Team:Utah State/Notebook

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Latest revision as of 21:52, 28 September 2011

Notebook

Protocols

Introduction

The majority of iGEM BioBrick assemblies will follow a repeating cycle of procedures, at each cycle more BioBricks are added to the growing construct. For simplicity, we will start with DNA transformation. We have a tube of purified DNA with our BioBrick construct in an antibiotic resistant transformation plasmid, and we insert this into the E. coli cells. Once we get the DNA into the cells, we spread the cells onto a plate and let them grow up overnight. In the morning, we select colonies from the plate and create a stock streak plate and a liquid culture. Once the liquid culture is grown, a portion is used to make a long-term frozen glycerol stock solution, and the rest is used to extract a large amount of plasmid DNA. Once the DNA is extracted, we cut with the correct restriction enzymes for our assembly to give a tube of the linear BioBrick and linear plasmid. These linear pieces of DNA are then separated by size on a gel, and the DNA from the gel is extracted and purified. The DNA pieces of interest are then combined in an overnight ligation reaction, which can then be transformed into E. coli, continuing the cyclic procedure.

I. Electroporation Transformation of E. coli

Transformations are any procedure used to insert DNA into a bacteria (if you use a virus, the term becomes transfection). Electroporation uses a pulse of electricity to disrupt the cell membrane and create holes that would allow the DNA to enter the cell. Cells need to be made competent before doing this procedure, in order for them to efficiently take up the DNA. Transformations generally utilize millions to billions of cells and DNA molecules and, for a transformation to be successful, only one molecule of DNA needs to enter into one cell, which then grows into a colony. One issue with transformations is selecting and verifying which colonies have the desired DNA. This is usually done using a marker, a characteristic possessed by cells that have the DNA (or lost by those cells) that distinguishes it from the rest of the colonies that grow up. Commonly, this is the expression of an antibiotic resistance gene included on the transformed DNA, which allows only the cells that have taken up the DNA to survive on a plate in the presence of that antibiotic. Sometimes pigment producing or fluorescent/luminescent proteins can also be used in place of antibiotic resistance to allow visual determination of transformed colonies. Other ways of selection exist, but will not be discussed here.

1. Turn on ice machine

2. Thaw DNA solutions

3. Clean and sterilize the electroporation cuvettes by washing with double distilled water (ddH2O) twice and then fill the cuvettes with ethanol.

4. Let cuvettes sit with ethanol for 5-10 minutes, then wash 4-8 times with ddH2O

5. Place cuvettes on ice

6. Take competent cells out of the -80 °C freezer, and thaw them on ice

7. Add 3 µL of DNA to the cell solution. (This should be around 100-250 ng of total DNA, too much DNA causes arcing, too little gives few transformed colonies).

8. Incubate on ice for 5 minutes.

9. Add 60 µL WB buffer (10% glycerol). This helps reduce arcing, although too much can lower numbers of transformed colonies.

10. Set the electroporation machine to 2500 V, 200 Ω, and 25 µF for E. coli.

11. Transfer the cell/DNA/WB solution into the cuvettes by pipetting up and down in the 1.5 mL tube first to mix. Make sure the pipette tip is between the metal plates on the cuvette before ejecting the solution. Keep the cuvettes on ice.

12. Before electroporating, dry the cuvettes of with a KimWipe, to ensure no liquid on the surface that could create other paths for the electric pulse (and could cause arcing).

13. Pulse the cells and return cuvette to the ice. Check the time constant on the machine, a constant of 4.5+ is a very good transformation, and will yield many colonies. A constant of 2.5-4.5 is okay, and will still work. Constants below 2.5 will yield very low colony numbers, and may need to be redone. NOTE: addition of extra WB or lower amounts of DNA will reduce the time constant as well, so it is only a rough measure.

14. To remove the cells from the cuvette add 1 mL LB media or SOC media (no antibiotic in this media) to the cuvette. Pipette up and down a few times to mix.

15. Remove the solution to just above the two plates in the first removal pipetting (~1/2 of the volume) and transfer it to the original cell tube (NOT THE DNA TUBE). Then, tip the cuvette on its side so that the space between the plates is vertical, place the 1000 µL pipette tip between the plates, and slowly draw up the solution, while tipping the cuvette further. This should draw up the rest of the liquid in the cuvette.

16. Incubate the cell solutions at 37 °C for 1-2 hours (can go up to three, but try to avoid doing it for that long).

17. Plate the cells on plates containing the correct antibiotic. Each transformation requires two plates. Add 500 µL of solution to one plate, spread with the spreading stick, and then spread the spreading stick on the second plate without adding any solution to it. This creates a dilution plate in case you have thousands of colonies on the first plate. It is roughly a 1:100 to 1:200 dilution.

18. Grow the plates up overnight at 37 °C. Do not leave for longer than 24 hours, as contaminants might have a chance to grow and the plates could dry out.

If your cuvette arcs (bright flash and loud popping noise during electroporation): 1. Clean the electroporator lid.

2. Wash out and sterile the cuvette with ethanol – the cells have been pretty much killed and will not be usable in plating, so you need to restart.

3. Add less DNA to the cells (reduce by 25%-33%).

4. Add an additional 15 µL of WB buffer to the solution.

II. Colony selection, Stock Streak Plating, and Culture Preparation

After allowing transformed cells to grow up overnight, we can then select for the colonies that exhibit the characteristics indicating that they have taken up our DNA of interest. It is important at this point to know all the possible DNA combinations that could be in these cells, and which set of characteristics we want. For example, if we are converting a construct from one plasmid to another and replacing an RFP expression system on the final plasmid with our construct, we would want to select cells that did not express RFP, as these are clearly wrong. One important thing to note is that when using a RFP/GFP color or fluorescence to indicate the correct or incorrect colonies, sometimes these characteristics are not visible at the small colony stage, and a second round of selection on a streak plate is necessary. Luckily, streak plating is necessary for all types of selection, as it acts as an intermediate storage stock until glycerol stocks are created.

1. Carefully decide which cell characteristics you are looking for.

2. Identify colonies with the characteristic that are separated by a small distance from other colonies – this makes it easier to pick only the colony you want

3. With a sterile toothpick or sterile pipette tip, lightly touch down on the colony to pick up cells

4. Keeping the toothpick or tip oriented so that the cells are on the bottom, gently streak the cells on a new plate with the correct antibiotic. DON’T DISCARD THE TIP/TOOTHPICK. The streak should be around 2 cm or so long, and should be separated from other streaks on the new plate.

5. If you want to grow the cells up in a liquid culture (for glycerol stock and DNA prep), you can place the tip/toothpick into a tube with 5-6 mL LB. There should be enough cells still on the tip/toothpick to have a dense culture overnight. If there is a strong chance that not all colonies picked will be correct (see RFP example above), 2-3 of these cultures should be made for each construct; otherwise 1-2 will usually be enough.

6. If colonies are used to grow up liquid culture, label the streak and the culture tube with the same number, so if the colony is incorrect, you know which tube is incorrect.

7. Repeat for as many colonies as you think you will need (15-25 is generally good for streaks)

8. Put the plate into the 37 °C incubator overnight

9. Re-evaluate the streaks for indications that they are not the right colony. Toss any liquid cultures corresponding to incorrect colonies.

III. Glycerol Stock Preparation

Glycerol Stocks are one of the best ways to store cells and DNA for long periods of time. These stocks are prepared so that the cells in the tubes are still alive, and capable of creating new liquid cultures from small amounts of frozen stock. This allows for quick growth and extraction of important DNA without spending an extra day and extra supplies re-transforming a construct every time it is needed. In order to freeze cells and still keep them alive, a cryoprotectant is added to the cultures. Cyroprotectants function by reducing the freezing point of the solution and act to reduce the formation of large ice crystals inside cells that could rupture membranes. Cryoprotectants are non-toxic to the cells, and are generally able to pass through the membranes into the cells.

1. Take 1 mL of overnight E. coli culture and add it to a clean, labeled 1.5 mL tube.

2. Add 200 µL of 80% glycerol to the tube (this creates a roughly 15% glycerol solution).

3. Mix well by inverting the tubes (unmixed glycerol will tend to stay separate from the cell solution.

4. Immediately place the cells into a -80 °C freezer box. There are often special freezing boxes that let the freezing occur more slowly, but for E. coli these are generally not needed.

5. To use the glycerol stock to establish a new culture, either scrape a very small amount of the frozen culture off with a pipette tip and add it to a culture (if it is still frozen), or add 10-20 µL of the liquid glycerol stock to the culture (if it is somewhat thawed). AVOID THAWING GLYCEROL STOCKS COMPLETELY – take what you need and quickly return them to the -80 °C freezer. If they do thaw completely, they can be re-frozen, but repeated thawing may reduce the number of live cells in the stock tube.

IV. DNA Extraction

DNA Extractions are used to generate large amounts of plasmid DNA from E. coli cell cultures. This DNA can then be used for restriction digesting, PCR reactions, gel electrophoresis, or further transformations. By using a kit (Qiagen Qiaprep Spin Miniprep Kit) we are able to effectively remove most of the protein and RNA from a solution, leaving us with very clean DNA solutions.

1. Grow up 5 mL culture overnight in LB (6 mL if you want glycerol stock – 1 mL will be used for the frozen stock, which should be removed before pelleting the cells in the next step)

2. Pellet 5-10 minutes in centrifuge at 3000-3500 rpm

3. Pour off supernatant

a. At this step, you can freeze the cell pellet if you don’t have time to finish the rest of the procedure. The pellet is fine for a week or two in the -20 °C freezer

4. Check P1 solution for the checked RNase added box

5. Add 250 µL solution P1 to the 10 mL culture tube

6. Suspend the pellet in the P1 by pipetting up and down

7. Transfer suspended pellet to a 1.5 mL tube

8. Add 250 µL solution P2 to the tube

9. Mix by inverting the tube by hand 10-20 times. Solution should turn blue throughout, if not, continue inverting until blue throughout

10. Allow lysis to occur for 3-4 minutes (no more than 5 minutes)

11. Add 350 µL solution N3 to the tube

12. Mix by inverting tube by hand 10-20 times. Solution should lose all blue color, if not, continue inverting until all blue is gone.

13. Centrifuge at 13,000 rpm for 10 minutes (keep the hinge out to get the pellet to form correctly)

14. Using a pipette, remove the supernatant from the tubes, and apply to a labeled blue spin column from the kit

15. Centrifuge at 13,000 rpm for 1 minute

16. Pour flow-through BACK into the column and centrifuge at 13,000 rpm for 1 minute 17. Discard flow through

18. Add 750 µL PE solution to tube and centrifuge at 13,000 rpm for 1 minute

19. Discard flowthrough and centrifuge again at 13,000 rpm for 1 minute

20. Transfer blue column to a fresh 1.5 mL tube (labeled)

21. Add 30-50 µL ddH2O (depending on how concentrated you want your final DNA). Buffer EB (supplied in the kit) can also be used.

22. Incubate on benchtop for 10-15 minutes

23. Centrifuge at 13,000 rpm for 1 minute and discard column

24. Nanodrop to determine DNA concentration

V. Restriction Digest

All restriction digests were carried out using Fermentas reagents and restriction enzymes. The Fermentas FastDigest® (FD) reagents were used.

In a tube the following was added:

4 µg DNA

4µl 10x FD buffer

1 µl FD restriction enzyme 1

1 µl FD restriction enzyme 2

Volume brought up to 40µl total with nuclease free water

Tube was placed at 37°C for 1hr.

The FastDigest® Green Buffer was also used. When it was used the DNA could directly be loaded onto an agarose gel.

VI. Gel Making

Agarose gels are useful for DNA purification and analysis of DNA sizes. The gels are made up of an agarose matrix composed of long strands of agarose, and gaps of various sizes between the strands. The larger the DNA molecule, the longer it takes to fit through the gaps, making its progress through a gel slower than a small DNA molecule. The DNA is drawn through the gel using an electric current; the negatively charged phosphates on the DNA backbone being attracted toward the cathode (“Run towards red” is a helpful mnemonic as the cathode is generally red colored). By varying the concentration of the agarose gel, it is also possible to increase the separation of bands of certain sizes on the gel. A 1% agarose gel is generally used as it provides separation of bands from 200 bp to 3000 bp. For separating larger bands, a 0.7% gel is typically used and the smaller DNA fragments are run completely off the bottom of the gel. For separating smaller bands, a 2% or 1.5% gel can be used, and run normally. Gels are useful for purifying DNA bands of a particular size from restriction digests (to prevent multiple products from forming during ligations) and also for removing proteins from a DNA sample (such as restriction enzymes that are not inactivated by heat). Gel purification has the downside of losing some DNA, and reducing overall DNA concentration (a 120 bp band of DNA in a 2000 bp plasmid will only give .06 µg of DNA if 1 µg of total DNA digest is added to the lane). For small band sizes (< 200 bp), it may be necessary to use CIP or TAP dephosphorylation and ligation using the digested DNA solution without gel purification.

1. Determine the number of lanes you wish to run. Always plan for 1-2 lanes of the DNA ladder (2 especially if this gel will be cut up and DNA removed from it), or another suitable control. Most lanes can hold 20-25 µL of sample, so larger samples may need to be run on two lanes, or use the larger lane combs (40-50 µL capacity). The small gel box can hold 6 lanes of large capacity or 10 lanes of smaller capacity. The larger gel box can hold 12 lanes of large capacity or 20 lanes of small capacity. The large gel box is also capable of having 2 combs at a time (the second placed ½ down the box), and so its capacity can double at the cost of distance over which it can separate bands.

2. Once your gel box is selected, determine the concentration of gel you wish to make (see description for details). The concentration is the mass of agarose/mL of gel x 100%. The small gel box supports gels of 50 mL (potentially up to 75 mL, but 50 is easier to use) and the large gel box supports gels of 200 mL.

3. Set up the gel box by removing the gel tray. Make sure the rubber seals are still in their grooves. Apply a small amount of water to the inside of the side walls of the gel box and to the ends of the gel tray that have the rubber seals. Slide the gel tray into the gel box so that the open ends of the tray are against the box walls, and so that the rubber seals have not rolled up out of their grooves (if the seals moved, return them to their grooves and try sliding the tray in again).

4. Add the correct gel combs to the gel. The main comb for both gels goes into the first notch on the gel tray (should be 1-2 cm from an end), the secondary comb for the large gel is placed in the notch in the middle of the tray. The small gel combs have two sides (one thinner than the other) the thinner side has about 2/3 of the capacity volume of the thick side (which has the capacities listed in #1), so choose what you need.

5. In a flask that can hold at least 4x your gel’s volume, add the correct volume of 1x TAE buffer. DO NOT ADD WATER – the gel will not work correctly.

6. Weigh out the correct mass of agarose (NOT AGAR) and add it to the flask.

7. Microwave the solution until it boils. There are two stages of boiling – where small white bubbles form (frothy) and where large clear bubbles form. You want to let it boil a bit past the frothy stage and into the clear bubble stage. These bubbles will pop naturally, and will keep you from having a bubble filled gel. It is necessary when using higher gel concentrations (and recommended for all other concentrations) that the microwaving occur in 30 second increments, with the solution being stirred by GENTLE rotation (wear protective heat gloves) after each 30 second period, to ensure proper agarose distribution.

8. After microwaving add Ethidium Bromide to the gel solution. WARNING – carcinogen, glove use is advised (if you get it on yourself, wash your skin with water for 5 minutes – its very water soluble). Add 1 µL of Ethidium bromide for EACH 10 mL of gel volume (5 µL for small gel, 20 µL for large gel).

9. Mix Ethidium Bromide into the solution by GENTLE swirling (to avoid bubbles).

10. The gel solution can be allowed to cool slightly before pouring into the gel box (pouring boiling solution into gel box can cause it to warp and bend over time).

11. Gently pour the gel into the gel tray by leaning it on the gel box wall farthest from the top comb, and slowly tipping it into the box. This prevents bubbles from forming in the gel, and if they do form, forms them near the bottom of the gel.

12. If additional bubbles form in the gel box, while the gel is still liquid take a pipette tip and push the bubbles to the bottom edge of the gel, where they won’t interfere with DNA movement.

13. Allow gel to cool for 40 minutes to 1 hour. Test solidification by gently pressing on the bottom corner of the gel with a finger, it should feel solid and gel-like (not liquid).

VII. Gel Purification

This procedure was carried out using the Qiagen QIAquick Gel Extraction Kit.

VIII. Ligation

Ligation reactions are used to combine two linear fragments of DNA into a circular plasmid. The ligation procedure can be modified based on how much backbone DNA you have and how much insert you have, as well as if a phosphatase such as CIP (calf intestinal phosphatase) or TAP (thermo-sensitive alkaline phosphatase) was used in preparing the linear DNA molecules.

Example protocol for ligation (to be added to a PCR tube)

10μl Insert DNA

3μl Vector DNA

2μl 10X ligation buffer

34μl H2O

1μl T4 DNA ligase

Note: This protocol can be adjusted depending on the concentration of insert and vector.

IV. PCR

Add the following reagents to a tube (50 μl reaction) in the following volumes and order:

32μl sterile H2O

5 μl 10X buffer

2μl dNTP Mix

3μl MgCl2

6μl cells/DNA

0.25μl Taq Polymerase

1μl Primer 1

1μl Primer 2

The thermocyler is setup beforehand with the desired protocol. Typically: 94˚C for denaturing, 50-60˚C for primer annealing, and 72˚C for polymerase extending.

Example setup:

Step 1. 94˚C 2min 1x

Step 2. 94˚C 45sec

Step 3. 55˚C 45 sec

Step 4. 72˚C 1min 15 sec

Step 5. 72˚C 5min 1x

Step 6. 4˚C indefinitely

Repeat Steps 2 and 4 35x

V. Luminescence Protocol

Reagents:

http://www.promega.com/products/reporter-assays-and-transfection/reporter-assays/dual_luciferase-reporter-assay-system/

1X Phosphate Buffered Saline solution (PBS)

1X Passive Lysis Buffer (PLB)

1X LARII

1X Stop & Glo

Luminometer :

http://www.promega.com/products/instruments/luminometers/glomax-20_20-luminometer/

Luminometer software protocol should be set for 10 second integration.

1. Innoculate overnight cultures, incubate for 12 to 16 hours.

2. Add 0.5ml of culture to 1.5 ml centrifuge tube.

3. Pellet cells for 5 minutes at 10,000 RCF.

4. Discard supernatant.

5. Suspend cell pellet in 0.5 ml 1X PBS by vortex.

6. Pellet cells for 5 minutes at 10,000 RCF.

7. Discard supernatant.

8. Suspend cell pellet in 0.5 ml 1X PLB by pipette.

9. Agitate cells on shaker table at room temperature for 15 minutes.

10. Add 50 ul of LARII to a new 1.5 ml centrifuge tube.

11. Place 50 ul tube in luminometer, add 10 ul of lysed cells. Pipette up and down several times to ensure mixing.

12. Record first luminometer measurement.

13. Add 50 ul of Stop & Glo reagent to tube, vortex for 1 second.

14. Record second luminometer measurement.