Now that we have prepared DNA encoding your mutant inverse pericams, we would like to produce the proteins themselves. Last time you were here, you transformed competent bacteria (called XL1-Blue) with mutagenized DNA prepared from a template plasmid. Successfully transformed bacteria grew into colonies on amipicillin-containing plates, and yesterday your oh-so devoted teaching staff picked two colonies per mutant to grow in liquid culture. The XL1-Blue cell line, although it now carries the inverse pericam DNA, cannot produce the inverse pericam protein. Today you will extract DNA from the XL1-Blue cells, prepare it for analysis, and transform your IPC mutant plasmids into a new bacterial system that can produce the protein directly.
The bacterial expression vector we are using (pRSET) contains the bacteriophage T7 promoter. This promoter is active only in the presence of T7 RNA polymerase (T7RNAP), an enzyme that therefore must be expressed by the bacterial strain used to make the protein of interest. We will use the BL21(DE3)pLysS strain, which has the following genotype: F-, ompT hsdSB (rB- mB-) gal dcm (DE3) pLysS (CamR). In BL21(DE3), T7RNAP is associated with a lac construct, and its expression is under the control of the lacUV5 promoter. Constitutively expressed lac repressor (lacI gene) blocks expression from the lac promoter; thus, the polymerase will not be produced except in the presence of repressor-binding lactose or a small-molecule lactose analogue such as IPTG (isopropyl β-D-thiogalactoside). To reduce ‘leaky’ expression of the protein of interest (in our case, inverse pericam), the pLysS version of BL21(DE3) contains T7 lysozyme, which inhibits basal transcription of T7RNAP. This gene is retained by chloramphenicol selection, while the pRSET plasmid itself (and thus inverse pericam) is retained by ampicillin selection - as you learned last time.
To isolate the inverse-pericam-containing pRSET plasmid from the overnight cultures, you will perform a miniprep. Because you are doing so few isolations compared to those done during Module 1, you will use a home-brew kit instead of the commercially available one. The major principle by which plasmid DNA is separated from everything else -- use of base and SDS, followed by acid and centrifugation -- remains the same. However, no silica column purification is done. One difference between the two approaches, then, is that the home-brew kit isolates a lot of RNA in addition to the plasmid DNA.
Production of mutant IPC protein.
The mutant DNA and protein are indicated by a green colour. Blue arrows/text indicate steps performed during class time; black arrows indicate steps performed by the teaching staff.
Once you have the plasmid DNA isolated, you can prepare it for sequencing and gel analysis, as well as use it immediately for transformation. In order to transform BL21(DE3) cells with your mutant IPC plasmids, you will first have to make the cells competent, i.e., able to efficiently take up foreign DNA. With the XL1-Blue strain, we used commercially available competent cells that did not need further treatment prior to DNA addition. Today, you will make chemically competent cells yourself using calcium chloride, then incubate them with plasmid DNA and heat shock them as before prior to plating. Shortly before you return from spring break, the teaching staff will pick colonies and set up liquid overnight cultures from your transformed cells. Next time, you will add IPTG to these liquid cultures to induce expression of your mutant proteins, which you will then isolate and characterize. Much of this process is summarized in the figure above.
Part 1: Prepare competent BL21(DE3) cells
- Pick up one 5 mL tube of BL21(DE3) cells. These cells should be in or near the early- or mid-log phase of growth, which is indicated by an OD600 value of 0.4-0.8.
- Measure the OD600 value of a 1:10 dilution of your cells (use a total volume of 650-700 μL). If the cells are not yet dense enough, return them to the rotary shaker in the incubator. Remember to balance with another tube! As a rule, your cells should double every 20-30 min.
Aspirate the supernatant, as shown, removing as few cells as possible
- Once your cells have reached the appropriate growth phase, pour them into 3 eppendorf tubes each containing ~ 1.5 mL culture volume. Spin for 1 min at max speed (~16,000 rcf/13,000 rpm), aspirate the supernatants as shown, and resuspend in an equal volume of ice-cold calcium chloride (100 mM). Note: you can balance these tubes in the centrifuge with three-way symmetry.
- You may find it easiest to resuspend the cells in a small volume first (say, 200 μL), then add the remaining volume of CaCl2 (e.g., in two steps of 650 μL) and invert the tubes to mix.
- Spin again for 1 min. The resultant pellets should occur as streaks down the side of the eppendorf tube, so be very careful not to disturb the cells when aspirating.
- This time, resuspend each pellet in 100 μL of CaCl2, then pool the cells together in one tube.
- Alternatively, resuspend the first pellet in 300 μL, then use this cell solution to resuspend the next pellet, and the next.
- Incubate on ice for 1 hour. You can work on parts 2, 4, and 5 of today's protocols now, as well as assemble the materials for part 3. Among other things, be sure to label four eppendorfs and pre-chill them on ice. The labels should indicate a (-) no DNA control, a (+) D24H/E67K/T79P/M124S transformation control (with minipreps prepared and vetted by the teaching staff), and your two mutant candidate transformations (X#Z -1 and -2).
- Pick up your two candidates cultures, growing in the test tubes labeled with your team colour. Label two eppendorf tubes to reflect your mutations and candidates (X#Z-1, X#Z-2).
- Vortex the bacteria and pour ~1.5 mL of each candidate into the appropriate eppendorf tube.
- Balance the tubes in the microfuge, spin for one minute, and resuspend the cells in 100 μL of Solution I, changing tips between samples.
- Note that the word "resuspend" implies aspiration beforehand.
- Prepare Solution II by mixing 250 μL of 2% SDS with 250 μL of 0.4M NaOH in an eppendorf tube. Add 200 μL of Solution II to each sample and invert the tubes five or six times to mix. The samples should become less opaque (almost clear) but don't worry if you don't see a big change.
- Place the tubes on ice for five minutes.
- Add 150 μL of Solution III to each sample and immediately vortex the tubes for 10 seconds at the highest setting. White clumps should appear in the solution.
- Spin for 4 minutes to separate the plasmid DNA from cell debris.
- While the tubes are spinning, label another set of eppendorf tubes with the candidate names and your team color.
- A white pellet should be visible when you retrieve your tubes. Transfer 400 μL of each supernatant to the appropriate clean eppendorf tube. It's OK to leave some of the supernatant behind; avoid transferring any of the white pellet.
- Add 1000 μL of room temperature 100% ethanol to each supernatant. The tubes will be quite full, so cap tightly before inverting at least five times to thoroughly mix the contents.
- Microfuge the samples for 2 minutes. It is now important to keep your tubes in a known orientation within the microfuge since the pellets from this spin may be barely visible.
- An orientation aligned with the tab of the cap may be easiest to remember.
- Remove the supernatants using your P1000, taking care not to disturb the pellet of plasmid DNA that is at the bottom of the tube, and put them in a 15 mL conical waste collection tube. Remove as much of the supernatant as possible, but you do not need to remove every drop since you will be washing the pellet in the next step.
- Add 500 μL of 70% ethanol to each pellet. You do not need to fully resuspend the pellet, but you might invert or flick the tube a few times. Spin the samples one minute, orienting the tubes in the microfuge so you will know where to find the pellet.
- Immediately remove the supernatant with your P1000, making sure to keep the tip on the side of the tube that doesn't have your pellet. Remove as much liquid as possible, using your P200 set to 100 μL to remove the last few droplets and/or to streak them up the side of the tube to promote evaporation.
- To completely dry the pellets, place your rack in the hood with the tube caps open for ~ 10 minutes. When the pellets are completely dry, add 50 μL of sterile water to each sample and vortex for 2 X 30 seconds (with a quick-spin in between and afterward) to completely dissolve the pellets. Store the DNA on ice, and do NOT throw it away after completing Part 3. (The teaching faculty will save the minipreps for potential future use.)
Part 3: Transform BL21(DE3) with mutant DNA
- Prewarm and dry 4 LB+Amp/Cam plates by placing them in the 37°C incubator, media side up with the lids ajar. You will perform one transformation for each of your four samples.
- When your competent cells are ready, aliquot 70 μL of cells per pre-chilled eppendorf.
- Add 2 μL of the appropriate DNA to each tube. Remember, you are testing plasmid DNA that was prepared from two different colonies for your X#Z mutant, along with DNA from a colony that is already known to be M124S/E67K/T79P. You will also perform a no DNA control.
- Flick to mix the contents and leave the tubes on ice for at least 5 minutes.
- Heat shock the cells on the 42°C heat block for 90 seconds exactly and then put on ice for two minutes.
- Move the samples to a rack on your bench, add 0.5 ml of LB media to each one, and invert each tube to mix.
- Incubate the tubes in the 37°C incubator for at least 30 minutes. This gives the antibiotic-resistance genes some time be expressed in the transformed bacterial cells.
- While you are waiting, label 3 large glass test tubes with your team color and sample names. (You can also finish part 5 of the protocol if you have not yet done so.)
- Normally we would have you combine LB culture broth with both ampicillin and chloramphenical, and then aliquot 2.5 mL of this mixture per tube. Due to the long break before your colonies are actually picked, we will do this part for you closer to M2D5.
- Also prepare 4 eppendorf tubes containing 180 μL of LB each. You will use these to dilute your transformed cells 1:10 when you retrieve them from the incubator.
- If you label these tubes with stickers rather than directly on the cap, you can then transfer each sticker to the appropriate plate as you go, saving one labeling step.
- Note that we are reducing the cell concentration because miniprep DNA is much more concentrated than the DNA resulting from mutagenesis; it also does not require repair, further increasing the transformation efficiency.
- Plate 200 μl of each (1:10 diluted) transformation mix on an LB+Amp/Cam plate.
- Safety reminder: After dipping the glass spreader in the ethanol jar, then pass it through the flame of the alcohol burner just long enough to ignite the ethanol. After letting the ethanol burn off, the spreader may still be very hot, and it is advisable to tap it gently on a portion of the agar plate without cells in order to equilibrate it with the agar.
- Once the plates are done, wrap them with colored tape and incubate them in the 37°C incubator overnight. One of the teaching faculty will remove them from the incubator and set up liquid cultures for you to use next time.
Part 4: Count mutant colonies
When you have a spare moment today, count the colonies that arose on your transformed XL1-Blue plate, as well as on your positive and negative control plates. Does the negative control have any colonies? How does your mutation efficiency compare to that of the positive control? Please put your colony counts on today's Talk page.
Part 5: Prepare DNA for Evaluation
You will perform diagnostic digests on the following samples: the inverse pericam parent plasmid (pRSET-IPC), a known mutant pRSET-M124S or -T79P or -E67K or -D24H, and two candidates for your X#Z mutation. "Digest 1" (D1) will be used to show that the reference mutant DNA contains the correct mutation, and "Digest 2" (D2) will be used to test whether your X#Z candidates do. Thus, you will need enough D1 mixture for two reactions (IPC and M124S/T79P/E67K/D24H), and enough D2 mixture for three reactions (IPC and the two X#Z candidates). To avoid pipetting very small volumes of enzymes, and in order to have at least a little extra of each reaction (so you don't run out due to pipetting error), make enough of each digest for four reactions.
The table below is for one reaction and assumes that each digest will consist of a single enzyme. Note that enzyme stock concentrations can be found on the NEB product page for that enzyme.
|10X NEB buffer
||2.5 μL of buffer#3.1
||2.5 μL of buffer#_____
||2.5 μL of buffer#_____
||2.5 U = 0.25 μL of PvuI
||2.5 U = __ μL of _____
||2.5 U = __ μL of _____
||For a total volume of 25 μL
- Prepare a reaction cocktail for each of the above reactions (digest 1 and digest 2) that includes water, buffer and enzyme. Prepare enough of each cocktail for 4 digests. Leave the cocktails on ice.
- Aliquot 4 μL of the appropriate plasmids into five well-labeled eppendorf tubes. The labels should include the plasmid name, the enzyme(s) to be added and your team color.
- Add 21 μL of the appropriate cocktail to each tube. Flick the tubes to mix the contents, touch-spin, then incubate the mixtures at 37°C for at least one hour.
- While your samples are digesting, you can return to Part 3 of the protocol.
- Before leaving lab today, please add 2.5 μL of loading dye to each of the digests you have assembled. You should also prepare undigested samples of parent IPC and each mutant candidate, containing 4 μL of plasmid, 21 μL of water, and loading dye. We will store the digests and the remaining DNA at –20°C.
As you learned during Module 1, sequencing reactions require a primer for initiation. Legible readout of the gene typically begins about 40-50 bp downstream of the primer site, and continues for ~1000 bp at most. Thus, multiple primers must be used to fully view genes > 1 Kbp in size. How many basepairs long is inverse pericam? (Check the ApE file linked in your previous homework.)
Luckily, we only care about the back end of IPC, i.e., the part containing calmodulin. The primer you will use today starts at base pair 823 of pRSET-IPC, a couple hundred base pairs upstream of CaM. As for the positive controls, everyone will be given the same sequencing data to analyze, because you are all working with DNA from the same four candidates.
The recommended composition of sequencing reactions is ~500 ng of plasmid DNA and 25 pmoles of sequencing primer in a final volume of 15 μL. The miniprep'd plasmid should have ~1 μg of nucleic acid/μL but that will be a mixture of RNA and DNA, so we will estimate the amount appropriate for our reactions.
For each reaction, combine the following reagents directly in the appropriate tube within the 8-PCR-tube strip, as noted in the table below:
- 2.5 μL of your plasmid DNA candidate
- 12.5 μL of the primer solution on the teaching bench, which contains
- 5 μL of sequencing primer at 5 *mu;M
- 7.5 μL sterile water
The side of each tube is numerically labeled and you should use only the two tubes assigned to your group. The teaching faculty will turn in the strips at the Genewiz company drop-off box for sequencing.
For next time
1. BL21(DE3) E. coli are often used for protein expression. In contrast, XL1-Blue E. coli are ‘workhorse’ cells useful for plasmid propagation. One gene modification in XL1-Blue is the hsdR mutation. What are the two other modified genes in XL1-Blue that make them ideal for the task of propagating a desired DNA? Briefly explain why each is important (1-2 sentences). It may help you to refer to the cell manual: (pdf download), but be sure to answer the question in your own words. Note that this question is about propagation, not screening.
2. The major assessment for this module will be a research article describing your protein design work. For this assignment, you will write start outlining and drafting the introduction to your report. Recall that the introduction provides a framework for the story you are about to tell, with respect to both background and motivation. You are encouraged to revisit the scientific writing guidelines before you begin.
- What might the big picture section consist of? Will you focus on engineering sensors in general, the utility of sensing calcium specifically, or ... what?
- What background would the reader like to have as you zoom in?
- When you define your specific study, be sure to share your hypotheses about both your unique mutant and one of the reference mutants.
At a minimum, you should hand in some high-level ideas and supporting notes for the first two sections, and write a draft of the third section in complete sentences.
3. Prepare a schematic that depicts your mutagenesis strategy and write a short caption for it. You might show proposed changes at both the nucleotide and amino acid level for D24H/E67K/T79P/M124S and X#Z. If you have switched mutants, be sure to write about the one that you are going forward with.
4. Please download the midsemester evaluation form found here (DOC download). Complete the questionnaire and then print it out without including your name to turn in. If there is something you'd like to see done differently for the rest of the course, this is your chance to lobby for that change. Similarly, if there is something you think the class has to keep doing, let us know that too. These will be collected in a separate folder from your FNTs, to preserve anonymity.
- 100 mM CaCl2, sterile
- LB (Luria-Bertani broth)
- 1% Tryptone
- 0.5% Yeast Extract
- 1% NaCl
- autoclaved for sterility
- Ampicillin: 100 mg/mL, aqueous, sterile-filtered
- Chloramphenicol: 34 mg/mL in ethanol
- LB+AMP+CAM plates
- LB with 2% agar and 100 μg/ml Ampicillin and 34 μg/ml Chloramphenicol
- Solution I
- 25 mM Tris pH8
- 10 mM EDTA pH8
- 5 mM Glucose
- Solution II
- Solution III
- Parental plasmid (pRSET-IPC)
- Mutant plasmids (pRSET-M124S or -T79P or -E67K, and your two minipreps)
- NEB buffers 1-4
- NEB enzymes
DNA Sequencing materials
- Sequencing primer "IPC-seq-f-823" dilute to 5 pmol/μL in water (original stock 100 pmol/μL)