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Protocol for LB medium and LB agar plates

Written by Katariina Jaenes

INTRODUCTION

In order for bacteria to be successfully cultured, they must be grown in the appropriate media. LB, also known as Lysogeny broth, is a nutrient rich broth that is a standard for culturing Escherichia coli, as it allows for quick growth and high yields. Therefore the proper preparation of LB will be crucial to maintaining our bacterial stock throughout the summer. Furthermore, addition of agar to LB broth creates a gel for bacteria to grow upon, and is therefore used for plating bacterial cultures on petri dishes. We will be preparing our own LB as well as our own LB agar plates, which will be used to culture E.coli K-12.

BASIC TERMINOLOGY AND CONCEPTS

Agar vs. Agarose: Agar is used for making petri plates to culture organisms, while agarose is used for making gels, in the likes of SDS-PAGE and gel electrophoresis

SAFETY PRECAUTIONS

SDS (safety data sheet): Refer to the SDS sheets for all listed materials before entering the lab. Be prepared to answer any questions regarding the information on these sheets.

PPE (Personal protective equipment): Proper lab attire should be worn throughout the experiment: This means that upon entering the lab you should be wearing long pants and close-toed shoes. Contact lenses should not be worn. Furthermore, a lab coat, goggles, and gloves should be worn at all times, and long hair should be tied back.

Autoclave: The autoclave should only be handled by execs. Note that any autoclaved materials may still be hot and should therefore be handled with caution. Be careful not to burn yourself.

MATERIALS AND EQUIPMENT

Note: this recipe is for 500ml of LB, which makes approximately 18 plates. LB can be kept for 3-4 months

Reagents

• 5g NaCl
• 5g Tryptone
• 2.5g Yeast extract
• 7.5g of agar This is only necessary if you are making LB agar plates
• 500ml of dH20 (distilled water)

Materials

• 1L pyrex bottle
• 1L graduated cylinder
• Filter paper and scupula
• Stack of sterile plates (this protocol makes approx 25.)
• Bunsen Burner/Ethanol burner
• 70% EtOH wash bottle
• paper towels/wipes

LAB PROTOCOL

Part 1. Making the LB broth

This step can be carried out at a regular lab bench.

1. Obtain a clean 1L pyrex bottle
2. Obtain a graduated cylinder with 500ml of dH20 (distilled water) and add to the bottle. Record the amount added.
3. Using filter paper, separately measure out 5g of NaCl, 5 g of Tryptone, and 2.5g of yeast extract on a scale and add them to the bottle. Swirl the bottle in a circular motion to mix. Remember to recalibrate your scales in between measurements.
4. If you are making LB agar plates, weigh and add 7.5g of agar and swirl to mix. Record the amount added. Note that the contents do not necessarily need to be completely in solution before autoclaving.

Part 2. Autoclaving *ONLY DONE BY EXECS*

1. Lightly seal the top of the beaker with aluminium foil, and label the beaker with autoclave tape. Include LB (agar) – your name – date - iGEM. Unless you have been trained to use the autoclave, you will not be conducting the following steps 2 - 5.
2. Use appropriate transportation protocols to bring the LB bottle into the autoclave room. Remember to store the beaker in an autoclavable basin, in case of spills.
3. Check the water level on the autoclave, if necessary. Autoclave on the liquid setting for approximately 20mins.
4. The contents of the beaker will be hot after autoclaving, therefore take necessary measures to prevent burns. After autoclaving, allow the LB media too cool to 55oC before handling.
5. The LB broth can be stored in sterile conditions at room temperature, and should be good for a 3-4 months. Flame the lip of the bottle each time the LB is used. If the LB is contaminated it will appear cloudy. If the LB contains antibiotics: store in a -4C freezer - (However it is not recommended to store LB with antibiotics as the antibiotic will degrade over time)

Part 3. Pouring the plates (if you are making LB agar)

While pouring the plates it is crucial to maintain a sterile environment. This should be done in room WB303, with a sterile environment provided by a lit bunsen burner.

Note: While you are waiting for the autoclave, steps 1- 3 can be done in the meantime, in addition to the clean up from Part 1.

1. Sterilize the workspace with 70% EtOH before depositing your materials. Light the bunsen burner.
2. Obtain a stack/roll of empty plates. The plates should still be in their plastic sleeve/wrapping, as they should be sterile. Don't throw out the wrapping as it can be used to store the plates. It is essential that you minimize any chance of contaminating the plates. Make sure that you open the package at the top, and expose the plates as minimally as possible. Note that this protocol makes approximately 25 plates.
3. Once you take the plates out, store them upside down on your lab bench. Label the plates with your name – iGEM 2015 – date prepared – designated stripe* (if are using antibiotic). Once labelled, you may stack the plates to free up workspace.
4. Allow the LB media to cool before pouring. The LB will start to settle at about 30oC.
5. If you are preparing selective media, add antibiotic to the mixture. Use the recommended antibiotic concentrations (see table below) Swirl the flask in a circular motion to mix. If you don't know whether or not you are preparing selective media, ask.

Antibiotic	Recommended Concentration
Chloramphenicol (CAM)	25ug/mL
Ampicillin (AMP)	100ug/mL

One stripe along the sides corresponds to CAM, two stripes corresponds to AMP

1. Take an empty plate and open it slightly. You do not need to open it all the way to pour the agar. When pouring the agar, pour until 2/3 of the plate has been covered, or approximately half of the plate has been filled when viewed from the side. Pour the agar slowly, to prevent the formation of bubbles. Swirl the plate in a circular motion to distribute the media evenly on the plate.
2. After pouring, set the plates to cool in stacks of 4-5, as this takes up less space. Don't stack the plates too high – we want to prevent spills.
3. Rinse the pyrex bottle with water before the remnants solidify and become hard to remove.
4. Allow the plates to cool for at least 20 minutes until the agar has solidified. Flip the plates to prevent condensation forming on the agar. The plates can then be stacked and stored in plastic bags (ideally re-use the plastic bags that the plates come in.)
5. The LB agar plates should be stored in a 4oC freezer, should be good for 1-2 months

LEAVING THE LAB

Prior to leaving the lab, you should:

1. Clean dirty glassware, or at least set aside the glassware to be cleaned by a designated individual.
2. Wipe down your workspace.
3. Ensure that all materials have been returned to their places, and that the plates have been properly stored in the fridge.

REFERENCES

Addgene: Protocol – Making LB Agar Plates for Bacteria. From: https://www.addgene.org/plasmid- protocols/bacterial-plates/

Department of Ecology and Evolutionary Biology, UCLA. Making LB Agar Plates. From https://www.eeb.ucla.edu/Faculty/Barber/Web%20Protocols/LB%20Agar%20Plates.pdf

U of T iGEM Luria Broth (LB) protocol

Sezonov G, Joseleau-Petit D, D’Ari R. Escherichia coli Physiology in Luria-Bertani Broth . Journal of Bacteriology. 2007;189(23):8746-8749. doi:10.1128/JB.01368-07.

Protocol for SOC medium

Written by Katariina Jaenes

INTRODUCTION

SOC is a variant of the rich media SOB (super optimal broth) with catabolite repression. This means that glucose is supplemented in the media, allowing for optimal metabolic conditions for the bacteria. SOC increases the transformation efficiency of cells, as it provides ample nutrients to cells that have recently undergone stress as result of having been made competent. Accordingly, it will be used in bacterial transformation to stabilize the cells and to increase transformation yields. Since SOC is high in nutrients, it is more easily contaminated than LB or King's B media.

SAFETY PRECAUTIONS

SDS (safety data sheet): Refer to the SDS sheets for all listed materials before entering the lab. Be prepared to answer any questions regarding the information on these sheets.

PPE (Personal protective equipment): Proper lab attire should be worn throughout the experiment: This means that upon entering the lab you should be wearing long pants and close-toed shoes. Contact lenses should not be worn. Furthermore, a lab coat, goggles, and gloves should be worn at all times, and long hair should be tied back.

Autoclave: The autoclave should only be handled by execs. Note that any autoclaved materials may still be hot and should therefore be handled with caution. Be careful not to burn yourself.

MATERIALS AND EQUIPMENT

Note: this recipe is for 500mL of SOC.

Reagents

• 1.802g glucose
• 10g tryptone
• 2.5g Yeast extract
• 0.584 g NaCl
• 0.093g KCl
• _g MgCl2 (anhydrous)
• 1.234g MgSO4 .7H20
• 500ml of dH20 (distilled water)

Materials

• 2 x 1L pyrex bottle (must have cap)
• Note: A smaller pyrex bottle may be used to accommodate the glucose solution.
• 1L graduated cylinder
• weighing boats and scupula
• 70% EtOH wash bottle
• paper towels/wipes

LAB PROTOCOL

Part 1. Making the SOC broth

This step can be carried out at a regular lab bench.

1. Obtain two 1L mL pyrex bottles. Ensure that the bottles can be sealed with a cap - this will help prevent contamination and enable long term storage.
2. Obtain a graduated cylinder with 500ml of dH20 (distilled water).
3. Using filter paper, separately measure out 10g tryptone, 2.5g yeast extract, 0.292g NaCl, 0.093g KCl, _g MgCl2 anhydrate, 1.234g MgSO4, heptahydrate. on a scale and add them to the 1L bottle. Add 400mL dH20. Swirl the flask in a circular motion to mix. Remember to recalibrate your scales in between measurements.
4. In the separate bottle, and measure out 1.802g glucose on a scale and add the rest of the 100mL dH20. Recalibrate your scales in between measurements.

* The 1L autoclaved separately, as the contents will react if autoclaved together.

Part 2. Autoclaving ONLY DONE BY EXECS

1. Lightly seal the top of the bottles with aluminium foil, or unscrew the caps. Label both with autoclave tape. Include a label with SOC – your name – date - iGEM 2015. Unless you have been trained to use the autoclave, you will not be conducting the following steps 2 - 5.
2. Use appropriate transportation protocols to bring the bottles into the autoclave room. Remember to store the beaker in an autoclavable basin, in case of spills.
3. Check the water level on the autoclave, if necessary. Autoclave on the liquid setting for approximately 20mins.
4. The contents of the bottles will be hot after autoclaving, therefore take necessary measures to prevent burns. After autoclaving, allow the media too cool to 55oC before handling.

Part 3

This part should be done in room WB303, in sterile conditions close to a bunsen burner.

1. In a sterile environment, slowly add the autoclaved 1.802g glucose and dH20 to the beaker containing the autoclaved _g MgCl2 and 1.234g MgSO4, 10g tryptone, 2.5g yeast extract, 0.292g NaCl, 0.093g Kcl and dH20. Flame the lip of the bottle before transferring the contents. Swirl to mix, and seal tightly to prevent contamination. Flame the cap before sealing.
2. The SOC broth can be stored in sterile conditions at room temperature, and should be good for a 3-4 months. Flame the lip of the bottle each time the SOC is used. SOC should be handled carefully, as it is especially prone to contamination.

LEAVING THE LAB

Prior to leaving the lab, you should:

1. Clean dirty glassware, or at least set aside the glassware to be cleaned by a designated individual.
2. Wipe down your workspace.
3. Ensure that all materials have been returned to their places, and that the plates have been properly stored in the fridge.

REFERENCES

Cold Spring Harbour Protocols. SOC Medium for E.coli. From: http://cshprotocols.cshlp.org/content/2012/6/pdb.rec069732.short

Sun QY et al. Culture of Escherichia coli in SOC medium improves the cloning efficiency of toxic protein genes. Anal Biochem. 2009: From http://www.ncbi.nlm.nih.gov/pubmed/19622338

Qiagen. Recipe for SOC Medium. From: https://www.qiagen.com/ca/resources/faq?id=d3be05d0- ec02-4ced-aabe-dcd2a1061920&lang=en

King's B Protocol

Written by Katariina Jaenes

INTRODUCTION

In order for bacteria to be successfully cultured, they must be grown in the appropriate media. King's B media, formulated by King et al,. Is a nonselective media that is a standard for culturing Pseudomonas species, as it allows for quick growth and high yields. Therefore, the proper preparation of King's B will be crucial to maintaining our bacterial stock throughout the summer. Furthermore, addition of agar to King's B creates a gel for bacteria to grow upon, and is therefore used for plating bacterial cultures on petri dishes. We will be preparing our own King's B as well as our own King's B agar plates, which will be used to culture Pseudomonas putida F1.

BASIC TERMINOLOGY AND CONCEPTS

Agar vs. Agarose: Agar is used for making petri plates to culture organisms, while agarose is used for making gels, in the likes of SDS-PAGE and gel electrophoresis

SAFETY PRECAUTIONS

SDS (safety data sheet): Refer to the SDS sheets for all listed materials before entering the lab. Be prepared to answer any questions regarding the information on these sheets.

PPE (Personal protective equipment): Proper lab attire should be worn throughout the experiment: This means that upon entering the lab you should be wearing long pants and close-toed shoes. Contact lenses should not be worn. Furthermore, a lab coat, goggles, and gloves should be worn at all times, and long hair should be tied back.

Autoclave: The autoclave should only be handled by execs. Note that any autoclaved materials may still be hot and should therefore be handled with caution. Be careful not to burn yourself.

MATERIALS AND EQUIPMENT

Note: this recipe is for 500ml of LB, which makes approximately 18 plates. King's B can be kept for 3-4 months

Reagents

• 5g proteose peptone
• 0.75g anhydrous K2HPO4
• 7.5g glycerol
• 1.6mL MgSO4 (of 1M solution )
• 500mL of dH20 (distilled water)
• 7.5g of agar This is only necessary if you are making King's B agar plates

Materials

• 2 x1L pyrex bottle ◦ Note: the MgSO4 solution may kept in a smaller bottle
• 70% EtOH wash bottle
• Filter paper and scupula
• Stack of sterile plates (this protocol makes approx 25.)
• paper towels/wipes
• Bunsen burner/Ethanol burner

LAB PROTOCOL

makes 500mL of King's B, should be good for 3-4 months

Part 1. Making King's B broth

This step can be carried out at a regular lab bench.

1. Obtain 2 clean 1L Erlenmeyer flask.
2. Obtain a graduated cylinder with around 500ml of dH20 (distilled water) and add it to one bottle. Record the amount added.
3. Using filter paper, separately measure out 5g of proteose peptone, 0.75 g anhydrous K2HPO4 7.5 g glycerol on a scale and add them to the bottle with 500mL dH20. Swirl the beaker in a circular motion to mix. Remember to recalibrate your scales in between measurements.
4. In a separate flask, add 12g of MgSO4 and 100mL of dH20. This will be the stock solution of 1M
5. If you are making agar plates, weigh and add 7.5g of agar to the 500mL solution, and swirl to mix. Record the amount added. Note that the contents do not necessarily need to be completely in solution before autoclaving.

Part 2. Autoclaving *ONLY DONE BY EXECS*

1. Lightly seal the top of the flask with aluminium foil, and label it with autoclave tape. Include King's B (agar) – your name – date - iGEM 2015. Unless you have been trained to use the autoclave, you will not be conducting the following steps 2 - 5.
2. Use appropriate transportation protocols to bring the flask into the autoclave room. Remember to store the beaker in an autoclavable basin, in case of spills.
3. Check the water level on the autoclave, if necessary. Autoclave on the liquid setting for approximately 20mins.
4. The contents of the beaker will be hot after autoclaving, therefore take necessary measures to prevent burns. After autoclaving, allow the media too cool to 55oC
5. After autoclaving, add 1.6 mL of sterile 1 M MgSO4 in sterile conditions. If it is added before autoclaving, the media will appear cloudy.
6. The broth can be stored in sterile conditions at room temperature, and should be good for a 3-4 months. Flame the lip of the bottle each time the media is used.

Part 3. Pouring the plates (if you are making agar plates)

While pouring the plates it is crucial to maintain a sterile environment. This should be done in WB303, in sterile conditions provided by a lit bunsen burner. Alternatively, can be done in 403 with an Ethanol burner

Note: While you are waiting for the autoclave, steps 1- 3 can be done in the meantime, in addition to the clean up from Part 1.

1. Sterilize the workspace with 70% EtOH before depositing your materials.
2. Obtain a stack/roll of empty plates. The plates should still be in their plastic sleeve/wrapping, as they should be sterile. Don't throw out the wrapping as it can be used to store the plates. It is essential that you minimize any chance of contaminating the plates. Make sure that you open the package at the top, and expose the plates as minimally as possible. Note that this protocol makes approximately 18 plates.
3. Once you take the plates out, store them on the lab bench and keep them upside down. Label the plates with King's B - your name – iGEM 2015 – date prepared
4. Make sure that the media has cooled down sufficiently before beginning. However, don't wait too long, as the media will start to settle at around 30C.
5. Take an empty plate and open it slightly. You do not need to open it all the way to pour the agar. When pouring the agar, pour until 2/3 of the plate has been covered, or approximately half of the plate has been filled when viewed from the side. Pour the agar slowly, to prevent the formation of bubbles. Swirl the plate in a circular motion to distribute the media evenly on the plate.
6. After pouring, set the plates to cool in stacks of 2 or 3, as this takes up less space. Don't stack the plates too high – we want to prevent spills.
7. Rinse the bottle with water before the remnants solidify and become hard to remove.
8. Allow the plates to cool for at least 20 minutes until the agar has solidified. Flip the plates to prevent condensation forming on the agar. The plates can then be stacked and stored in plastic bags (ideally re- use the plastic bags that the plates come in.)
9. The agar plates should be stored in a 4oC freezer, should be good for 1-2 months

LEAVING THE LAB

Prior to leaving the lab, you should:

1. Clean dirty glassware, or at least set aside the glassware to be cleaned by a designated individual.
2. Wipe down your workspace.
3. Ensure that all materials have been returned to their places, and that the plates have been properly stored in the fridge.

REFERENCES

From: Cold Springs Harbour Protocol. King's B Medium. From: http://cshprotocols.cshlp.org/content/2009/7/pdb.rec11326.full?text_only=true

Recipe for King's B medium. Received from Maggie Middleton, Lab technician at the David Guttman Lab.

Antibiotic Stock Preparation

Written by Dr. Tim Lee

Background

The preparation of antibiotic stock is a relatively simple series of mixing and dilutions. The antibiotic stocks are typically made from four antibiotics : Ampicillin, Kanamycin, Tetracycline and Chloramphenicol. With the exception of chloramphenicol, the other antibiotics are light sensitive and once stock is prepared they are wrapped in foil to reduce light exposure.

An antibiotic comes in a powder form as the basic stock and is mixed with milli-Q water to form a stock solution. Antibiotic stock preparation follows the same basic process for each antibiotic, varying only in concentration which is dependent on the type of antibiotic used.

Antibiotic stocks are prepared for use either in liquid or solid media. In the case of liquid media, the antibiotic stock is simply added and mixed in. For solid media, usually agar and LB, the media is sent for autoclaving, and while the media is still hot the antibiotic is added and mixed. This is to ensure that the antibiotic can be mixed evenly amongst the media before it solidifies into a solid state.

Antibiotics are used for the purpose of selection. In terms of synthetic biology, plasmids confer selective antibiotic resistance when successfully transformed into their target bacteria. Using liquid or solid growth media that has been treated with antibiotics provides a selection control for those bacteria that have successfully incorporated the plasmid.

Basic Terminology and Concepts

There are some terms which will be commonly used to describe the preparation of antibiotic stocks. Familiarity and understanding of these terms is key to comprehend the protocol.

Milli-Q- Water that has been purified through successive steps of filtration and deionization. The standard used in our lab is typically 18.2 MΩ·cm at 25 °C, measured in resistance due to the lack of ions. The filters used are 0.22 μm in size to ensure a high level of purity. This water is used for preparing our antibacterial stock to ensure purity.

LB – Lysogeny Broth, is a very standard and simple media to create due to its recipe consisting of 3 components - tryptone, yeast extract, and sodium chloride. This mix of anhydrous ingredients is added to water and then autoclaved, producing liquid media. The production of solid LB for plates is done by adding agar, a protein isolated from certain species of seaweed which coagulates the liquid into a gel like form when cooled. This is done prior to autoclaving and is poured into the plates while still hot, where it will cool into the plate shape.

Aliquot - An aliquot is a term to denote a certain quantity of something. In this case an aliquot of antibiotic stock would denote either 1mL or 0.5mL, depending on the antibiotic.

Antibiotic- Compounds which inhibit bacterial growth. They act either bacteriostatically by preventing reproduction of the bacteria, or bacteriocidally where they directly kill the bacteria. Generally bacteriocides work by interfering with the synthesis of peptidoglycan in the bacteria's cell walls. Tetracycline is an example of a bacteriostatic, where it acts by binding to the ribosomes of prokaryotic bacteria and inhibits translation.

Autoclave- A piece of equipment used for sterilization. The autoclave performs much like a pressure cooker: it subjects the contents inside it to a high temperature and high pressure steam bath. Usually the temperature is 121 Celsius and at 15lbs/in2 20 minutes is enough to kill most microorganisms and render equipment sterile. When adding antibiotics to media, it is done after autoclaving so that the heat does not destroy the antibiotic activity.

Safety Precautions

• Be careful when handling the antibiotic, and avoid contact with eyes, mouth and skin
• When pouring liquid, commit to the pour to prevent spills.
• Use proper PPE

Materials and Equipment

• 100mL Pyrex Bottles
• Magnetic Stir Bar
• Magnetic Stirrer
• Antistatic Weighing Boat
• Analytical Balance
• Milli-Q Water Dispenser
• 50mL Falcon Tubes
• Aluminum Foil
• 20mL Syringe
• 0.22 μm Filter
• 1mL Microcentrifuge tubes

Reagents

• 4g Amphicilin
• MilliQ water
• 400mg tetracycline
• 80mL 70% EtOH
• 2.72g chloramphenicol
• 80mL 100% EtOH

Procedure

Ampicillin

• Stocks and Usage Stock Concentration 50 mg/mL in milliQ water
• Aliquots 500μL (use a P1000 set to 0500)
• Working Concentration 50 μg/mL Preparation of 80mL stock solution

Ampicillin is kept in the 4°C fridge, and is light sensitive. To ensure your stock solution is not degraded, cover all microcentrifuge tubes used for storing the solution with foil.

1. Weigh 4g ampicillin onto an antistatic weighing boat.
2. Add 80mL milliQ water to a 100mL Pyrex bottle.
3. Add the ampicillin to the milliQ.
4. Place a small magnetic stir bar into the solution and place the Pyrex bottle on the stirrer. Set at 300-600 rpm and stir until dissolved.
5. Filter sterilise the solution into 50mL Falcon tubes using a 20mL syringe outfitted with a 0.22 μm filter.
6. Aliquot (500μL) into the appropriate microcentrifuge tubes, labelled with an "A" on top, and store in the Nalgene racks found in the 20oC.

Kanamycin

• Stocks and Usage Stock Concentration 10 mg/mL in milliQ water
• Aliquots 1 mL (use a P1000 set to 1000)
• Working Concentration 50 μg/mL
• Add 5 mL of stock per litre of LB. Preparation of 80mL stock solution

Kanamycin is kept in the 4°C fridge, and is light sensitive. To ensure your stock solution is not degraded, cover all microcentrifuge tubes used for storing the solution with foil.

1. Weigh 800mg kanamycin onto an antistatic weighing boat.
2. Add 80mL milliQ water to a 100mL Pyrex bottle.
3. Add the kanamycin to the milliQ.
4. Place a small magnetic stir bar into the solution and place the Pyrex bottle on the stirrer. Set at 300-600 rpm and stir until dissolved.
5. Filter sterilise the solution into 50mL Falcon tubes using a 20mL syringe outfitted with a 0.22 μm filter
6. Aliquot (1mL) into the appropriate microcentrifuge tubes, labelled with a "K" on top. Store in the Nalgene racks found in the 20oC. (Make sure microcentrifuge tubes are covered with foil)

Tetracycline

• Stocks and Usage Stock Concentration 5 mg/mL in 70% EtOH (N.B. 70% EtOH, not 100% EtOH!)
• Aliquots 1mL (use a P1000 set to 1000)
• Working Concentration 20 μg/mL Preparation of 80mL stock solution

Tetracycline is kept in the 4°C fridge, and is light sensitive. To ensure your stock solution is not degraded, cover all microcentrifuge tubes used for storing the solution with foil.

1. Weigh 400mg tetracycline onto an antistatic weighing boat.
2. Add 80mL 70% EtOH to a 100mL Pyrex bottle.
3. Add the tetracycline to the 70% EtOH.
4. Place a small magnetic stir bar into the solution and place the Pyrex bottle on the stirrer. Set at 300-600 rpm and stir until dissolved.
5. Recommended, but optional because of storage in 70% EtOH: Filter sterilise the solution into 50mL Falcon tubes using a 20mL syringe outfitted with a 0.22 μm filter.
6. Aliquot into the appropriate microcentrifuge tubes, labelled with a "T" on top. Store in the Nalgene racks found in the 20oC.

Chloramphenicol

• Stocks and Usage Stock Concentration 34 mg/mL in 100% EtOH (N.B. 100% EtOH, not 70%!)
• Aliquots 1 mL (P1000 set to 1000)
• Working Concentration 25μL/mL. Preparation of 80mL stock solution

Chloramphenicol is kept with the general chemicals, and is not light sensitive. The microcentrifuge tubes do not need to be covered with foil to store chloramphenicol.

1. Weigh 2.72g chloramphenicol onto an antistatic weighing boat.
2. Add 80mL 100% EtOH to a 100mL Pyrex bottle.
3. Add the chloramphenicol to the 100% EtOH.
4. Place a small magnetic stir bar into the solution and place the Pyrex bottle on the stirrer. Set at 300-600 rpm and stir until dissolved.
5. Aliquot (1mL) into the appropriate microcentrifuge tubes, labelled with a "C" on top. Store in the Nalgene racks found in the 20oC. Note: No filter sterilization is needed because it is stored in 100% EtOH.

Leaving the Lab

Prior to leaving the lab, you should:

• Clean dirty glassware, or at least set aside the glassware to be cleaned by a designated individual.
• Wipe down your workspace.
• Ensure that all materials have been returned to their places, and that the plates have been properly stored in the fridge.

Acknowledgement

Protocols of previous iGEM teams were used to make this guideline.

Gel Electrophoresis

Written by Seray Cicek

Part 1: Casting the gel

1. Add 0.5g of Agarose and 50ml of 1X TAE buffer to an Erlenmeyer flask. (Note that TAE buffer in the stock has 50X concentration and needs to be diluted with autoclaved Milli-Q water)
2. Place the flask into the microwave. After this step flask will be hot. Use heat gloves
3. Set the timer to 2 min. Start the microwave oven but stop every 30 sec to swirl the contents of the flask. This helps suspend the undissolved agarose. Continue until agarose particles are dissolved.
4. While hot, add 3ul of 10,000x SYBR Safe into the agarose solution and mix throughly.
5. Set the flask aside for cooling. Occasionally swirl to make the cooling even.
6. Set up the gel tray and make sure that there is no leak by testing it with dH20 and kimwipes.
7. When the flask is cool enough for you to handle easily, pour the solution into the tray.
8. Wait until the gel solidifies.

Part 2: Loading the gel

1. Make a note of what is going into each well and put that in your lab notebook.
2. Determine the DNA concentrations via Nanodrop. (Approximately 100ng DNA per band is required.)
3. Cut a piece of parafilm to use as a mixing surface. Alternatively can use PCR tubes.
4. Mix DNA with loading dye in a 1:1 ratio directly on the parafilm or in the PCR tubes using a pipette tip and gently aspirating. (i.e. 10uL of DNA with 2uL of 6x loading buffer) Mix by pipetting up and down. Avoid bubbles.
5. Carefully remove the comb of the gel.
6. Place the gel into the gel electrophoresis apparatus with its tray.
7. Add more 1X TAE buffer to cover the gel completely. Buffer should be 1-2mm above the gel. Make sure that all the wells are submerged.
8. Load 10uL of DNA ladder to the first well. Add 10uL of your sample DNA to corresponding wells. (Remember record which wells you load)
9. Put the lid on. The negative (-/black) electrode should be closer to the DNA wells as DNA migrates towards the positive (+/red) electrode. 10.Set the rig to 100V & 60 min. Hit start. 11.Make sure that your DNA does not run off the gel. The loading buffer enables you to track the DNA. 12.The gel apparatus will stop on its own when the time is up. Proceed to imaging the gel. It can sit in the rig overnight if need be.

References:

igem wiki: http://local.biochemistry.utoronto.ca/igem/index.php/Analytical_Gel_Electrophoresis

jove: http://www.jove.com/video/3923/agarose-gel-electrophoresis-for-the-separation-of-dna- fragments

open wetware: http://openwetware.org/wiki/Agarose_gel_electrophoresis

lifetechno: https://www.lifetechnologies.com/ca/en/home/life-science/pcr/elevate-pcr- research/agarose-content-with-tips-and-tricks.html

Syber green catalog: http://www.lifetechnologies.com/order/catalog/product/S33102

Syber green staining protocols: http://www.lumiprobe.com/protocols/sybr-green-gel-staining

Making Chemically Competent Cells

Protocol obtained from openwetware: http://openwetware.org/wiki/Preparing_chemically_competent_cells

Materials

• Plate of cells to be made competent
• TSS buffer
• LB media
• Ice

Glassware & Equipment

• Falcon tubes
• 500μl Eppendorf tubes, on ice
• 200ml conical flask
• 200μl pipetman or repeating pipettor
• 5ml pipette

Protocol

Part 1: TSS Buffer

To make 50 mL:

• 5g PEG 8000
• 1.5 mL 1M MgCl2 (or 0.30g MgCl2*6H20)
• 2.5 mL DMSO
• Add LB to 50 mL
• Filter sterilize (0.22 μm filter)

Part 2: Making Chemically Competent Cells

1. Grow a 5ml overnight culture of cells in LB media. In the morning, dilute this culture back into 25-50ml of fresh LB media in a 200ml conical flask. You should aim to dilute the overnight culture by at least 1/100.
2. Grow the diluted culture to an OD600 of 0.2 - 0.5. (You will get a very small pellet if you grow 25ml to OD600 0.2)
3. Put eppendorf tubes on ice now so that they are cold when cells are aliquoted into them later. If your culture is X ml, you will need X tubes. At this point you should also make sure that your TSS is being chilled (it should be stored at 4 fresh then put it in an ice bath).
4. Split the culture into two 50ml falcon tubes and incubate on ice for 10 min. All subsequent steps should be carried out at 4 wherever possible
5. Centrifuge for 10 minutes at 3000 rpm and 4
6. Remove supernatant. The cell pellets should be sufficiently solid that you can just pour off the supernatant if you are careful. Pipette out any remaining media.
7. Resuspend in chilled TSS buffer. The volume of TSS to use is 10% of the culture volume that you spun down. You may need to vortex gently to fully resuspend the culture, keep an eye out for small cell aggregates even after the pellet is completely off the wall.

8. Add 100 μl aliquots to your chilled eppendorfs and store at − 80oC.

   o Freeze the cells immediately using a dry ice bath.

   o If you run a control every time you clone (i.e. a vector-only ligation), you can as you can as well freeze cells in 200 μl aliquots. Unused cells can be frozen back once and reused, albeit with some loss of competence.

Lab protocol for bacterial transformation

Written by Matt D'Iorio

Introduction

In this protocol, you will be using chemically competent E coli that have been placed in a calcium chloride solution prior to freezing. The solution acts to neutralize the cells so that they don't repel each other. The bacteria and plasmid mixture will be chilled on an ice for 30 minutes. Placing the mixture in a 42 degrees Celsius water bath for 30 seconds will heat shock the mixture, which will cause the transformation. Once the LB is added and mixed with the transformed bacteria, it can be plated on the LB plate with an antibiotic. Since a lot of the bacteria will not be transformed after the heat shock, plating E. coli with an antibiotic will ensure only the transformed E. coli survive. Separate reagents from materials

Basic Terminology and concepts

Transformation – The process in which the genetic makeup of a cell is changed by introduction of DNA from the surrounding environment.

Competent E. coli – E. coli in a state where it can allow the uptake of DNA

Heat Shock – A sudden increase in temperature used to propel a plasmid into a bacterium

Safety precautions

This is a benign lab protocol, but don't forget that PPE is necessary at all times. The E. coli you will be working with will be non-pathogenic but it should still be handled properly, which means it should not come into contact with your skin or gloves. If you are removing materials from the -80 degrees Celsius freezer, do not use your bare hands or regular lab gloves, use gloves designated specifically for the freezer.

Reagents

• Competent E. coli cells
• 250 μL of SOC
• LB plate supplemented with antibiotics
• Materials and equipment
• Set of micropipettes and pipette tips
• 5 x1.5mL – microcentrifuge tube
• Ice bath
• Thermomixer
• Streaker/Glass beads

Lab protocol

1. Thaw competent E. coli on ice
   • Competent cells till be in the -80 degrees Celsius freezer
   • Do NOT thaw cells by hand, if they are warmed by hand they will no longer be competent
2. Mix the cells gently with a pipette tip (do not pipette up and down)
3. Aliquot 50μL of bacteria into a pre-chilled 1.5mL-microcentrifuge tube.
4. Add 1-2 μL of the plasmid to the bacterial sample, and mix gently with the pipette tip (don't pipette up and down)
5. Incubate the tube on ice for 30 minutes.
6. While the bacteria are on ice, ensure the water bath is set to 42°C
7. Put 250μL of LB in a microcentrifuge tube and place it in the 37°C incubator to warm up during this time
8. After the 30 minute incubation period, place the tube in the 42°C water bath (mixing OFF) for exactly 30 seconds (45 seconds for E.coli DH10 beta)
9. Place the tube on ice for two (2) minutes 10.Add 250μL of pre-warmed SOC
10. Shake in 37°C incubator, 300 rpm, for 1-2 hours
11. Spread 100μL of the transformed bacteria on an LB plate supplemented with the appropriate antibiotic
12. Label plate with name, date, and what it is (iGEM 2015 – Transformed E. Coli – Your Name)
13. Allow the plates to dry inverted, then place in the 37°C incubator O/N

Leaving the lab

• Clean dirty glassware, or at least set aside the glassware to be cleaned by a designated individual.
• Wipe down your workspace.
• Ensure that all materials have been returned to their places, and that the plates have been properly stored in the fridge.
• Dispose of all disposable tubes and pipette tips used in biohazard containers.
• Make sure your plates are labelled and put in a place they can be found.

Cell Lysis Protocol

Written by Kayla Nemr

In preparation have the desired cells grown out on plates, with single colonies.

Day 1

1. Create overnight cultures of each from a single colony in 5mL of LB + antibiotic at 37C and 300rpm

Day 2

1. label 250mL (or larger) baffled flasks, pre-autoclaved.
2. Add 50mL of LB + antibiotic
3. Add 500uL of starter culture to the corresponding flask.
4. Grow at 37C until the OD is approximately 0.1-0.6
5. Reduce the temperature to 30C (can go as low as 16C)
6. At the end of the day, add 1mM IPTG

Day 3 - Extraction of cell lysates

1. Pellet 25mL in 50mL falcon tubes at 5000g for 10 min at 4C.

2. Resuspend in 2mL of lysis buffer

   • 50mM tris-HCl pH 7.5
   • 50-200mM NaCl
   • 5% glycerol (v/v)
   • 1mM DTT

3. Add 20uL 100mM PMSF

4. Add lysozyme to final concentration of 300ug/mL
5. Incubate tubes on ice horizontally in ice buckets and place on orbiter (for 3 hours)
6. Transfer to 2mL eppendorf tubes
7. Pellet in microcentrifuge at 13000g for 10 minutes at 4C.

SDS-PAGE Protocol

Written by Kayla Nemr, with Seray Cicek, Sarah Bi, and Roy Lee

Introduction

Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) is a technique commonly used in molecular biology which allows for the separation of proteins depending on the size of each protein and helps identify the proteins that are being separated.

Please watch videos shown below to understand the protocol more in depth.

Making the gel: https://www.youtube.com/watch?v=EDi_n_0NiF4

Running the gel: https://www.youtube.com/watch?v=XUjLO-ek2C8

Staining the gel: https://www.youtube.com/watch?v=b-1dXzU4iOw

Basic Terminology and Concepts

Protein ladder: a premade solution containing proteins of known sizes. Analyzing the electrophoresis alongside the ladder makes it easier to identify the various sizes of the fragments in the tested protein samples by comparing them to the known protein sizes of the ladder.

Resolving gel: this section of the gel will be the section that separates the proteins based on weight.

Stacking gel: this section of the gel will be the section where the protein samples are loaded. This section does not separate the proteins based on weight and after the electrophoresis is complete, provides no information and may be thrown out.

Buffer: the buffer allows for current to run through the gel Materials and Equipment

Materials

• 6.04g of Tris base
• 28.8g of glycine
• 2g SDS
• ddH2O
• 1.5 M Tris, pH 8.8
• 1.0 M Tris, pH 6.8
• 30% bis-acrylamide
• 10% SDS
• 10% APS
• TEMED
• 30% Isopropanol
• Methanol
• Coomassie Brilliant Blue R250
• 100% ethanol
• Glacial acetic acid
• Protein samples
• Protein ladder
• SDS loading dye 6x

Equipment

• Micropipette and tips
• Glass plates
• Casting frame
• Casting stand
• Gel comb
• Tweezers/spatula
• Filter paper/nitrocellulose membrane
• Erlenmeyer flasks
• Electrophoresis apparatus (Gel apparatus/Cassette/Tank)
• Power supply
• Wide containers

Lab Protocol

Preparation of 1X SDS-PAGE Running Buffer.

This recipe makes 2L of 1X SDS-PAGE running buffer.

CAUTION - SDS powder is hazardous. Prepare solution in a ventilated fume hood.

• Dissolve 6.04g of Tris base and 28.8g of glycine together in 1.8 L of ddH2O.
• Add 2g SDS and mix.
• Add ddH2O to a final volume of 2 L.

Preparation of SDS-PAGE gel

• Insert glass plates (0.75 mm) into the casting frame and secure them into place. Make sure that the shorter
• plate is in front and that the bottom is flat.
• Put the casting frame on the casting stand.
• Insert the comb in between the glass plates. Take a marker and make a mark approximately 1
• cm beneath the comb. Take the comb out.
• If necessary, test the leakiness of the apparatus by adding water. Draw out the water using filter
• paper.
• Start making the resolving gel in a separate flask

The recipe for the resolving/separating gel (10mL, 12%) is as follows:

This will create 10 mL of gel. Around >3.5 mL is needed for each gel.TEMED must be added last and the gel should be casted immediately after its addition.

• 3.35mL of dH2O (milliQ)
• 2.5 mL of 1.5 M Tris, pH 6.8
• 100 uL of 10% SDS
• 4mL of 30% bis-acrylamide
• 50 uL of 10% Ammonium persulfate (APS)

• 5 uL of TEMED

  Add gel into the apparatus until it reaches the previously created mark

• Remove any air bubbles by adding a layer of isopropanol on top of the gel. Wait until the

• resolving gel has polymerized (usually takes 30-45 minutes) and remove the isopropanol by pouring it or by using filter paper.
• Start making stacking gel in a separate flask

The recipe for the stacking gel (10mL, 4%) is as follows:

This will create 10 mL of gel. Around 1.2 mL is needed for each gel. TEMED must be added last and the gel should be casted immediately after its addition.

• 7.3 mL of dH20
• 1.25 mL of 1.0 M Tris, pH 6.8
• 100uL of 10% SDS
• 1.34 mL of 30% bis-acrylamide
• 50uL of 10% APS
• 10uL of TEMED

• Fill the rest of the apparatus with stacking gel and place the gel comb in. Ensure that there are

  no air bubbles. Wait until the gel solidifies (around 30-45 minutes).

Running the SDS-PAGE gel

1. Take out the gel (with the plates) from the casting apparatus and place it in the gel apparatus.
2. Make sure that the short plate faces inwards to the apparatus. If only one gel is being run, make
3. sure to close the other side of the apparatus with a dummy plate.
4. Once the two plates are secured, insert them into the cassette and lock the apparatus in place.
5. Insert the cassette into the tank.
6. Prepare the samples. Add 3uL of 6x sample buffer to 15uL of sample, then boil the samples. This can be done by setting a thermocycler to 95C for 10 minutes.
7. Meanwhile, fill the inner chamber (in between the glass plates) with buffer until the buffer
8. reaches the top of the inner chamber. Fill the bottom of the outer chambers with more buffer
9. so that it covers the bottom of the gels by at least 2cm. A large amount is not needed. Take the
10. gel comb out gently. straighten any lopsided well walls with
11. pipette tip.
12. Add the protein ladder to the first well and the protein samples in the other
13. wells. Load 10-20ul per well (same volume for every well). Be careful not to overfill a well!
14. Record order of loading in lab notebook!
15. Dip the pipette tip through the surface of the buffer, centre it near the bottom of the well and
16. SLOWLY depress the pipette plunger into the well. Make sure all of the solution actually goes
17. into the well as to not contaminate the other wells.
18. Place the lid on the apparatus and connect the apparatus to the power supply.
19. Make sure the cathode and anode are connected correctly (black to black, red to red)
20. Set the voltage to 140 and run the gel for around 70 minutes.
21. Monitor movement of loading dye through gel. Stop running when dye front reaches bottom of
22. gel. Once the process is finished, take off the lid, remove the inner chamber and pour out the
23. buffer carefully.
24. Remove the glass carefully.
25. Rinse the plates with water and separate the plates
26. The stacking gel should be removed by cutting with a spatula or razor blade. Try not to tear the
27. resolving gel.

Staining the SDS-PAGE gel

1. Fill a wide container with deionized water and separate the resolving gel from the glass plate. Leave the
2. gel in the container.
3. Rinse the gel with deionized water gently. Repeat 2-3 times.
4. In a container with deionized water, microwave the gel for 30 seconds.
5. Transfer the gel to the staining solution and microwave for 30 seconds.

To make the staining solution:

• In a new container, add 2.5 g of Coomassie Brilliant Blue R250 to 450mL 100% ethanol. Dissolve
• the Brilliant Blue with a magnetic stir bar.
• Add 100 mL of glacial acetic acid
• Add 450 mL dH2O
• Optional: filter through Whatman No.1 filter
• Leave the gel on an orbital shaker for 20-60 minutes, until strong bands are visible.

To make the staining solution

• In a new container, add 800 mL of 100% ethanol and 400 mL of glacial acetic acid to 2800 mL of

  dH2O (destaining solution)

• Transfer the gel to the new destaining solution. Add kimwipes to aid in the destaining process.

Leaving the lab

• Ensure that the gels are disposed of in the proper waste bins
• Dispose of any excess buffer according to regulations
• Disconnect power supply and ensure that all machinery is turned off

References

Protocol for separating and stacking gels, sample preparation and gel staining obtained from Kayla Nemr.

Preparing gels: http://www.ruf.rice.edu/~bioslabs/studies/sds-page/gellab2a.html

Assembling, loading, running gels: http://www.ruf.rice.edu/~bioslabs/studies/sds-page/gellab2b.html

Jove: http://www.jove.com/science-education/5058/separating-protein-with-sds-page

PureLink® PCR Purification protocol

Minor changes were made to the protocol supplied with the Invitrogen PureLink® PCR Purification Kit

1. Add 4 volumes of PureLink ® Binding Buffer (B2) with isopropanol to 1 volume of the PCR product (50–100 μL). Mix well.

2. Remove a PureLink ® Spin Column in a Collection Tube from the package.

3. Add the sample with the appropriate Binding Buffer (from step 1 of this procedure) to the PureLink ® Spin Column.

4. Centrifuge the column at room temperature at 13,000 × g for 1 minute.

5. Discard the flow through and place the spin column into the collection tube.

6. Add 650 μL of Wash Buffer with ethanol to the column.

7. Centrifuge th e column at room temperature at 13,000 × g for 1 minute. Discard the flow through from the collection tube and place the column into the tube.

8. Centrifuge the column at maximum speed (15, 000 x g)at room temperature for 2–3 minutes to remove any residual Wash Buffer. Discard the collection tube.

9. Place the spin column in a clean 1.7-mL PureLink ® Elution Tube supplied with the kit.

10. Add 50 μL of Elution Bu ffer (10 mM Tris-HCl, pH 8.5) or sterile, distilled water (pH >7.0) to the center of the column.

11. Incubate the column at room temperature for 1 minute. Centrifuge the column at maximum speed for 2 minutes.

12. The elution tube contains the purified PCR product. Remove and discard the column. Store the purified PCR product at –20°C or use the PCR product for the desired downstream application.

References

PureLink® PCR Purification protocol. From: http://tools.thermofisher.com/content/sfs/manuals/purelink_pcr_man.pdf

Quantum Prep Plasmid Miniprep Protocol

Minor changes were made to the protocol supplied with the Bio-Rad Quantum Prep Plasmid Miniprep Kit

All centrifugation steps are performed at maximum speed (12,000–14,000 x g) unless otherwise stated.

1. Transfer an overnight culture (1–2 ml) of plasmid-containing cells to a microcentrifuge tube. Pellet the cells by centrifugation for 15–30 secs. If the overnight culture is larger than 2mL, centrifuge the falcon tubes at 3000g for 10 minutes and proceed. Remove all of the supernatant by pipeting.

2. Add 200 μl of the cell resuspension solution and vortex until the cell pellet is completely resuspended.

3. Add 250 μl of the cell lysis solution and mix by gently inverting the capped tube about ten times (do not vortex). The solution should become viscous and slightly clear if cell lysis has occurred.

4. Add 250 μl of the neutralization solution and mix by gently inverting the capped tube about ten times (do not vortex). A visible precipitate should form.

5. Pellet the cell debris for 5 mins in a microcentrifuge. (If used falcon tubes centrifuge for the same amount of time.) A compact white debris pellet will form along the side or at the bottom of the tube. The supernatant (cleared lysate) at this step contains the plasmid DNA.

6. While waiting for the centrifugation step at step 5, insert a spin filter into one of the 2 ml microcentrifuge wash tubes supplied with the kit. Mix the Quantum Prep matrix by vortexing or repeated shaking and inversion of the bottle to insure that it is completely suspended.

7. Transfer the cleared lysate (supernatant) from step 5 to a spin filter. Split the total volume (700uL) into two, adding 350uL of lysate into each filter. Add 100 μl of thoroughly suspended Quantum Prep matrix to each, then pipet up and down to mix. If you have multiple samples, transfer the lysates first, then add matrix and mix. When matrix has been added to all samples and mixed, centrifuge for 30 secs.

8. Remove the spin filter from the 2 ml tube, discard the filtrate at the bottom of the tube, and replace the spin filter in the same tube. Add 250 μl of wash buffer and wash the matrix by centrifugation for 30 seconds.

9. Remove the spin filter from the 2 ml tube, discard the filtrate at the bottom of the tube and replace the spin filter in the same tube. Add 250 μl of wash buffer and wash the matrix by centrifugation for a full 2 mins to remove residual traces of ethanol.

10. Remove the spin filter and discard the microcentrifuge tube. Place the spin filter in one of the 1.5 ml collection tubes supplied with the kit or in any standard 1.5 ml microcentrifuge tube that will accomodate the spin filter. Add 50 μl of nuclease free H 2 O or TE. Elute the DNA by centrifugation for 1 min at top speed.

1. Discard the spin filter and store the eluted DNA at -20°. Make sure to label properly.

References

Bio-Rad Quantum Prep Plasmid Miniprep Kit Instruction Manual. From: http://www.bio-rad.com/webroot/web/pdf/lsr/literature/MS4100066F.pdf