Difference between revisions of "20.109(S09):Prepare expression system (Day4)"

Revision as of 02:47, 13 October 2008

Introduction

Now that we have prepared DNA encoding your mutant inverse pericams, we would like to actually produce the physical proteins. Last time you were here, you transformed competent bacteria (called XL1-Blue) with mutagenized DNA prepared from a template plasmid prepared. 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. Thus, 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.

From Invitrogen manual

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 (r_B^- m_B^-) gal dcm (DE3) pLysS (Cam^R). In BL21(DE3), T7RNAP is associated with a lac construct, and its expression is under the control of the lacUV5 promoter. Due to the action of the lac repressor (lacI gene), the polymerase will not be produced except in the presence of lactose or a small-molecule lactose analogue such as IPTG (isopropyl β-D-thiogalactoside). To further reduce ‘leaky’ expression of the protein of interest (in our case, inverse pericam), the pLysS version of BL21(DE3) contains T7 lysozyme, which also 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.

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.

To isolate the inverse-pericam-containing pRSET plasmid from the overnight cultures, you will perform what is commonly called a “mini-prep.” This term distinguishes the procedure from a “maxi-” or “large scale-prep” which involves a larger volume of cells and additional steps of purification. The overall goal of each “prep” is the same--to separate the plasmid DNA from the chromosomal DNA and cellular debris, allowing the plasmid DNA to be studied further. In the traditional mini-prep protocol, the media is removed from the cells by centrifugation. The cells are resuspended in “Solution I” which contains Tris to buffer the cells and EDTA to bind divalent cations in the lipid bilayer, thereby weakening the cell envelope. A solution of sodium hydroxide and SDS is then added. The base denatures the cell’s DNA, both chromosomal and plasmid, while the detergent dissolves the cellular proteins and lipids. The pH of the solution is returned to neutral by the acetic acid and potassium acetate in “Solution III.” At neutral pH the SDS precipitates from solution, carrying with it the dissolved proteins and lipids. In addition, the DNA strands renature at neutral pH. The chromosomal DNA, which is much longer than the plasmid DNA, renatures as a tangle that gets trapped in the SDS precipitate. The plasmid DNA renatures normally and stays in solution, effectively separating plasmid DNA from the chromosomal DNA and the proteins and lipids of the cell.

Once you have the plasmid DNA isolated, you can prepare it for sequencing and gel analysis, as well 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 using calcium chloride, then incubate them with plasmid DNA and heat shock them as before prior to plating. Tomorrow, 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 at left.

Protocols

Part 1: Prepare competent BL21(DE3) cells

1. Pick up two 3 mL tubes of BL21(DE3) cells. These cells should be in or close to the mid-log phase of growth, which is indicated by an OD₆₀₀ value of 04.-06.
2. Measure the OD₆₀₀ value of a 1:10 dilution of your cells (use 50 μL of cells per tube). If the cells are not yet dense enough, return them to the rotary shaker in the incubator. Remember to balance your tubes! As a rule, your cells should double every 20-30 min.
3. Once your cells have reached the appropriate growth phase, pour them into eppendorf tubes. Spin down 4 tubes of ~ 1.5 mL each for 1 min at max speed, aspirate the supernatants, and resuspend in an equal volume of ice-cold calcium chloride (100 mM).
4. 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.
5. This time, resuspend each pellet in 100 μL of CaCl₂, then pool the cells together in one tube.
6. Incubate on ice for 1 hour. (You might work on parts 3-5 of today's protocols now.)
7. Meanwhile, label five eppendorfs (for four transformations and a no DNA control) and pre-chill them on ice. (You can label your mutant tubes as X#Z, where # is the residue number you are modifying, X is the original amino acid, and Z is the mutant acid. The example shown on Day 1 would be Y64D.)

Part 2: DNA extraction (mini-prep)

1. Pick up your four candidates cultures, growing in the test tubes labeled with your team colour. Label four eppendorf tubes to reflect your mutations and candidates (X#Y-1, X#Y-2, Y64D-1, and Y64D-2).
2. Vortex the bacteria and pour ~1.5 mL of each candidate into the appropriate eppendorf tube. If you are nervous about pouring the liquid, you can use your P1000 to pipet 750 μL into each eppendorf twice. Either way, the eppendorf should be quite full when you try to close the cap. You can wear gloves to keep the bacteria from splashing your skin or you can wash your hands after closing all the caps.
3. Balance the tubes in the microfuge, and then spin them for one minute.
4. Aspirate the supernatant, as shown, removing as few cells as possible.
   

   

   Aspirate the supernatant, as shown, removing as few cells as possible
5. Resuspend the cells in 100 μL of Solution I, changing tips between samples.
6. Prepare Solution II by mixing 500 μL of 2% SDS with 500 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. In some cases the samples may appear to "clear" but don't worry if you don't see a big change. Place the tubes on ice for five minutes.
7. Add 150 μL of Solution III to each tube and immediately vortex each tube for 10 seconds with your vortex set at the highest setting. White clumps should appear in the solution after you vortex it. Place the tubes in the room temperature microfuge and spin them for 4 minutes.
8. While the tubes are spinning, label another set of eppendorf tubes with the plasmid names and your team color.
9. A white pellet should be visible when you remove your tubes from the microfuge. Use your P1000 to 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.
10. Add 1000 μL of room temperature 100% ethanol to each new tube. The tubes will be quite full. Close the caps and invert the tubes at least five times to thoroughly mix the contents.
11. Microfuge the samples for 2 minutes. It is important to orient your tubes in the microfuge this time since the pellets from this spin will be barely visible.
12. Remove the supernatants using your P1000 or the aspirator, but be careful not to disturb the pellet of plasmid DNA that is at the bottom of the 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.
13. Add 500 μL of 70% ethanol to each pellet. 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.
14. To completely dry the pellets, place your rack in the hood with the caps open for a few minutes. When the pellets are completely dry, add 50 μL of sterile water to each sample and vortex each tube for 2 X 30 seconds to completely dissolve the pellets. The liquid can be brought back to the bottom of the tubes by spinning them in the microfuge for a few seconds. Store the DNA on ice.

Part 3: Transform BL21(DE3) with mutant DNA

maybe skip concentrated stock this year? 10X worked for everyone last time i think

1. Prewarm and dry ten LB+Amp/Cam plates by placing them in the 37°C incubator, media side up with the lids ajar. You will perform two transformations for each of your five samples – one with a concentrated cell stock, and one with a 1:10 dilution of cells.
2. When your competent cells are ready, aliquot 75 μL of cells per pre-chilled eppendorf.
3. Add 2 μL of the appropriate DNA to each tube. Remember, you are testing plasmid DNA that was prepared from two different colonies for each of your two mutants, along with a no DNA control.
4. Flick to mix the contents and leave the tubes on ice for at least 5 minutes.
5. Heat shock the cells at 42°C for 90 seconds exactly and then put on ice for two minutes. Use your timer.
6. Move the samples to a rack on your bench then use your P1000 to add 0.5 ml of LB media to each eppendorf tube. Invert each tube to mix.
7. 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.
8. While you are waiting, prepare 4 large glass test tubes containing LB+Amp/Cam, and label them with your team color and sample name.
9. Also prepare 5 eppendorf tubes containing 180 μL of LB each. Use these to dilute your transformed cells 1:10 when you retrieve them from the incubator.
10. Plate 200 μl of each transformation mix on an LB+Amp/Cam plate. Make sure to label which concentration of cells was used on each 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 (if it sizzles, it's way too hot).
11. 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 each of your transformed XL1-Blue plates. Do the control samples have no colonies? Do the two mutations appear to have different efficiencies?

Part 5: Prepare sequencing reactions

also digest DNA today?

As we will discuss in lab today, 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? (Try using the Word Count feature on the the sequence document.)

The recommended composition of sequencing reactions is 200-500 ng of plasmid DNA and 3.2 pmoles of sequencing primer in a final volume of 12 μL. You will prepare two reactions per candidate (8 reactions total), one with a forward-reading primer and one with a reverse primer. Do not mix the primers together in one tube! 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 guess at the amount of plasmid DNA appropriate for our reactions.

For each reaction, combine the following reagents in an eppendorf tube:

• 2 μL of your plasmid DNA candidate
• 5.3 μL of a 1:100 dilution of the sequencing primer
• 12.7 μL sterile water

For each solution, mix by pipetting and then transfer 12 μL to an 8-PCR-tube strip. Keep track of which sample is in which tube (A-H), and label your tubes according to the table below. The teaching faculty will turn in the strips at the Biopolymers Laboratory in E17 for sequencing.

For next time

Reagent list