Team:UChicago/Protocols

General Lab Best Practices

Labeling

Label tubes at all time before storage. Label clearly with:

1. Contents in the tube
2. Date
3. Your name/Initials
4. Number

Notes on Sterile Techniques

• Turn the flame on before opening anything sterile.

• Aliquot everything you use to prevent contamination. Before pouring aliquots, flame the bottle rim quickly to keep sterile. Make new aliquots in new tubes with stock when running low.

• Unscrew but do not remove all caps to make it easier to work with in the moment.

• When using growth media, flame the bottle rim quickly and then aliquot by pouring into 50ml conical tube. Never stick anything into bottles of growth medium, always pour.

• Flame bottle rim again and close bottle of growth medium.

• Take what you need from 50ml tube, keep sterile so it can be used repeatedly.

Lab Notebook Best Practices

Every lab notebook entry should contain…

1. Title: what technique you’re using, what you’re using it on (refer to specific tubes*)
2. Reference to protocol: are you using the standard protocol found in the masterlist?
3. Changes in protocol: have you done anything different from the standard protocol?
4. Conditions: did you run at 120V? for how long?
5. Mistakes: in case an experiment doesn’t work as a result of a mistake
6. Conclusion: was your experiment successful? why not? what’s next? what did you expect to see?
7. Your name: so others can ask questions to the right person. This is one reason why it’s critical to label tubes with the date, description, and your name

Best practices from Thomson’s [http://www.iphandbook.org/handbook/chPDFs/ch08/ipHandbook-Ch%2008%2002%20Thomson%20Laboratory%20Notebooks.pdf How to Start–and Keep–a Laboratory Notebook: Policy and Practical Guidelines] (see link and scroll down for examples):

“Although you may think you will remember what you did and why you did a certain experiment in a week’s time, YOU WILL NOT! And nor will anyone else in your laboratory. Hence the need for laboratory notebooks. In short, a laboratory notebooks is:

• a daily record of every experiment you do, think of doing, or plan to do
• a daily record of your thoughts about each experiment and the results thereof
• a record that would enable successive scientists, working on the same project, to pick up where you left off or reproduce your results”

“What goes into a laboratory notebook?

• a detailed account of every planned and executed experiment with the amount of detail that would enable a skilled scientist to determine what had been done, why it had been done, and what the results were
• dates accompanying every entry, account, or record
• [Links to outside resources used, formulas used for any calculations. -Alice]
• explanations of the significance of each experiment, as well as the observations, results and conclusions of the experiment
• details of each experiment (Remember, what may seem trivial or obvious at the time your experiment was conducted, may later be of critical importance.)
• personal comments (It is a living document, so stamp it with your own personality. Comments such as “SUCCESS AT LAST!! THIRD TIME LUCKY :)” are highly appropriate.
• photographs, computer generated data, and so forth should all be stuck into your notebook. a good photo matters!
• cross-references [For example, if you are starting a new experiment on 8/21 entry and are using the same protocol as already described in the Binder of Lab Practices, write on 8/21 entry, “following the protocol as described in the Binder of Lab Practices” -Alice]"

“A laboratory notebook is an important tool that goes well beyond research management, and keeping good records has implications for issues ranging from intellectual property management to the prevention of fraud.”

Steps to 3A Assembly & Labeling Guidelines

If from BioBrick in distribution kit plates (is an upstream or downstream part?):

1. resuspend plasmid DNA from kit plates
   • tube label e.g. lid: 18E plate 3, side: Pveg [date] [name]
2. transform plasmid DNA into DH5a cells (why those cells?)
   • plate label e.g.

   bottom of agar plate: [resistance] Pveg 18E3 [date] [name] [number if plating >1]

3. make overnight culture of a few isolated colonies
   • 15ml O.N. tube label e.g.

   side: [resistance] Pveg 18E3 [date] [name] [number if >1]

4. miniprep overnight culture
   • tube label e.g. lid: 18E3 mp [#], side: Pveg [date] [name] [concentration]ng/ul
5. Measure the concentration of DNA using the nanodrop and write concentration on side of tube and in lab notebook, return 1ul to tube // if you have time, run 2ul on gel to estimate concentration based on band size.
6. digest some of the miniprep
   • tube label e.g. lid: 18E3 dig [E, S] + [X, P], side: Pveg [date] [name]
7. run 2ul of the digest on a 1% gel, take photo and label
8. if digestion successful, then it’s ready to ligate.

If you see label rubbing off, re-label it! Always use permanent marker.

Resuspending DNA from iGEM Kit Plates

Protocol for resuspending DNA from the iGEM kit plates:

1. Poke through the aluminum foil on the well from which you want to take DNA with a 10ul tip with 10ul of nuclease-free water. Don’t take the foil off, it might cause cross-contamination between the wells.
2. Add 10 µL of dH2O. The water will turn red. Pipette in and out a few times to mix. Leave for 5 mins to make sure all the DNA dissolves.
3. DNA is ready to use for transformation. Use 2 µL for transformation. Acc. to the iGEM website, the concentration is high enough to use for transformation.

Recipes

Making Agar Plates

for 12 plates of 2 different antibiotics

Weigh out:

• 5.0 g tryptone
• 2.5 g yeast extract
• 5.0 g NaCl
• 7.5 g agar

1. Add 250 mL of dH2O to a graduated cylinder.
2. Mix powder well to bring into solution.
3. Add dH2O to total volume of 500 mL and transfer to 1 L flask. All powder except agar should be dissolved.
4. Put on stirring hot plate and heat to boil for 1 min while stirring.
5. Autoclave at liquid setting for 15 minutes in a basin with some water making sure to loosen top.
6. Let cool to point you can pick up with gloves comfortably and split between 2 500ml jars, each containing 250ml LB-agar with flame on.
7. Let agar cool to ~55C (you should be able to pick up the jar without a glove) and add antibiotics

Pouring the Plates

1. Remove sterile Petri dishes from plastic bag (save the bag for storage)
2. Pour a thin layer (5mm) of LB Agar (~10mL) into each plate being careful to not lift the cover off excessively (you should be able to just open up enough to pour).
3. Swirl plate in a circular motion to distribute agar on bottom completely.
4. Let each plate cool until its solid (~20 minutes) then flip so as to avoid condensation on the agar.
5. Store plates in plastic bags in fridge with: date and contents (note any additive).

Making LB (500mL)

Weight out:

• 5g tryptone
• 5g NaCl
• 2.5g yeast extract

1. Pour 250mL dH2O into 1L bottle.
2. Add ingredients and shake to mix
3. Pour 250mL dH2O to make total of 500mL
4. Autoclave on liquid setting

Agar Stab Protocols

1. Incubate agar stab from iGEM HG
2. With gloved hand, take sterile P1000 pipette tip and dip in the deep agar stab.
3. Streak on plate.
4. Streak again using another sterile pipette tip to spread bacteria over the plate.

Ligation Protocol

from [http://www.addgene.org/plasmid_protocols/DNA_ligation/ AddGene]:

1. Add amounts from highest to lowest (so water first). Flick to mix and pulse spin for 3 seconds.
2. Incubate at room temperature overnight.
3. If you’re not going to transform right away, heat inactivate tube at 65C for 10min.

Make controls if you can:

Transformation Procedures

TOP10 heat shock transformation procedure:

1. Let the cells thaw on ice.
2. Add your plasmid to the competent cells. Flick gently using index finger alone to mix. Do NOT vortex cells.
3. Leave on ice for 20 mins.
4. Place on the 42ºC heat block for 1 minute to heat shock.
5. Immediately place on ice for 2 minutes.
6. Add 1 mL 2X YT or SOC media.
7. Incubate in the 37ºC shaker for 1 hour. (We have no shaker, so we left it in a heat plate).
8. Centrifuge at 8 rcf for 30 seconds. Decant all but ~100 µL supernatant and resuspend.
9. Plate onto chloramphenicol (or antibiotic of choice) plates and use a sterile P1000 pipette tip to gently spread by hand while wearing gloves (no pipetman). Do not heat pipette tip. It’s okay if the tip leaves marks on the agar, but be as gentle as possible.

Protocol for competent cells in PCR tubes:

1. Let the cells thaw on ice.
2. Add your plasmid (~2ul) to the competent cells.
3. Leave on ice for 10 mins. [30min if you’re doing a ligation]
4. Heat shock at 42 degrees for 45 seconds

   a. Set the thermal cycler (PCR Machine) to 42 degrees

   b. Carry tubes in ice bucket to thermal cycler.

   c. Place tube inside for 45 seconds to heat shock.

5. Immediately put on ice for 2 minutes.
6. Add 1 mL SOC (or LB) media.
7. Incubate in the 37ºC shaker for 1 hour.
8. Centrifuge at 8 rpm for 30 seconds (pulse spin). Decant all but ~100 µL supernatant and resuspend.
9. Plate onto antibiotic of choice plates and use sterile, disposable plastic L-spreader to spread. Do not push on agar, just gently skim surface, it shouldn’t leave marks. If no spreaders are available, use a sterile P1000 pipette tip to gently spread by hand while wearing gloves (no pipetman). Do not heat pipette tip. It’s okay if the tip leaves marks on the agar, but be as gentle as possible.

B. subtilis Electroporation Protocol

Protocol from “High osmolarity improves the electro-transformation efficiency of the gram-positive bacteria Bacillus subtilis and Bacillus licheniformis” (Xue et al. 1999):

1. Cool cells in 50ml Falcon tubes in ice-water, 10min
2. Spin down 3,000rpm for 5min (Nora’s lab)
3. Resuspend in 650ul electroporation medium and transfer to 1.5ml tubes
4. Spin down 5,000xg for 5min (BSLC lab)
5. Wash 3x with electroporation medium (500ul)
6. Resuspend in 1/40 of culture volume of electroporation medium

Reusing Electroporation Cuvettes

• we need 1mm or 2mm (we have protocols for both)

Adapted from:

[http://www.protocol-online.org/biology-forums/posts/322.html http://www.protocol-online.org/biology-forums/posts/322.html]

[http://bitesizebio.com/articles/re-cycling-electroporation-cuvettes/ http://bitesizebio.com/articles/re-cycling-electroporation-cuvettes/]

• Focus on the metal parts of the cuvettes – clean them of DNA and cells and dry again quickly to prevent corrosion.
• Wash and dry as soon as possible after transformation.

1. Rinse the cuvette five times with purified water. Ensure, especially with 1-2mm gap cuvettes that the chamber is fully washed out in all washing steps.
2. Fill the cuvette with 0.2M HCl and allow to stand for 10 minutes (but no longer as this will promote corrosion).
3. Rinse the cuvette five times with 70% ethanol, again ensuring the chambers are fully washed.
4. Under a sterile hood if possible, decant the ethanol and use a pipette (or syringe and hypodermic needle for small gaps) to remove as much residual ethanol as possible.
5. Air dry for 60 minutes, then replace the cap.

…and now your cuvettes are ready to go again.

Making Overnight Cultures

1. Using Kron lab stick™, select a single colony from your [http://www.addgene.org/recipient_instructions/streak_plate/ streaked LB agar plate].
2. Drop the Kron lab stick™ into the liquid LB + antibiotic, swirl, and carefully break off the tip so it fits in the 15ml tube.
3. Loosely cover the culture with cap that is not air tight.
4. Incubate bacterial culture at 37oC for 12-18hr in a shaking incubator.

Miniprep Protocols

Note whether you are using E. coli or B. subtilis with Thermo Kit

(FYI: The protocol is in the kit box)

Day 1: Make O.N. culture: USE STERILE TECHNIQUE

1. Use the aliquoted LB already 3ml in 15ml tubes; make more aliquots if needed.
2. Add appropriate amount of antibiotic:

   1.5ul cam (stock 50mg/ml), 3ul kan (stock 50mg/ml), 3ul amp (stock 100mg/ml)

3. Select single, well isolated colony on plate (do not pick satellite colonies-small colonies clustered around a larger colony) and pick it with sterile P1000 pipette .
4. Using sterile technique, drop pipette in 15ml tube and vortex.

Day 2: Protocol from Thermo kit:

1. Retrieve overnight culture and apply to a microcentrifuge tube. Aliquot 30ul of Elution Buffer into eppendorf tube and warm to 50C on heating block.
2. Add 1mL at a time. Centrifuge the tube at 12,000rpm for 1 minute and dump out the supernatant. Then, apply another 1 mL or however much is left in tube from the overnight culture and repeat until all of the overnight sample has been pelleted.

   B. subtilis ONLY: Made 1mg/ml of lysozyme in resuspension buffer from stock (50mg/ml)

   a. Stock solution is in fridge at 50mg/ml

   b. Vortex stock solution.

   c. Put 245ul Resuspension Buffer in tube with pellet.

   d. Add 5ul of lysozyme from stock solution (50mg/ml) to reach final concentration of 1mg/ml

3. Resuspend completely the pelleted cells in 250 uL of Resuspension Buffer (with 1mg/ml lysozyme if B. subtilis culture) and pipet up and down until no cell clumps remain
4. Add 250 uL of the Lysis Solution and mix thoroughly by inverting the tube 4-6 times until the solution becomes viscous and slightly clear

   (BE SURE TO NOT VORTEX OR INCUBATE FOR OVER 5 MINUTES)

5. Add 350 uL of the Neutralization Solution and mix immediately and thoroughly by inverting the tube 4-6 times

   (THIS ENSURES COMPLETE PRECIPITATION OF BACTERIAL CELL DEBRIS)

6. Centrifuge sample for 5 minutes at 12,000rpm to pellet cell debris and chromosomal DNA
7. Transfer supernatant to the labelled translucent blue columns with labelled collecting tube

   (AVOID PIPETTING OR DISTURBING THE WHITE PRECIPITATE)

8. Centrifuge the sample in the column for 1 minute at 12,000rpm. Discard the flow-through and place the column back into the same collection tube
9. Add 500 uL of the Wash Solution to the column. Centrifuge for 1 minute at 12,000rpm and discard the flow-through. Place the column back into the same collection tube.
10. Repeat the wash step in step 9. using 500 uL of the Wash Solution
11. Discard the flow-through and centrifuge for an additional 1 minute at 12,000rpm to remove residual Wash Solution.
12. Discard the collection tube and place the column into a fresh 1.5 mL microcentrifuge tube.
13. Add 15 uL of the Elution Buffer to the center of the column membrane to elute the DNA. Be sure not to contact the membrane with the pipette tip. Incubate the column for 2 minutes at room temperature and centrifuge for 2 minutes. Repeat with remaining 15ul of elution buffer.
14. Nanodrop concentration (nucleic acid setting, blank with dH2O) in W110 GCIS.
15. Place the tube and column in our beaker and store the purified DNA at -20˚C

Making a 1% agarose gel for gel purification

1. Weigh out 0.5 g agarose.
2. Place in an EM flask.
3. Add 50 mL of 1X TAE.
4. Microwave for 1 minute.
5. Allow to cool for a bit. Should still be hot, but not too hot to touch.
6. Add 2.5 µL EtBr.
7. Pour out onto the cast with the large combs. Allow to solidify.

Note: It’s okay to reuse agarose gels, we just need to run the sample off. The EtBr will need to be re-added to the TAE buffer in the apparatus each time.The gels can be reused for a few weeks.

EtBr BATH: Made 100ml stock, used empty tip top to hold the bath. Remember to cover in aluminum foil. 5ul EtBr/100ml dH2O. Shake by hand gently for 15min, box wrapped in aluminum foil. Pour bath into aluminum-covered container and wash gel with ddis H2O. Shake gently again for 15min, covered with foil to protect EtBr in gel.

Enzymatic digestion

Total Volume: 15 uL

• 500ng DNA
• 1.5ul 2.1 buffer
• 0.5ul AflII
• 0.5ul NdeI
• add nuclease-free H2O for a total volume of 15 uL

If digesting overnight, place in heat block in 37C incubator for even heat

Note:

• If performing a double digest, use NEB's double digest finder to find the most compatible buffer for both enzymes.
• Enzymes used will differ depending on the biobrick part (see here)
• It may be necessary to heat inactivate your enzymatic reactions depending on if the enzymes have star activity (check here).

Using FastDigest Enzymes:

[http://www.thermoscientificbio.com/uploadedFiles/Resources/fast-digestion-dna.pdf Protocol]

[http://www.thermoscientificbio.com/restriction-enzymes/fastdigest-aflii-bspti/ AflII]

[http://www.thermoscientificbio.com/restriction-enzymes/fastdigest-ndei/ NdeI]

Gel extraction with the QIAGEN kit

1. Run the DNA from your digest in a 1 % agarose gel with large wells.
2. Excise the DNA from the gel by cutting with a blade and place into a microfuge tube.
3. Weigh the gel slice. Add 3 volumes of Buffer QG for every volume of gel. (100 µg ~ 100 µL)
4. Incubate at 50ºC for 10 mins (or until gel slice has dissolved). Invert/vertex as necessary to aid dissolution.
5. Add 1 volume isopropanol to the sample and mix.
6. Place the sample into the QIAGEN spin column and centrifuge for 1 min. at 13,000 rpm. If the volume is greater than 750 µL, simply spin in multiple increments of 750 µL or less. Discard flow-through after each spin.
7. Add 500 µL of Buffer QG and spin for 1 min at 13,000 rpm. Discard flow-through.
8. Wash by adding 750 µL Buffer PE and spinning at 13,000 rpm for 1 min. Discard flow-through.
9. Spin again at maximum speed to get rid of residual ethanol.
10. Place the column into a clean microfuge tube.
11. To elute, heat Buffer EB or water to about 50ºC. Put 50 µL of Buffer EB onto the center of the membrane, and let sit for 1-4 mins. Spin 1 min @ 13,000 rpm. (Can use a smaller amount of EB, e.g. 20 µL, to achieve higher DNA concentrations).
12. Nanodrop to check the concentration. DNA is now ready to use for ligation.

Tips for gel extraction

• Remember to heat up elution buffer to 50C before use and let the binding buffer sit on the column for a few minutes, let the EtOH evaporate off too by keeping the cap open.
• If you have low yield, try using smaller columns and eluting twice with warm elution buffer. Take the total volume you want and divide it in half. For the first round of elution, let the EB sit for 10min and then spin down. And then do it again. The silica in the spin column 50 tubes is too large compared to the volume of elution going through, so DNA tends to get struck.
• Gel purification does not work with 2% agarose gels