Alkaline lysis plasmid extraction

Solutions

Alkaline Lysis Solution 1

50 mM glucose

25 mM Tris-Cl (pH 8.0)

10 mM EDTA (pH 8.0)

Prepare Solution I from standard stocks in batches of -100 ml, autoclave for 15 minutes at 15 psi (l.05 kg/cm2 ) on liquid cycle, and store at 4 degrees C.

Alkaline Lysis Solution 2

0.2 N NaOH (freshly diluted from a 10 N stock)

1% (w/v ) SDS

Prepare Solution II fresh and use at room temperature.

Alkaline Lysis Solution 3

5 M potassium acetate - 60.0 ml

Glacial acetic acid - 11.5 mL

H2O - 28.5 ml

The resulting solution is 3 M with respect to potassium and 5 M with respect to acetate. Store the solution at 4 degrees C and transfer it to an ice bucket just before use.

Protocol

1. Pellet the overnight cultures in a 1.5ml eppendorf tube for 1 min. Spin down 4 to 4.5 ml of culture. (For low copy spin down about 6 ml.

2. Resuspend each pellet in 200ul of Alkaline Lysis Sol I, RnaseA added (final RNaseA concentration should be 100ug/ml) Make sure there are no lumps, homogenized.

3. Add 400ul Alkaline Lysis Sol II. Invert 4-6 times to mix. Do not allow reaction to lyse for more than 5min. Sample should clarify.

4. Add 300ul Alkaline Lysis Sol III. Invert 4-6 times to mix. Sample should have a white precipitate.

5. Add 100ul of chloroform. Do this in a fume hood. Invert 4-6 times to mix.

6. Rest on ice for 5-10min. This step is so that the chloroform does not get too hot in the centrifuges and leak out of the tubes. If you want to skip this step you might consider using less cholorform. I put the tubes at -20 for a couple of minutes.

7. Centrifuge at max. speed (14,000rpm) for 10min.

8. Pipet 750ul of supernatant/aqueous layer into a fresh tube. I do up to 800ul.

9. Add 1/10 volume (75ul) 3M NaOAc, pH 5.2. Flick to mix. 80ul if have 800ul of supernatant.

10. Add 0.7-1.0 volume COLD isopronanol (in freezer). Vortex/Flick to mix. If in a hurry go straight to step 11, otherwise rest on ice for 10-30minutes. I have even let it precipitate overnight at 4C if convenient. 600ul isopropanol. Then I put it at -20C for 5minuetes up to over the weekend if needed.

11. Centrifudge at max. speed for 25min. Most miniprep protocols say to do this at 4C, but I have not noticed decreased yield by centrifuging at room temp.

12. Remove and discard the supernatant. Don’t disturb the pellet. Sometimes I can’t see a pellet and more often than not I still have DNA.

13. Add 1ml of 70% EtOH (at room temp.) Invert 4-6 times to rinse the tube.

14. Centrifuge at max speed for ~5 minutes. Room temp. is fine. Remove and discard the EtOH.

15. Repeat steps 14 and 15 to remove all traces of isopropanol. Pulse spin after removing bulk of final EtOH wash and pipet off remaining EtOH.

16. Air dry the pellet for ~15min (pellet will change from white to clear as it dries). Resuspend desired volume (~30ul) of H2O or EB or T10E1 depending on downstream applications. If you pipet off the EtOH well, then I have done this for as little as 2 min. For fosmids, I usually resuspend the pellet in 20ul water.

Colony PCR

Colony PCR is an efficient way to screen for the construct with the desired insert. Colony PCR uses primers that attaches on the outside of the cloning site and amplify only the cloning region.

Master Mix
This is for 10ul reactions. The volume below is for one 10ul reaction. If doing more than one, multiply the amount by how many colonies you will be screening, and probably add one extra reaction worth to account for pipetting error.

Reagent Volume

Sterile H20 4ul

GoTaq 5.0ul

FWD primer 0.5ul

REV primer 0.5ul

In each PCR tube, place 1 colony DNA and 10ul of Master Mix.

Thermal Cycler Protocol

Step Temperature Time

1. 95C 3 min

2. 95C 30sec

3. 55C 30sec

4. 72C 3min (depends on length 1min/kb)

5. Go to step 2 24x

6. 72C 5 min

7. 4C Forever

After the thermal cycler is done, run on a 1% gel at 100V for 30min. Use colonies that display the correct band to make overnight cultures with antibiotic to miniprep the next day for sequencing.

Digestion for cloning

Sterile Milli-Q Water (20 - x) uL x=amount of reagents used Total volume should be 20uL

DNA (2000ng/[DNA])	_ uL
BSA* [10X]	2 uL
B4 Buffer	2 uL
Enzyme #1	1 uL
Enzyme #2	1 uL
ddH2O	_ uL
Total	20uL

*BSA should only be added if needed. Check the NEB poster on the -80 to see if it is needed.
Incubate for 1 hour in the 37 degree water bath
Notes:
Keep everything on ice when thawing and mixing reagents. Do Not vortex the master mix; you can gently bump it with your fingers and then spin it down for less than 10 seconds.
If your DNA [ ] is lower and your volume will exceed 20uL, scale up the amounts of B4 Buffer and BSA to 3uL each and make a 30uL total volume reaction.
IMPORTANT: Enzymes must be kept in the blue cold box.

Dishwashing protocol

1. Bleach / wait / remove stickers

2. Pour down drain

3. Rinse 2-3 times with tap water

4. Add alconox and brush if necessary

5. Rinse 3x with DI, put upside down on a rack

Electroporation

Use the electro-competent cells you just made for transformation. ALL OF THIS IS ON ICE

1. 100uL electrocompetent cell aliquots and UV sterilized cuvettes should already be in an ice bucket. (UV means 3 mins in the gel doc on full)

2. Label 13mm plastic test tubes beforehand, have 900uL of LB ready to pipette into the cuvette immediately after shocking

3. Set electroporator machine to bacteria

4. If plasmid DNA is ~150 ng/uL use 2.5uL. If 200 ng/uL, use 2uL (~400ng) into 100uL aliquots of electocomp cells

5. Use a kimwipe to wipe off any condensation from the exterior of the cuvette - important!!

6. Place the cuvette in the holder in the right orientation, slide in until you hear the click. The two electrodes should touch the sides of the cuvette

7. Take up 900uL of LB into the pipette tip

8. Press the SHOCK BUTTON

9. Immediately, pull out the cuvette and add the 900uL LB directly to the slit inside

10. Pipette up and down twice very gently. Transfer to the labelled 13mm tube (Don't make bubbles)

11. Repeat for all samples, INCLUDING YOUR NEGATIVE AND POSITIVE CONTROLS

12. The time constant reading should be between 3.8 and 5 ms (If it arcs, it is not usable)

13. Incubate in the gentle shaker (37 degrees and 100rpm) for 1 hour

14. Plate 100uL aliquots onto appropriate media, leave the remaining 800uL in the tubes to sit on the bench top overnight

15. For more difficult transformations, pellet 700uL of the culture, pour off supernatant and re-suspend pellet in 100uL of LB. Plate the concentrated 100uL of transformation.

16. To the remaining 200uL of transformation, add 1mL of LB and let sit on bench top overnight

17. Incubate all plates in 37oC incubator overnight, inspect for colonies in morning.

Ethanol Precipitation

1. 1/10 volume of Sodium Acetate (3 M, pH 5.2).

2. Add 2.5-3.0 X volume (calculated after addition of sodium acetate) of at least 95% (ice-cold) ethanol.

3. Incubate on ice for 15-30 minutes. In case of small DNA fragments (<100bp) or high dilutions overnight incubation gives best results, incubation below 0 °C does not significantly improve efficiency.

4. Centrifuge at > 14,000 x g (max speed) for 10 minutes at room temperature or 0 °C.

5. Remove supernatant being careful not to disturb the DNA pellet, which may be invisible. Remove any drops of liquid that adhere to the walls of the tube. (It is best to save the supernatant from valuable DNA samples until recovery of the precipitated DNA has been verified).

6. Fill tube 1/2 way with 70% Ethanol to rinse and recentrifuge at maximum speed for 2 minutes.

7. Repeat step 5.

8. If necessary, store the open tube on the bench at room temperature just until the last traces of fluid have evaporated.

9. Dissolve pellet (which is often invisible) in desired volume of buffer. (Usually TE [pH between 7.6 and 8.0], or water)

NOTE: Make sure to rinse the wall of the tube well with the buffer since up to 50% of the DNA will have been smeared on the wall. This can be done by pushing a bead of fluid over the surface of the wall using a disposable pipette tip.

Freezer stock preparation

Freezer Stocks

750ul of culture 250ul of 60% Glycerol Filter Sterilized using equipment in the Pfleger Lab Store them in -80C Combine in labelled (strain, date, initials) cryotube (lids & tubes in separate blue boxes on center of benches)

Gel extraction

If you are planning on extracting more than 2 fragments, make sure you add a ladder to the left of each well in order to split up the gel when you excise the band. This will lower the time for UV exposure.

1. Make a 1% gel, except increase the weight to 0.3g of agarose and 30mL of 1X TAE, and the 8 welled comb.

2. Pre-Weigh a micro-centrifuge tube for later use

3. To load a 30uL digestion, add 6uL of 6X Loading Dye to the digestion tube

4. Also load 1uL of dyed uncut DNA into another row (as the control), and 2Log Ladder into a third row to compare band sizes and to confirm the DNA has been digested correctly.

5. Run the gel at 100V for 30min (unless the gel bands are extremely close in size

6. Put up the UV shield on the gel imager, and find some gloves

7. During extraction, use the Prep UV setting on the gel imager so the UV isn’t as strong

8. Also you need to cut out (excise) the band very quickly (~5 sec)

9. Then take a picture of the rest of the gel to record the excised band

Note: The picture is taken after excision to reduce the amount of exposure time to the DNA. Too much exposure (~1min) will potentially cause thiamine dimmers and other mutations in your DNA. After this proceed using the direction found in the gel purification/extraction kit. Finally, elute in 25 uL of water.

Gibson Protocol

Written by Jackie in the Pfleger Lab

1. Generate forward and reverse primers to amplify insert and vector, refer to figure 1, can also use gibthon.org to design primers

Figure 1. Primer Design for 1 insert into a vector. Include at least 20 base pair homologous region over hang (pictured as the orange section on the gene of interest primers). For the insertion of more genes, just be sure each gene has an overlap region for the genes around it

Edit I (Ryan) designed primers with the gfp flappy ends for the vector as well, which aren't necessarily, but will most likely increase successful annealing.

2. PCR amplify insert and vector using the Tm of section that is actually annealing to the template and not the overall Tm that includes the overhang region

3. Gel extract the products to ensure correct amplification and to remove residual template plasmid. Can possibly Dpn1 the reaction then PCR purify to reduce background noise, but I have not tried yet

4. Add the following amount of Gibson Master Mix and vector and insert and mix together, then incubate at 50°C for 1 hour

Components	Suggested Amount
Gibson Master Mix	15uL
Plasmid	10-100ng
Insert (each)	2-3x of plasmid

5. Add 2 μL of reaction mix to competent cells (chemically competent DH5α), and transform according to standard protocol.
From the NEB site
I'm not sure if this is the same mix as Jackie, but NEB wrote this:
"When using the Gibson Assembly Master Mix product for electroporation, it is necessary to dilute the reaction 3-fold and use 1 μl for transformation"
How many Parts can we assemble?
Q4: How many fragments of DNA can be assembled in one reaction?
A4: The number of DNA segments that can be assembled in one reaction is dependent on the length and sequence of the fragments. Gibson Assembly has been used to efficiently assemble up to twelve 0.4 kb inserts into a vector at one time. However, we recommend the assembly of five or fewer inserts into a vector in one reaction in order to produce a clone with the correct insert. A strategy involving sequential assembly can be used if all of the fragments cannot be assembled in a single reaction.
Matt's gibson protocol

1) Run PCR rxn
- take small aliquot (3 PCR µl + 7 µl dH20 + 2 µl 6x dye) and run on a gel
- take the rest and PCR clean-up all of it
2) Dpn1 digest the PCR cleaned-up DNA
- (1 hr, 37º C, make sure you add the NEB buffer 4)
3) Run out a gel (extraction wells) of both insert PCR, and vector PCR
4) finish gel extraction protocol
5) Nanodrop time
6) Setup Gibson rxn - run for 1 hr at 50º C

Plasmid	10-100 ng (shoot for 50-75)
Insert	2-3 times whatever you put in for plasmid
dH20	20-(plasmid + insert)
Total	20µl

7) Transformation time, take 3µl from gibson rxn and transform directly using that handy protocol
8) Stay classy

Ligation

Calculations

For the ligation, you should follow the following calculations to determine the ideal amount to achieve the typical ratio of 3:1, insert:vector. Also the Ligation buffer vial should be made into 5uL aliquots to prevent constant freeze thaw and keep it active. The freeze thawing will eventually cause the sulfur ends to degrade. As a quick check, smell the vial to see if it still smells like rotten eggs (sulfur smell). A sulfur smell means the buffer is still good to use.
(Length of insert/ Length of vector) X 100ng of vector X 3 = ng of insert for a 3:1 ratio
On top of the standard ligation, run a control ligation by replacing the insert with water

Reaction

For 20 uL reaction (in this order):
• 2 uL ligase buffer (tubes marked with a green dash, in a box marked with a green dash in the -20)
• 17 uL of the above-calculated mixture of insert & vector & water
• 1 uL T4 DNA ligase (in enzyme box)
Run benchtop ligation 15 minutes. Take 3uL from this, and tranform it into competent cells. Take the remaining 17uL and place in strip tubes, then thermocycler for an overnight ligation at 16 degrees.




Limonene Production Assay w/ GC-MS Chromatography

This protocol is work in progress.


Day 1 :


1. Freshly prepare transformed strains which are capable of : (Cultures to be produced in triplicates)
a. LIMS1 synthesis in LB medium. (pBba5c + LIMS1)
b. ADS synthesis in LB medium. (pBba5c + ADS )
c. Untransformed cells in LB medium. (DH10B)


Make 5 ml liquid cultures of each in triplicates and incubate overnight @ 37 C.


Day 2


2. Overnight cultures to be diluted 1:100 in LB medium to 80 ml.
Induce with 500mM IPTG
Overlay the solution with (20%) 20 ml dodecane
Also, prepare a 80ml flask (blank) containing LB and overlay with 20% dodecane (20ml).
Add Limonene in the medium of the following volumes (to the three flasks) :
1. 0.15 ml
2. 0.075 ml
3. 0.0 ml


Incubated at 37*C for 72 h.


Day 5
At the end of the production period, dodecane layers are to be sampled for limonene content by gas chromatography/mass spectrometry on a DB-5 column using the internal standard caryophyllene to normalize between samples.


Amendment for amorphadiene:


GC-MS analysis of amorphadiene. Amorphadiene production by the various strains was measured by GC-MS as previously described21 by scanning only for two ions, the molecular ion (204 m/z) and the 189 m/z ion. Cells were grown in LB medium at 37 °C for 2 h and induced to express the ADS and the mevalonate pathway by the simultaneous addition of 0.5 mM IPTG and varying concentrations of mevalonate. Amorphadiene concentrations were converted to caryophyllene equivalents using a caryophyllene standard curve and the relative abundance of ions 189 and 204 m/z to their total ions. The sesquiterpene caryophyllene was purchased from Sigma-Aldrich.