20.109(S13):Induce protein and evaluate DNA (Day5)

Introduction

Last time you transformed your mutant DNA into BL21(DE3) cells. The colonies that arose were moved to liquid cultures, grown to saturation overnight, and then sub-cultured. Today you will add IPTG to these sub-cultures in mid-log phase in order to induce protein expression by the bacteria. Next time you will purify the resultant protein. We won’t shy away from telling you that there are many things that can go wrong at this stage! However, each one is certainly a learning experience.

As evidenced by Nagai’s work, wild-type inverse pericam is not toxic to BL21(DE3) cells. Although it is unlikely for your small mutation to dramatically change this fact, in general a novel protein may turn out to be toxic. If this is the case, only very small amounts of protein are produced before the bacteria die. Keep in mind that overexpressing a single protein may come at the expense of producing proteins needed for survival, and will most likely cause cell death eventually; however, toxic proteins hasten this demise. Aberrant toxicity can sometimes be alleviated by reducing the culture temperature (e.g., to 30 °C).

Based on its fluorescence activity, wild-type inverse pericam allows proper folding of (cp)EYFP, and based on its response to calcium, it also allows calmodulin to fold. One problem you may encounter is that your mutant proteins will no longer fold correctly. Since you made mutations in the calcium sensor part of IPC, rather than the fluorescent part, it is unlikely that your protein will destroy EYFP fluorescence. However, a common problem with misfolded proteins is the formation of insoluble aggregates, due for instance to improperly exposed hydrophobic surfaces. Proteins can be purified from these aggregates – called inclusion bodies – but the process is more labor-intensive than for soluble proteins. (The proteins must be extracted under more harsh conditions than you will use next time, then purified under denaturing conditions, before finally attempting to renature the proteins.) Inclusion bodies sometimes form simply due to very high expression of the protein of interest, causing it to pass its solubility limit. This outcome can be prevented by lowering the culture temperature or time, the amount of IPTG, or the growth phase of the bacteria.

One final point to keep in mind is that not all proteins can be produced in bacteria. Eukaryotic proteins that require post-translational modifications (such as glycosylation) for activity require eukaryotic hosts (such as yeast, or the ubiquitous CHO – Chinese hamster ovary – cells). Sometimes eukaryote-derived proteins will be truncated or otherwise mistranslated by E. coli due to differential codon bias; errors in translation can be prevented by providing additional tRNAs to the culture or directly to the bacteria via plasmids. Despite all this complexity, prokaryotic hosts have been plenty good enough to produce proteins for certain therapies, notably the cytokine G-CSF. This cytokine is taken by patients needing to replenish their white blood cells (e.g., after chemotherapy), and sold as Neupogen by the company Amgen.

After you induce your cells with IPTG, you will let the resultant protein factories do their work for 2-3 hours. During this time, you will evaluate the DNA from your two X#Z candidates and from the reference mutant. First, you will run your diagnostic digests from last time out on a gel. The banding patterns will allow you to determine (i.e., diagnose) whether either of your putative X#Z mutants actually contains the new restriction site that you introduced. Of course, there is a slim possibility that the silent mutation was incorporated but the non-silent mutation wasn’t. To get more direct evidence for whether the site-directed mutagenesis worked, you will analyze data from the sequencing reactions that you set up last time.

You're already sequencing pros after your Module 1 experience, of course, but feel free to review the relevant background here (last couple of paragraphs).

Now you might be wondering why you would ever go through the trouble of designing and performing diagnostic digests, when sequencing is relatively simple and yields more information. Here, the idea of scale becomes important. Sequencing costs $6-8 per reaction, which can add up if you need to examine, say, 10 or more candidates. Agarose gel electrophoresis, by comparison, costs perhaps $1 per candidate. Since both methods require DNA isolation, one is not dramatically more labor intensive than the other. (A method called colony PCR avoids this labor. Can you guess what it might entail?) Finally, banding patterns can give a quick readout of many candidate colonies at once compared to the time it takes for the individual sequencing analyses you will perform today. Of course, there's no reason one couldn't automate the analysis process with a bit of (computer, not DNA) code!

Protocols

Part 1: Cell measurement and IPTG induction

1. Pick up a 6 mL aliquot of BL21(DE3) cells carrying each mutant plasmid (X#Z 1 and 2, and the reference mutant) as well as an aliquot with wild-type inverse pericam. These cells should be in or close to the mid-log phase of growth for good induction, just as they were for transformation. Like last time, check the OD₆₀₀ values of your cells (650-700 μL of a 1:10 dilution) until they fall between 0.4 and 0.8.
   • OD values at the higher end should favor more protein production.
2. Once your cells have reached the appropriate growth phase, set aside - on ice - 1.5 mL of cells from each tube as a control (no IPTG) sample.
3. Take an aliquot of cold IPTG (0.1 M), and add to your remaining 4.5 mL of cells at a final concentration of 1 mM. You should prepare three mutant and one wild-type tube.
4. Return your tubes to the rotary shaker in the 37 °C incubator, and note down the time.
5. While your IPTG-stimulated cells are producing protein, you will analyze the sequence data and restriction digests of the plasmids they are carrying. At the end of the day, you will choose only one of your X#Z candidates to save, and aspirate the other into your bleach flask.

Part 2: Run diagnostic gel

The scheme below assumes that both Digest 1 (D1, used to analyze the positive control mutant) and Digest 2 (D2, used to analyze your X#Z mutant candidates) use only one enzyme. If you are doing double-digests and need to run single enzyme controls, hopefully you spoke to the teaching faculty about this plan last time. Load your samples on a 1% agarose gel (or a higher-weight percent gel, for those groups who are expecting a small band size) in the following order, using 10 μL per ladder and 20 μL per plasmid:

Lane	Sample	Digest
1	IPC	uncut
2	IPC	D1
3	(+) control	D1
4	1 Kb marker
5	IPC	D2
6	X#Z-1	D2
7	X#Z-2	D2
8	100 bp marker	(if relevant for your band sizes)
9	X#Z-1	uncut
10	X#Z-2	uncut

Once all the samples are loaded, the power will be applied (100 V for 45 minutes), and then you will photograph your gels.

When the gel is ready, you will compare the band sizes in the photograph with the expected band sizes that you previously calculated. In the meantime, you can analyze your sequence data.

Before you leave today, please upload your gel image according to the filenames on today's Talk page. The code should automatically display your image on the Talk page after the upload is complete.

Part 3: Analyze sequence data

Your goal today is to analyze the sequencing data for three samples - two independent colonies from your X#Z mutant, and one reference mutant clone for practice - and then decide which colony to proceed with for the X#Z mutant.

You will want to have the pRSET-IPC ApE file handy, and to mark and/or note down the expected location of your mutation before proceeding. (Just compare to your annotation of the IPC alone ApE file that you prepared on Day 1 of the module.) The data from Genewiz is available at this link. Choose the "Login" link and then use "astachow@mit.edu" and "be20109" to log in. At the bottom right should be a link to download your sequencing results.

Begin by analyzing your reference mutant, and then move on to your unique mutant.

• Order date 03-25-13, #10-220995825 has T/R data as well as some reference mutant data. In particular, sample 3 is D24H and sample 4 is E67K (not WT and D24H as written).
• Order date 03-25-13, #10-220996747 has W/F data.
• Order date 04-04-2012 (select "More..." and then use the search function) has T79P and M124S data.

The quickest way to start working with your data is to follow the "View" link under the Seq File heading. For ambiguous data, you may want to look directly at the Trace File as well.

Recall from Module 1 that you can align your sequencing data with a known sequence, in this case inverse pericam, and the differences will be quickly identified. There are several web-based programs for aligning sequences and still more programs that can be purchased. The steps for using one web-based tool, the same one you used in Module 1, are sketched below. Another popular alignment program is CLUSTAL.

Align with "bl2seq" from NCBI

1. The alignment program can be accessed through the NCBI BLAST page or directly from this link. The default settings should be fine.
2. Paste the sequence text from your sequencing run into the "Query" box. This will now be the "query." If there were ambiguous areas of your sequencing results, these will be listed as "N" rather than "A" "T" "G" or "C" and it's fine to include Ns in the query.
3. Paste the inverse pericam sequence into the "Subject" box.
4. Click on the BLAST button. Matches will be shown by vertical lines between the aligned sequences. You should see a long stream of matches, followed by lots of errors in the last ~200bp of the sequence – ignore the error-ridden part of the data, as it may not accurately reflect your mutant plasmid. In this stream of matches, the 1-3 missing lines indicating your mutant codon should stand out. If they don’t, use the numbering or Find tool to locate the appropriate codon.
5. You should print a screenshot of each alignment to pdf (and to paper if you desire). These will be used to prepare a figure showing what you found today. You might want to email yourself the alignment screen shots or post them to your wiki userpage.

If both colonies for your mutant have the correct sequence, flip a coin and proceed with one or the other; ditto if both are inconclusive. If one appears right and the other doesn’t, of course proceed with the former. Finally, if both are clearly wrong, talk to a member of the teaching faculty.

Part 4: Observe mutant colonies

Last time you transformed BL21(DE3) cells with three different plasmids (two candidates for the X#Z mutant, and one D24H/E67K/T79P/M124S clone). Compare the relative colony formation of cells carrying the different plasmids. If all the plates have dense cell growth, there is no need for you to get an exact colony count; just do your best to get a relative estimate, and describe any findings in your notebook.

Part 5: Cell observation and collection

1. After ~2-3 hours, you will pour 1.5 mL from each tube (from Part 1) into a labeled eppendorf, then spin for 1 min. at maximum speed. Save the other 3 mL!
   • A minimum of 2.5 hours is best, assuming that timing doesn't take you past 4:45 pm.
2. Aspirate the supernatant from each eppendorf, using a fresh yellow pipet tip on the end of the glass pipet each time.
3. Observe the color of each of your pellets, and compare to the examples on today's Talk page. If the wild-type and both mutant pellets all appear yellow-greenish to the eye, proceed as follows:
   • Do NOT toss the rest of the liquid cultures. First, measure their OD₆₀₀ values, according to part 6 of today's protocol.
   • Next, pour 1.5 mL more of the relevant liquid culture on top of each pellet, spin again, and aspirate the supernatant.
   • The last 1.5 mL of culture may be aspirated in your vacuum flask, to be later bleached and discarded.
4. If one or more of your pellets are white or only dimly colored, please ask one of the teaching staff to show you the room temperature rotary shaker. You will continue to grow your bacteria overnight. Tomorrow morning, the teaching staff will collect your pellets for you and freeze them. As you can see above, the +IPTG pellets are from 3 mL of culture, while the -IPTG pellets come from 1.5 mL of culture.

Part 6: Preparation for next time

Next time, you will lyse your bacterial samples to release their proteins, and prepare to run these out on a protein gel. In order to compare the amount of protein in the -IPTG versus +IPTG samples, you would like to normalize by the number of cells. At the end of today, you may have only three samples ready (-IPTG only), or you may have all six. In either case, measure the OD₆₀₀ of a 1:10 dilution of cells for each finished sample (actually, for the -IPTG samples you have done so already), and write this number down in your notebook and on today's Talk page. Then spin down the cells and aspirate the supernatant. Give the cell pellets to the teaching faculty; they will be stored frozen at -20 °C. (Be sure to make a 2X pellet for the +IPTG samples.)

For next time

1. Your Module 1 report revision is due by 11 AM on Friday, to the 20109.submit address and using a TeamColor_LabSection_Mod1-REV.doc naming scheme.
2. Your second reflection, written individually about lessons learned from the report revision process, is also nominally due Friday. We are happy to accept these within about 24 hours after you submit your report (no late penalty), but don't want to delay further than that as you may start to forget about the revision experience.
3. Day 6 of this module is pretty packed, and has the potential to run long. Thankfully, some changes implemented last year dramatically shortened the day. Nevertheless, you may want to do a little reading in advance, and perhaps even fill in the table under "Advance preparation for PAGE" based on the OD values you measured. Be sure to post these to the Day 5 Talk page also if you haven't already.

Reagent List

• IPTG (isopropyl β-D-1-thiogalactoside), 0.1M

• Loading Dye
  • 0.25% xylene cyanol
  • 30% glycerol
  • RNase

• 1% and 1.2% agarose gels with SYBR Safe

• Gels made and run in 1X TAE buffer
  • 40 mM Tris
  • 20 mM Acetic Acid
  • 1 mM EDTA, pH 8.3