Introductory Lab Procedures

Preliminary Lab activities

The main objectives of these protocols are for you to become familiar with measuring small volumes of liquids,  making buffers and other solutions, and becoming familiar with some of the equipment in the lab. During the first two weeks of lab, each group will perform all of these protocols. Look in Appendix 1 for more detailed information on equipment and measuring liquids.

Protocol I: Working with very small volumes of liquid (using micropipettes)

Although you will be working with your team, each student should individually do each of these micropipetting exercises.

Throughout the entire semester, we will often be pipetting very small volumes (< 1 ml) of clear, colorless liquids that contain critical reagents. Careful and accurate pipetting will determine the success of those procedures, so it is useful to practice working with very small volumes of “colored”, visible liquid before it’s time to pipette an important (and expensive!) reagent.

The instructor will demonstrate pipetting with the micropipettes, and either the instructor or TAs will be available for assistance as needed. But as with anything…practice is essential.

Each team should obtain an aliquot of blue dye, yellow dye, and red dye (already distributed at the stations). Fill the ‘larger’ beaker with dH2O from the carboy or the MilliQ unit.

For each exercise below, make sure you are using the appropriate pipette with the appropriate tip!

IMG_3367
Micropipets

a.Into each of five separate microfuge tubes*, pipette 100 µl of water. Using your p2 pipetman, pipette 2.0 µl, 1.5 µl, 1.0 µl, and 0.5 µl of blue dye into the individual water samples. Mix well (vortex the closed tube) and observe the differences in color intensity.

b. Label an eppendorf tube “A”; Make a solution that contains 1.0 ml H2O, 7 µl blue dye, plus 23 µl yellow dye. Cap the tube, vortex, and place it in the rack; note color of resulting solution.

c. Label an eppendorf tube “B”; Make a solution that contains 1.0 ml H2O, 20 µl red dye, plus 35 µl yellow dye. Cap the tube, vortex, and place it in the rack; note color of resulting solution.

The following 2 exercises begin with a fixed volume of liquid in a tube; then after successive removal of the specified volumes (below) with the appropriate micropipettes, there should be some liquid remaining in the original tube. This will help you to determine if your technique is correct. (If you ‘run out’ of liquid before the end of the exercise, you will know that you pipetted incorrectly.)

d. Each student: pipette 1000 ul of H2O into a clean microfuge tube. (Pulse spin the tube to make sure that all the liquid is at the bottom!) Then, from this tube, pipette out the following volumes (and simply discard in waste beaker): 486 µl, 258 µl, 125 µl, 98 µl, 16.5 µl, and 1.5 µl. Upon finishing, you should still have a little liquid left over in the original tube (~20 µl). If you have ‘run out’, please try again!

e. (The team can work together on this one): obtain a tube with 300 µl blue dye (prepared by instructor; already distributed to the stations). Pulse spin the tube to bring down any condensation to the bottom. Then, each team member taking turns, pipette out (and discard) 118 µl of the dye; then pipette out 77 µl, 58 µl, 17 µl, 7 µl, 2 µl, and 1 µl. At the end of this exercise, you should still have ~20 ul of blue dye left in the original tube. Do you? Familiarize yourself with the appearance of those volumes of liquid in the pipette tips.

f.  In addition to the dyes, each team will also have an aliquot of isopropanol; each person should simply practice pipetting up and down a few times into the same tube (with any size micropipette) as needed, to get a feel for pipetting an alcoholic solution. You may notice that the reduced surface tension of the alcohol (relative to water and other aqueous buffers) makes it more prone to ‘drip’ from the pipette tip, and therefore you must work more quickly when pipetting from one vessel to another.

g. A bottle of detergent (NP-40) will be available at the “TA” station in front of the lab; using a serological pipette (any size), practice pipetting the concentrated detergent up and down to get a feel for pipetting very viscous solutions.

Protocol 2: Buffer preparation

This exercise takes you through the typical steps of preparing a very commonly used buffer in biological research laboratories.

Each group will prepare 100 ml of a Tris buffer solution [“Trizma Base”1, NH2C(CH2OH)3, molecular weight 121.1 g/M]. Choose a molarity between 20 mM and 100 mM. Choose a pH between 7.5 and 8.8 (the optimum buffering range for Tris buffers is between ~7.4 and 9).

  1. Calculate the appropriate amount of Trizma base (Tris) to weigh out, using its molecular weight and the final volume (100 ml), and final concentration (in mM) of the solution you are preparing. Weigh out the Tris on the balance. Be sure to tare (zero) the balance with the weighing paper/boat.
  2. Place a stir-bar into a 250 ml beaker, then add ~80 ml of dH2O (Milli-Q water or water from carboy). Begin stirring on the stir-plate between the balances and pH meter. Then, pour the weighed Tris into the beaker and wait for it to dissolve. (While you’re waiting, the pH meter can be calibrated.)
  3. Observe the starting pH of the solution using a pH meter that has been calibrated with standard pH buffers. Let the pH electrode rest in the beaker, safely away from the rotating stir-bar. Then, as the buffer stirs, add acid (1:10 HCL) or base (0.1 M NaOH) dropwise, as necessary, to achieve the desired pH.
  4. Once the correct pH has been attained, pour the buffer (minus the stir- bar) into a 100 ml graduated cylinder for accurate measurement of final volume. Add enough dH2O to reach 100 ml (use the dH2O carboy and just ‘eyeball’ it). Cover with parafilm and invert a couple of times to mix completely.
  1. Using the labeling tape and sharpie markers, label the covered cylinder with the date, name of the buffer (“Tris-Cl”), concentration (molarity), the pH, and your group’s name (for example, “25 mM Tris-Cl, pH 7.6, Team Awesome”, etc).

Protocol 3: Protein serial dilution

For this protocol, each team will obtain a tube containing bovine serum albumin (BSA) at 1 mg/ml (in dH2O).

The following protocol illustrates the power of spectrophotometry as a tool with which to measure the concentrations of macromolecules that absorb light at specific wavelengths. (It is also a test of your accuracy in pipetting.) Both proteins and nucleic acids in solution will absorb light at characteristic wavelengths, and within a certain concentration range, there will be a direct linear relationship between the concentration of the substance and its absorbance at the appropriate wavelength.

1. Label 6 *12 X 75 mm tubes with numbers “1” through “6”.

2.  Pipet 750 µl dH20 into each tube.

3.  Pipet 750 µl of the protein solution into the first tube (“1”); mix well (vortex carefully).

4. Transfer 750 µl of tube #1 contents to tube #2. Mix well as above.

Repeat the 1:2 dilutions with remaining tubes.

5. When you have completed the dilutions, go to the uv/vis spectrophotometer at back of 406 (NINJA will assist).

6. “Blank” the machine by inserting the Quartz cuvette filled with 600 ul water (same solvent in which the protein is dissolved).

7.  Read your samples at an absorbance of 280 nm. Write down the readings. Note: if you begin with your most dilute sample and work backwards (towards the most concentrated), you won’t need to rinse the cuvette between readings.

Questions:

  1. What concentration of protein is in each tube? (ug/ml is the most convenient unit)
  2. Plot your absorbance (A280) vs. protein concentration; are the absorbance readings linear with respect to the protein concentrations?

Protocol 4 (week 2): LB-Antibiotic (Ampicillin) agar plates for bacteria

This procedures provides each student hands-on experience in pouring of nutrient agar plates for bacterial culture; you are also preparing plates that you will use in future protocols.

The following steps will be completed prior to lab:

  1. Place eight pre-measured LB-Agar pellets in a 2L bottle or flask. b. Add 1L of Milli-Q water.
  2. Place cap on loosely, apply autoclave tape.
  3. On the liquid (slow exhaust) cycle, autoclave for 20 min.
  4. Remove promptly from the autoclave so that the liquid doesn’t caramelize. f. Swirl carefully to evenly distribute the pellet material.

(The LB-agar will be stored in a 55°C bath to prevent solidification)

  1. During lab: when the liquid has cooled to the point where the flask can comfortably be handled with gloves, instructors will add 1 ml of 100 mg/ml Ampicillin (for 100 µg/ml final).

Each student then does the following steps (after a demonstration):

  1. Poor the liquid into ~1-3 petri dishes, one at a time, as demonstrated by instructors/TAs (we want everyone to get a chance to pour at least a few plates before the solution runs out).
  2. If bubbles form, flame the bubbles.
  3. Cover the plates and allow them to solidify and dry at room temperature

(~1-2 days).

  1. Once the plates have hardened and there is no condensation on the lid, we will return them to the plastic sleeves and store at 4°C, inverted, for up to 2 months.

Protocol 5: Cell culture (Observation only, as time allows)

Look at U2OS human osteosarcoma cells (and other cell lines as available) under the inverted microscope at various magnifications; ideally we will have 1 or 2 plates of cells at different densities: log phase (subconfluent, meaning cells have room to grow), and confluent (cells are all in contact on all sides, no more space to proliferate). Note any differences between the morphology of individual cells within a dish, which could be due to heterogeneity of cell types and/or being in different stages of the cell cycle. Note also the differences in morphology of cells that are in log phase of growth and those that are confluent (or overgrown).

Which appear to be “more healthy”?

Protocol 6: Understanding Dilutions- make Tris-EDTA (TE) buffer from stock solutions.

TE buffer = 10 mM Tris-Cl, 1 mM EDTA, pH 8

TE buffer is commonly used to dilute and store DNA preparations at -200C; DNA can be diluted in water for short-term storage, but use of this alkaline TE buffer protects the DNA from degradation and is preferred for long-term storage.

Each team will need its own supply of TE buffer for use throughout the semester. In the event of contamination of the buffer (with DNA, environmental microorganisms, etc.), maintaining your own buffer eliminates the risk of the contamination spreading to other groups (which has happened in the past, when sharing from a common stock!).

Using your understanding of dilutions from Appendix-1 and from the lab discussions, prepare 10 ml of your own TE buffer using the following stocks (which will be provided to you): (no pH adjustment will be necessary)

  • 1M Tris-Cl, pH 8
  • 0.5M EDTA, pH 8
  • use MilliQ dH2O to bring up the final volume to 10 ml; ask instructors if you are unsure of the calculations.2

You can use a 15-ml blue-top tube for the initial solution. Then, to sterilize, use the provided 10-ml syringe and syringe filter to filter the solution into a new, labeled 15 ml tube (this will be demonstrated). Label this tube with your team name (in addition to “TE buffer, pH 8”) and leave it at your stations.

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