Team:Stanford-Brown/Protocols

Methods

General Protocols

PCR (Polymerase Chain Reaction)

PCR vs. Colony PCR

There are a couple of different ways to introduce your template DNA, or the DNA you want to amplify, into a PCR mixture, and the method you use will depend on the source of the template DNA. The way I see it, there are four different scenarios:

1. Amplifying a genetic segment within a plasmid or isolated sample of DNA

This is the most basic scenario, where you have a tube of linear or plasmid DNA. This situation will definitely come up if you are trying to amplify a part from the registry directly, without going through the time-intensive process of transforming the part into e.coli and waiting for it to grow (quick note: you should go through the time-intensive transformation in parallel regardless, because you will need to replicate the part and make cryostocks of it, therefore saving it for later). In this case, you need first to know the concentration of your sample. If you don’t know it or it was not provided, you can learn the concentration for your sample by using the nanodrop machine located in room 347. It depends on the size of your template, but as a general rule, you need on the order of 25-50 ng template minimum for a successful PCR, so adjust the volume of your template in your PCR accordingly.

2. Colony PCR

You can also amplify both plasmid and genomic DNA straight from live cultures of organisms containing your desired sequence. You will usually have cultures in one of two forms: either in liquid culture, or spread on an agar plate. If you are amplifying from liquid culture, grow it up as much as you can and add 1μL of the culture to the PCR mix. If you're amplifying from the plate, there is no need to add a volume; instead, simply take a pipette with a pipette tip from the green box, gently touch the pipette tip to the desired colony on the plate (try to take as little from the plate as possible; agar can screw up PCRs), and then insert your pipette tip into the PCR mixture, pipette in and out to mix, and you’re done.

Polymerases and Supermixes

Platinum Blue PCR Supermix:

Platinum Blue is a 1.1X supermix, complete with dNTPs, buffer, and Taq polymerase. Taq is the polymerase that was isolated from extremophiles living in hot springs in Yellowstone, which you surely have all learned about in previous bio classes. Taq is the most pervasive polymerase used in molecular bio, and there are a host of downstream workflow kits and applications that are specifically designed to work with PCR amplicons produced by Taq polymerase (e.g. TOPO TA cloning kit). This specializations stems from the fact that Taq polymerase adds a single Adenine (A) base overhang on the 3’ ends of the amplicons; these overhangs need to be accounted for in subsequent ligation steps, and any ligation partner needs to have matching T overhangs to hook up with the Adenines.

For 20 μL rxn:

18 μL supermix

1 μL template DNA (assuming an adequate concentration)

.5 μL forward primer

.5 μL reverse primer

For 50 μL rxns:

45 μL supermix

1-3 μL template 1 μL forward primer (if using 1/10 dilution, this can be .7 μL)

1 μL reverse primer (if using 1/10 dilution, this can be .7 μL)

Add nuclease free water to 50 μL volume

rTaq PCR Supermix:

rTaq is a 2X supermix, meaning it needs to take up 50% of your reaction volume. It is inferior to platinum blue, as I think it produces more errors, and therefore it should not be used for accurate sequencing, or for any downstream application. That being said, it is really good for validation and testing to make sure constructs have made it into plasmids and stuff like that.

For 20 μL rxns:

10 μL supermix

1 μL DNA template (can be more if need be, since we have more leeway with rTaq and the requisite volumes of components)

.5 μL forward primer

.5 μL reverse primer

8 μL nuclease free water (or less, depending on what your volume of template is)

N.B.: This reaction can be scaled up to 50 μL rxns.

Phusion Polymerase:

Phusion is a high-fidelity, proofreading polymerase that absolutely flies: it is about 4x faster than your typical Taq polymerase you find in supermixes, and this speed will need to be accounted for in the Thermocycle conditions (see below). Also unlike Taq, Phusion produces blunt-end amplicons, and consequentially you may need to make adjustments to any downstream workflow plans (e.g. blunt end ligations, gibson cloning, etc.)

For 20 μL rxn:

4 μL 5x HF reaction buffer (I have never used GC buffer and don’t really bother with it.)

.4 μL 10mM dNTPS

.5 μL forward primer

.5 μL reverse primer

.5-14.4 μL template DNA

.1-.6 μL DMSO (OPTIONAL) - I use DMSO when I am doing a colony PCR sometimes, especially if I am redoing it because it didn’t work the first time. DMSO serves to relax genomic DNA, making it easier to read and amplify.

Nuclease water filled to 19.8 μL

.2 μL Phusion enzyme

N.B.: You will never add this much template but when putting together your own PCR mixtures and not relying on supermixes, you have a ton of leeway on how much template you can add, so long as it is suspended in water and none of that buffer EB crap that comes in the Qiagen kits. Never use that stuff, as it does have effects on the success of your PCR reactions.

Thermocycler Conditions

Taq polymerase (Platinum Blue and rTaq supermixes)

1. Initial Denature: 95 ̊C 2 min - The official Platinum Blue protocol calls for 94 ̊C for 3 min, use judgement as either will work.

2. Denature: 94 ̊C 15-30 secs - Use a shorter time if the amplicon is a relatively short segment of DNA, and a longer time if it is a relatively long piece of DNA.

3. Annealing X ̊C 15-30 secs - This is the most crucial step of the thermocycle! Your annealing temperature will be determined by the melting temperature of your primers. As a general rule, your annealing temperature should be about 5 ̊ lower than the lowest melting temperature of your primer pair. Additionally, if you are trying to add tails to your amplicon (e.g. you are trying to add restriction sites to the ends of your DNA template), you may need to drop the annealing temperature down even more. I have had primers with melting temperatures above 65 ̊ that needed to be annealed at 42 ̊. Additionally, if a primer may be difficult to anneal to the template, you can increase the annealing time for better results.

4. Extension 72 ̊ X seconds - Taq extension runs at 1kb per minute. Therefore, allowing the extension step enough time to fully copy the entire amplicon.

5. Repeat steps 2-4 32X

6. Final Extension 72 ̊C 5 min

7. Hold at 4 ̊C forever.

Phusion

1. Initial Denature 98 ̊C 30 sec

2. Denature 98 ̊C 10 sec

3. Annealing X ̊C 15-30 sec - Same principles of annealing with Taq polymerase apply here.

4. Extension 72 ̊C X seconds - Phusion is much faster than Taq, and requires 15-30 sec per kb. 5. Repeat steps 2-4 32X

6. Final extension 72 ̊C 5 min - You want to be a bit careful here; I wouldn’t run the final extension step for any longer than 5 min; Phusion is supposedly a very active enzyme, and I’ve been told it is best to not let it run for too long. Although I have never seen it happen, supposedly it can add random bases to the ends of amplicons.

7. Hold 4 ̊C forever.

N.B.: The standard protocols for various polymerases can be found at these addresses:

Platinum Blue Supermix: (http://tools.invitrogen.com/content/sfs/manuals/ platinumbluesupermix_man.pdf) rTaq: Phusion: (http://www.finnzymes.com/pdf/ f530_phusion_high_fidelity_dna_polymerase_prodinfo_low.pdf)

Primers

Designing Primers:

-Choose a forward and reverse primer from a location in the gene or plasmid that is sure to include the portion desired for amplification or sequencing

-For sequencing, it is desirable if possible to have primers that fall 50-150bp outside your desired region, to ensure that accurate reading occurs for the whole gene (often the first and last ~100bp in the read are very inaccurate). While for PCR, remember that the sequence portion corresponding to the primers themselves will be amplified, and also primers should normally be between 15-30bp in length (around 20bp is ideal) -desired melting temperatures are generally between 55-65C as you will see, melting temperature is a function of length and GC content, so it is often difficult to design primers in regions much greater than 50% AT. Forward and reverse primers should have the same melting temperature, or with a difference of no more than 2 degrees.

-The annealing temperature used for a pair of primers should be set at 5 degrees below the lower melting point of the primer pair. Using a tool like ApE or Geneious makes it easy to select certain sections of a sequence to check for primer features like melting point and GC content. IDTs 'Oligo Analyzer' is a great tool to check for primer dimerization, hairpin structures, etc. (http://www.idtdna.com/analyzer/Applications/OligoAnalyzer/). Use this tool or something like it as a final check to make sure your primers will not be likely to react with themselves or each other around the temperatures they will be active for gene interaction

-NCBIs Primer Blast is another great tool. It can be used both to help design the primers and to ensure that the primers you choose will not amplify any genomic DNA in a colony based amplification (http://www.ncbi.nlm.nih.gov/tools/primer-blast/)

Ordering Primers:

-All primer orders are from ELIM Biopharm (http://elimbio.com/) -Usually any primers you order during normal work hours will arrive next day

Primer Dilution (stock preparation):

-Once you receive your primers, you need to dilute them; They can be dilutions can be made 1/10 or 1/20, both work well as long as you keep track and adjust your reactions appropriately. -Typically we create 100-200μl working stocks; it will take a long time to use up that much primer

PCR Cleanup (using Wizard SV Gel and PCR Purification System)  Sample Prep

A. Gel Extraction:

- Following electrophoresis, excise DNA band from gel and place gel slice in a 1.5ml microcentrifuge tube.

- Mass gel slice (by massing the tube containing the slice and subtracting the mass of an empty tube)

-Add 10μl Membrane Binding Solution per 10 mg of gel slice. Vortex and incubate at 50–65°C until gel slice is completely dissolved (usually 10-15 minutes)

B. PCR Amplifications:

-Add an equal volume of Membrane Binding Solution to the PCR amplification.

Binding of DNA

1. Insert SV Minicolumn into Collection Tube.

2. Transfer dissolved gel mixture or prepared PCR product to the Minicolumn assembly. Incubate at room temperature for 1 minute.

3. Centrifuge at 12,000 × g for 1 minute. Discard flowthrough and reinsert Minicolumn into Collection Tube. If you are worried about the final concentration of your purified product, you can repeat this step to maximize the amount of DNA bound to the filter.

Washing

4. Add 700μl Membrane Wash Solution (ethanol added). Centrifuge at 12,000 × g for 1 minute. Discard flowthrough and reinsert Minicolumn into Collection Tube.

5. Repeat Step 4 with 500μl Membrane Wash Solution. Centrifuge at 12,000 × g for 5 minutes.

6. Empty the Collection Tube and recentrifuge the column assembly for 1 minute with the microcentrifuge lid open (or off) to allow evaporation of any residual ethanol.

Elution

7. Carefully transfer Minicolumn to a clean 1.5ml microcentrifuge tube.

8. Add 30-50 μl of Nuclease-Free Water to the Minicolumn. Incubate at room temperature for 1 minute. Centrifuge at 12,000 × g for 1 minute. By adding less water, like 30 μl, you will increase the concentration but decrease the total amount of product. On the flipside, if you want to maximize product, you can maximize elution volume so long as you don’t care about concentration.

N.B.: you can also increase yield by warming the elution water before hand. I usually warm it to 40 ̊C with good results.

9. Discard Minicolumn and take sample to nanodrop (see 'Nanodrop', below) 10. Store DNA at 4°C (temporary, ~few weeks) or –20°C (long-term).  The standard protocols for the SV Wizard Gel and PCR purification kit can be found here: (http://www.promega.com/resources/protocols/technical-bulletins/ 101/wizard-sv-gel-and-pcr-cleanup-system-protocol/)

Hell Cell

Venus Life

Construct Development

Potential cell-cycle dependent promoters in E. coli and their sequences were identified from literature (Quiñones 1997, Sun 1994, Ogawa 1994). Primers were designed to isolate these sequences and add on BioBrick cut sites according to the Silver lab method (Phillips 2006). Colonies of E. coli were selected from plates and sequences were isolated using colony PCR. Sequences were verified using BioPharm oligo sequencing and gel electrophoresis. PCR product was digested for 2 hours at 37C with EcoRI and SpeI. Digestion products were isolated via gel extraction; two samples were ligated with pSB1C3 cut with EcoRI and SpeI, and BBa_E0840 cut with EcoRI and XbaI, respectively. Ligation products were transformed into NEB-5a competent cells via heat-shock; promoter-pSB1C3 strains were plated then inoculated in LB+Chloramphenicol and promoter-E0840 strains were plated then inoculated in LB+Ampicillin.

Bulk Assay

50mL cultures of promoter-E0840 were grown overnight in 125mL flasks with LB+Amp, and some culture was diluted into 2 15mL samples with OD600 between 0.1 and 0.3. To synchronize cell cycle, serine hydroxamate (SHX) was added to both replicates to a final concentration of 10mg/mL and allowed to incubate at 37C for 1.5 hours (Ferullo 2009). After incubation, OD600 was taken to verify cell-cycle arrest. Samples were pelleted and resuspended in LB+Amp and replaced in the 37C shaking incubator.

Every 2.5 minutes, 1mL sample was removed from each replicate and OD600 was measured. The same sample was transferred into a microcentrifuge tube, pelleted, aspirated, and resuspended in M9. This sample was then measured in the fluorometer.

Microscopy Assay

4mL cultures of promoter-E0840 were grown overnight in 15mL conicals with LB+Amp. At an OD600 reading between 01. and 0.3, 40uL of 100X SHX was added to synchronize cell cycle. After 90 minute incubation, cells were pelleted and resuspended in LB+Amp without serine hydroxamate. 10uL of sample was pipetted onto a glass slide then covered with an agar pad. A timelapse was taken for 2 hours, taking images every 5 minutes.

Biomining

Flagella Removal Protocol

Procedure from Westerlund-Wikstrom Paper Cited

For large scale removal and purification of the flagella, we hoped to adopt the purification method stated in the paper entitled “Functional expression of adhesive peptides on flagellin”. The method outlined involved four steps:

1. Shearing of flagella from bacteria

2. Separating the cells from flagella

3. Purifying the flagella

4. Detection of the flagella

In the first step, agar plates of the bacteria were collected in Tris-HCL buffer. The cells then had their flagella sheared off by a Turrax homogenizer at 20,000 r.p.m. The next step included pelleting the cells, and re-centrifuging the supernatant to ensure that all residual bacterium were removed. The supernatant, which contained the flagella, was then ultracentrifuged to form a pellet which was reconstituted in 1ml 10mM Tris-HCL and 0.5% deoxycholate. The third step then involved purifying the flagella. This was accomplished by isopycnic ultracentrifugation in a 10-60% sucrose gradient in Tris-DOC. The complete conditions for ultracentrifugation are outlined within the article. Concentrated light was then used to illuminate and detect the flagella, which were then collected and dialysed against Tris and distilled water.

Once the flagella have been collected, we would need to separate the metal ions for collection and use. The purified intact flagella (complete with metal binding peptides and metal ions) would be collected on Gelman cellulose acetate filters. The Tris-DOC buffer would be washed through with 0.5M KCL. The filter and flagella were then immersed in 5M urea - 0.05M KCL for half an hour at 26 C. At this point, the filaments (flagellin) dissociated while the hook-basal body complexes that make up the rest of the flagella were eluted.