As you know, you take your Final Lab Practical next week (4/25/17-4/28/17). Below is a list of things compiled by our teaching staff that you need to review, i.e., need to know for our Final Lab Practical. Please be aware that you do not get to use your lab manual when you take the Final Lab Practical, but you can use your calculator.
BIOL 135 Final Lab Practical Review
**GENERAL NOTE: The setup of the lab practical consists of various stations that test the hands-on skills you were taught throughout the semester. You correctly demonstrate these skills in order to do well on the practical final. If you are too slow at performing any of the assessments, it will be assumed that you did not learn the material, and thus will have points deducted and you will be moved to the next station.
- Know “Basic Lab and Safety Procedures”
- How to discard specific lab wastes
- Specific examples to know:
- 10 mL of acid: neutralize with water and pour down the drain with lots of water
- Agar plate containing E.coli: tape closed and put in orange biohazard bag in biohazard bin
- Test tube containing B.subtilus: goes to media prep to be autoclaved
- Kimwipe used on pH meter: trash can
- **TIP: a question may be asked that seems general, but BEWARE. If a question is general, such as how to dispose of a glove or Kimwipe that has something biological on them (cells, bacteria, etc.), then they need to be disposed of in biohazard bags, NOT in the trash. Be aware of what substance is on an object you are disposing because it makes a difference as to whether or not it goes in the trash, in a special trash container (like for cobalt chloride), or into the biohazard container (or the sharps container, or the broken glass box).
- Using a beamed balance
- Maximum capacity of our beamed balances ~ 610 g
- Zeroing (taring) and using an electronic balance
- Maximum capacity electronic balances ≤ 300.000 g
- Know how to calibrate electronic balance using 200 g calibration weight
- Measuring with a Vernier caliper
- Measuring objects to the nearest 0.1 mm with a Vernier caliper. Helpful hint: first measure an object with a ruler to get an estimate of the linear dimension, and then measure with the Vernier Caliper for greater precision
- Know the steps/stages/phases of mitosis and meiosis
- Know the two alignments at metaphase I of meiosis
- Be able to perform the various stages of mitosis and meiosis with pipe cleaners and beads, as done in lab
- How to do serial dilutions
- **TIP: Be sure to use a clean graduated pipet/pipet tip every time you transfer a liquid to avoid contamination/carry-over, and thus inaccurate dilutions.
- Using a Pipetman along with checking the accuracy of a Pipetman.
- With the plunger at the 1st stop, immerse the tip into the liquid and allow plunger to slowly return to the original position. When dispensing, push the plunger to the 1st stop before fulling pushing the plunger to the 2nd stop to dispense all fluid from the tip.
- **TIP: Pay close attention to the different stop positions of the Pipetman, be sure to NOT let the plunger snap back to the original position (slowly release it by releasing thumb pressure), and make sure you know which Pipetman to use/how to set it to the correct volume/how to read the volume of each model of Pipetman. Not following these important points will result in point deductions.
- Check the accuracy of a Pipetman: depending on the capacity of a Pipetman, tare a weigh boat on the electronic balance, set volume of Pipetman, load volume of water into tip, dispense water into weigh boat
- Use the close approximation for the density of water that is 1 mL = 1000 μL = 1 g. Therefore, 100 μL = 0.1 g, 200 μL = 0.2 g, 1000 μL = 1 g. Depending on the model of Pipetman, one of these volumes will be good to check the Pipetman’s accuracy.
- If the weight of volume delivered by a Pipetman is > 1% of what it should be, then that Pipetman is not accurate and you thus need to adjust volume to be delivered to account for this inaccuracy, or the Pipetman needs to be serviced.
LABS 5, 6
- Calibrating a pH meter, and how to measure the pH of a solution
- Before measuring the pH of a solution using a pH meter, you should identify whether the solution is acidic or basic by using universal pH paper. DO NOT dip the strip of pH paper into a solution, use a clean Pasteur pipet to put a drop on the paper.
- If you are preparing to measure an acid, use the pH 7 and pH 4 buffers to calibrate. For a base, use the pH 7 and pH 10 buffers.
- First check to see that the stopper plug is slid open. Clean off the pH probe with electrode (it should be sitting in pH 4 buffer solution) with DI water in a wash bottle, allowing the rinse water to fall into a waste beaker. Gently wipe the pH probe with a Kimwipe. Place the pH probe into pH 7 buffer, and allow it to stabilize. Place the pH probe into pH 7 buffer, and allow it to stabilize. Press the STANDARDIZE button.Remove pH probe from the buffer solution and rinse the pH probe with DI water, and dry with a Kimwipe. Place the pH probe in the second buffer solution, either pH 4 or pH 10 buffer. Again, allow to stabilize then press the STANDARDIZE button. Rinse the pH probe with DI water, and dry with a Kimwipe. Place the pH probe in the solution and measure its pH. When finished, rinse the pH probe with DI water, and dry with a Kimwipe, the put the probe in pH 4 buffer for storage.
- Buffer solution color-code: pH 4 = red, pH 7 = yellow, pH 10 = blue
LABS 7, 8, 9
- Adjusting a spectrophotometer and measuring the absorbance of a solution.
- Be sure spectrophotometer has been on (warmed up) for 15 minutes. Set the wavelength with the Wavelength Knob. Set the Mode (Abs. or Trans.) with the MODE Switch. Insert reference or blank cuvette tube, and close sample compartment lid. Press the red 100%T/0A Button. Now ready to read the Absorbance or Transmittance of a substance in solution.
- Remember to always wipe off the outside of a cuvette tube with a Kimwipe before inserting tube into sample compartment; use forceps to remove a cuvette tube from the sample compartment; and whenever you change the wavelength of the spectrophotometer, you must “reset” it with the blank.
- **TIP: Remember to vortex solutions before reading their Absorbance or Transmittance to be sure the solution is “homogenous.” Remember cobalt chloride (CoCl2) is toxic, so wear gloves when handling it. Remember, wear gloves when doing the Bradford Assay to prevent proteins on your hands contaminating your solutions.
- Here is an overview of the basic guidelines for sterile technique: wash your hands with soap and water before you start and when you finish, wipe down your work area with 70% ethanol before you start and when you finish, set up your work area to minimize contamination, use sterile glassware, wear gloves if instructed, hold the opening of flasks, test tubes, or bottles at a 45° angle to minimize airborne contamination, and place a cap on a clean surface on the lab bench.
- Sterile technique using graduated pipets, and when pouring solutions
- **TIPS: Never allow the tip of a graduated pipet touch you or a surface. Remember to pass the bottom third of the pipet through the flame for 1-3 seconds BEFORE and AFTER loading/dispensing liquid microbial culture. All metal containers containing graduated pipets must be flamed before being uncapped and exposed to the air to reduce contamination.
- Streaking an agar plate using sterile technique, and single colony isolation
- **TIPS: Review the diagram in your manual for how to streak for single colony isolation. Be sure to fully flame an inoculation loop by holding it at the top of the blue flame cone and waiting until it turns bright red; this must be done for 2/3 of the tungsten wire with look, not just the loop end. After flaming a loop, be sure to allow the loop to cool (if picking up bacteria from a liquid culture) or cool it in “side agar” (if picking up from a solid culture) to avoid killing (frying) the bacteria. Always flame the inoculation loop BEFORE picking up bacteria and AFTER spreading it on the agar. Just lift the lid of an agar plate a bit (about 1-2 inches) when picking up bacteria or streaking.
- Inoculating a liquid culture
- First, flame the inoculation loop. Cool it by using a “side agar cooling spot,” before scooping up solid bacteria from an agar plate. For the broth, uncap the test tube and flame its opening. Be careful not to touch the inner sides of the test tube, then lower the cooled inoculation loop into the broth and give a few quick motions to “stir” the culture. Remove the loop. Flame the test tube opening and recap. Finish by flaming the inoculation loop.
- Light microscope
- Know all the parts to a light microscope
- How to focus on a specimen. Begin by observing a specimen at the lowest power objective (4x) and increase as necessary
- How to maximize resolution using Koehler Illumination
- When to and not to use immersion oil. Remember, immersion oil is ONLY used with the 100x objective.
- How to clean the lenses of a light microscope. Lens paper is the only material used to clean a microscope eyepiece and objective lenses, NEVER Kimwipes.
- Preparing bacterial smears using liquid and solid cultures
- To prepare a smear from an agar culture, put a very small drop of RO water on the center of the slide. Using sterile technique, remove a small amount of bacteria from an agar culture using an inoculation loop, then mix and spread the bacteria in the water droplet on the microscope slide. Remember with an agar culture, it takes a very little “scoop” of bacteria to give a usable smear.
- To prepare a smear from a liquid culture of bacteria, do not place a drop of RO water on the slide, instead use sterile technique to remove a drop of liquid culture with an inoculation loop, then use the loop with the drop of culture to smear the bacteria onto the cleaned slide.
- Allow the smear to air-dry completely. Once dry, quickly pass the slide through the flame three quick times to affix the bacteria to the slide. Once cooled, the bacterial smear is ready to stain.
- Staining bacterial smears using methylene blue and gram staining techniques.
- Methylene blue stain: apply methylene blue stain on the smear on a slide, a drop at a time until the smear is covered with stain. Allow the stain to sit on the smear for 60 seconds. If it begins to dry, add more stain. After 60 seconds gently wash off the stain with RO water. Remove excess water from the slide by using Kimwipes or paper towel. Use spray fixative on the moist stained smears and let dry for a couple minutes before examining with a microscope.
- Gram stain: Flood a slide with crystal violet for 60 seconds. Pour off the stain and gently wash with RO water. Next, flood the slide with iodine solution and leave for 30 seconds to 1 minutes. Poor off and wash. While holding the slide at an angle, apply 95% ethanol one drop at a time until the violet color no longer appears in the runoff. Quickly rinse off the alcohol with water and blot dry. Counter stain by flooding the slide with safranine for 60 seconds. Pour off and wash. Blot dry as previously described and use spray fixative on moist stained smear before examining bacteria.
- Direct methods used to measure bacterial growth:
- Viable plate counts
- Spread plate method: a volume of an appropriately diluted bacterial culture is spread over the surface of an agar plate using a sterile glass cell spreader (aka hockey stick). Allow the bacterial culture to soak into the agar for ~15 minutes before inverting the agar plate and incubating.
- **TIPS: Counts must be between 30 and 300 for both forms of direct methods. When sterilizing a glass cell spreader, first soak the spreader in ethanol then flame the spreader and allow all ethanol to burn off before using to spread the liquid culture evenly on the agar.
- Indirect methods used to measure bacterial growth:
- Turbidity measurements using Spec20
- Calibrate eyepiece graticule using a stage micrometer.
- To begin, focus on the scale on the stage micrometer. Next, align the lines of the eyepiece graticule with those of the stage micrometer so that they are parallel, by adjusting the stage micrometer or rotating the eyepiece with the graticule. Align the zero lines of the stage micrometer and eyepiece graticule. Count how many lines on the stage micrometer fit precisely in a given number of lines of the eyepiece graticule. From this record the value that represents how many graticule lines represent a designated length (with units) when using 10x, 40x or 100x objective.
- For example:
- If you counted 39 eyepiece graticule lines per 0.1 mm of the stage micrometer, then the distance between each eyepiece graticule line is 100 μm / 39 = 2.6 μm
- Measure length of object using calibrated eyepiece graticule
- Using the value you found that represents how many graticule lines represents a designated length, measure the length of an object on a slide
- Examine and identify histological sections