak plate 2 LB agar plates (100 mm) pet28a wgmdh in DH5a (DE3) plate 2 LB agar - Ampicillin (or Carb*) plates & 2 LB agar Kan *Carbenicillin ( Carb ) is an equivalent antibiotic, but more long- lasting (and expensive). In general, an unmarked plate (for this lab) is considered to contain LB agar without antibiotic. Plates with antibiotic (or other components) are typically striped on the side with a color code indicating additions to the plate. It is important that the media portion of the plate is marked and not only the lid. Otherwise you should mark the plate on the bottom. Ideally culture plates are removed from the refrigerator where they are stored long- term and allowed to warm to room temperature before use (incubating at 37 C). Too rapid warming (transfer directly from 4 C to 37 C) can cause bubbles to appear in or under the plate making it difficult to see colonies on the plate. If the plate is still cool, condensation may need to be wiped off immediately prior to writing with a marker. Note that plates should be stored with the lid down/media side up (unless wet) this is bacteriologically right side up this way any airborne contaminants are more likely to land on the lid than on the media in the plate. 1. Use a marker to label the bottom of each agar plate (the portion with the LB agar medium) with the name of the bacterial strain, your initials and date, and (as needed) the type of bacterial plate (LB or LB/Amp). See the illustration below ( Anatomy of a Bacterial Streak Plate ). Write small and near the edges of a plate. Streaking matrix: You will streak a total of 6 plates (2 LB, 2 LB/Amp, 2 LB/Kan) with two different bacteria (DH5a Amp sensitive, pet28a wgmdh /DH5a (DE3) Amp resistant). Each bacteria will be streaked on each plate. Indicate your expected results by writing in growth or no growth into the chart below: Culture plate type Bacterial Strain LB LB / Amp LB / Kan DH5a pet28a wgmdh /DH5a (DE3) 5
2. Light your Bunsen burner. 3. Hold the inoculating loop in the flame until it is red hot, tilted to sterilize the maximal part of the wire. Make sure the entire wire portion up to the handle has at least been passed through the flame. 4. Cool the loop in an unused part of the bacterial plate (under the writing is a good spot). Return the plate to its lid on the bench. 5. With the lid resting on the bench, lift a culture plate (media part with colonies) and scrape the sterile loop through several isolated colonies. Close the plate, and open your clean plate. Smear the bacteria in a single large streak back and forth at the top of the plate (near the loop cooling region see illustration). 6. Flame the loop again and cool. 7. Pass the loop ONCE through the initial smear region, and continue into a clean region of the plate. Slide the loop on the surface back and forth several times (actually 2 or 3 times more than the illustration indicates). Use about 1/3 1/4 of the plate area. Note that if you tilt the plate to reflect the ambient light just right you should be able to make out on the surface of the plate where you have previously streaked with your loop. 8. Flame the loop again, cool, and pass once through the end of the first streak, and again streak back and forth through about 1/3 1/4 of the plate area. 9. Repeat one more time to make a 3 rd streak. Figure: Anatomy of a Bacterial Streak Plate Notes: Each new streak begins in the final pass of the previous streak. (The number of back & forth spreading is actually much more than illustrated.) When labeling the plate, write small, near the edges, on the media- containing part of the plate (not the lid). Write the plate type if there is any potential for confusion. 10. Repeat this for the remaining 3 plates. Then place all your plates (lid down) in the 37 C incubator to grow overnight. Next Day (parafilm, scissors needed) 11. Remove plates from the incubator. Observe and record the growth on the plates (see chart below). If you do not have well isolated colonies for either of the plates we are keeping, repeat the streaking process and grow again. 6
Culture plate type Bacterial Strain LB LB / Amp LB / Kan DH5a pet28a wgmdh /DH5a (DE3) 12. Keep the appropriate plates for later use (hint: the correct plate with the plasmid in the correct antibiotic how will you know? Results!!!). Parafilm and place plates in the refrigerator. Discard the other plates in the autoclave bag. 7
II. Starting a Small Overnight Culture (Demonstrated during first lab period; perform evening before Plasmid DNA isolation lab) Cultures and Media bacterial streak plate LB broth, sterile (or previously prepared tube) antibiotic Supplies and Equipment bacterial loop Bunsen burner 37 C shaking incubator sterile culture tube, tube rack parafilm, scissors 1. Take your bacterial streak plate from the refrigerator and remove the parafilm. 2. Use your previously- prepared culture tube with 5 ml LB broth (assuming it is still sterile). Light the burner and flame the loop. Cool the loop in a clean part of the plate. 3. Pick most or all of one large isolated colony with a uniform appearance. (If you do not have a plate with any isolated colonies, arrange to use someone else s plate.) Without setting the cap down, place the colony into the broth, and agitate the loop to dislodge the bacteria into the LB broth. Re- flame the loop. 3. Replace the cap on the fingers to allow air to enter the tube. Do not close or the culture will grow poorly. 4. Place your culture tube into the rack in the 37 orbital incubator shaker (if it already shaking, it will stop when you open the lid). Close the lid and wait to see that the platform begins rotating again. Add your tube to the tube rack in the shaker at the bottom of the incubator. Ensure the shaking incubator is on. Check timer to ensure you didn t accidentally move it from a permanent on status to a limited hour on. 5. Take a few moments for proper cleanup: Return equipment to where you found it. Re- parafilm your bacterial plate and return to the refrigerator. Wash your hands before you leave. Note the time you started your overnight culture in your lab book. Work with your partners and the instructor to create a cell growth curve. Group One One colony diluted to a 5 ml tube. Group Two 0.5 ml of an overnight culture diluted into 50 ml tube In general, overnight cultures should be incubated closer to 16 hrs or less rather than 24 hrs or more (set up evening before, harvested the next morning). For example, a culture set up at 5 PM and harvested at 9 AM the next day would have grown for 16 hours. As the culture goes to 24 hrs and beyond, more death of bacterial cells occurs, compromising the quality of the culture for a number of purposes. If you cannot use the culture in the morning, it is best to remove it from the 37 C incubator and leave at room temperature or even refrigerate. 8
Why do shorter cultures? See the bacterial growth curve at right after the cells run out of nutrients and accumulate waste products, entering stationary phase in which no further cell division occurs, eventually cells begin to die. The actual course of the growth curve depends on the bacterial strain and the precise growth conditions. Figure modified from image found at http://pathmicro.med.sc.edu/fox/growth_c.jpg 9