Creating Sterile Agar Plates

Goal

• Goal: To create sterile agar plates using one of the four methods below.

Pre-Protocol Questions

1. Do you know how to make liquid media?
2. Do you have enough media ingredients?
3. Do you know how to use the pH meter?
4. Do you have enough empty glass bottles in which you can pour the liquid media?

Refer to this link for general guidance on making media: http://microbes.sites.haverford.edu/LaboratoryWiki/Media_Recipes

Version 1: Copied from ABRC's Seed Handling FAQ for Seed Handling

Citation: ABRC Seed Handling FAQ PDF [1], accessed Sept. 23, 2018.

1. Add 4.31 g of Murashige and Skoog (MS) basal salt mixture and 0.5 g of 2-(N-Morpholino) ethanesulfonic acid (MES) to a beaker containing 0.8 L of distilled water and stir to dissolve. Add distilled water to final volume of 1 L. Check and adjust pH to 5.7 using 1M KOH and pH meter.
2. Divide the media into two 1 L glass bottles, 500 mL in each. Add 5 g of agar granulated per bottle. Keep the lid loose.
3. Autoclave for 20 min at 121°C, 15 psi with a magnetic stirring device in the bottle.
4. Place the bottles on a stir plate at low speed, and allow the agar medium to cool to 45-50°C (until the container can be held with bare hands).
5. Starting from this step, perform all the steps in sterile conditions in a laminar flow hood. Add (optional) 1-2% sucrose and 1 mL Gamborg’s Vitamin Solution, stirring to evenly dissolve. Optional sucrose and vitamins should be added after autoclaving and only after the agar media cools, because vitamins are thermo-labile and 15-25% of the sucrose may be hydrolyzed to glucose and fructose at elevated temperatures. Plants grow more vigorously and quickly on media containing 1-2% of sucrose, however, fungal and bacterial contamination must be rigorously avoided by seed sterilization. Note that germination of some mutants might be delayed on sucrose-containing media.
6. Label the bottom of Petri plates with identification number or name, including the date.
7. Pour enough media into plates to cover approximately half of the depth of the plate.
8. Allow the plates to cool at room temperature for about an hour to allow the agar to solidify. If the plates are not to be used immediately, wrap them in plastic and store at 4°C (refrigerator temperature). Covered plates, boxes, or tubes with solidified agar can be stored for several weeks at 4°C in a container that prevents desiccation.
   1. NOTE: Our lab will likely be using Magnenta GA-7 plant culture boxes rather than petri dishes - if any changes to this protocol are necessary due to this difference, it will be addressed in Version 2 of this protocol.

Version 2

Note: This protocol version was set up to document the way that we first made MS media on 10/24/2018. It will be modified as we learn more about how MS media actually works! This protocol uses Magenta GA-7 3x3x4" boxes, not petri dishes.

All parts of this protocol are to be followed in accordance to the rules of a Biosafety Level 2 lab - lab coat and gloves are worn at all times.

1. Fill a 1 Liter granulated cylinder with 400 mL of MilliQ water - gently drop magnetic stir bar into the cylinder and put the cylinder on a magnetic plate.
2. Open the pouch of MS media powder and pour into the cylinder. Slowly turn on the magnetic plate and allow the media to dissolve. Use a small squirt bottle of MilliQ water to wash some of the leftover MS powder off the sides of the cylinder. (Note: The amount of powder is not written on the pouch, so we will weigh the powder the next time we make media. Based on previous papers, it is likely around 4 g.)
3. Fill the cylinder to 950 mL
4. Adjust pH to 5.7±0.1 using the 5M NaOH and pH meter (caution: very strong base)
5. Add water up to 1000ml with a squirt bottle.
6. Pour solution into clean 2 L erlenmeyer flask.
7. Weigh out 8 g of agar - add to the solution. This will create a 0.8% concentration of agar in the final MS media. An improved agar-plate method for studying root growth and response of Arabidopsis thaliana (Agar ID: BP1432-500, from Fischer BioReagents)
8. Cover with tin foil and use autoclave tape to secure it to the flask.
9. Place flask in the "To Be Autoclaved" section. As the agar will begin cooling as soon as it is in the autoclave, ask Nicole to put the flask in the waterbath at 55C so that the solution stays liquid.
10. To create individual grow boxes containing MS media, first sterilize the hood by generously spraying with 70% ethanol and wiping down with a paper towel. Do the same with a styrofoam lid and place in the hood.
11. Coat gloves with 70% ethanol and allow to dry. Transfer the autoclaved grow boxes that are to be filled with MS media into the hood. Transfer the autoclaved MS media into the hood and rest on the styrofoam lid.
12. Using a serological pipet, transfer as much media as desired into individual boxes and close the lids back when full. Do not move the boxes as they are solidifying.
13. Once solidified, place boxes on their plastic trays and move to their next location.

Version 3

Note: this protocol documents our decision to increase agar concentration from 0.8% to 1%, and also illustrates how to add cycloheximide to the final sterilized media.

All parts of this protocol are to be followed in accordance with the rules of a Biosafety Level 2 lab - lab coat and gloves are worn at all times. Be especially careful with the solution of cycloheximide and autoclave any instruments that come into contact with it.

1. Fill a 1 Liter granulated cylinder with 400 mL of MilliQ water - gently drop magnetic stir bar into the cylinder and put the cylinder on a magnetic plate.
2. Open the pouch of MS media powder and pour into the cylinder. Slowly turn on the magnetic plate and allow the media to dissolve. Use a small squirt bottle of MilliQ water to wash some of the leftover MS powder off the sides of the cylinder. (Note: The amount of powder is not written on the pouch, so we will weigh the powder the next time we make media. Based on previous papers, it is likely around 4 g.)
3. Fill the cylinder to 950 mL
4. Adjust pH to 5.7±0.1 using the 5M NaOH and pH meter (caution: very strong base)
5. Add water up to 1000ml with a squirt bottle.
6. Pour solution into clean 2 L erlenmeyer flask.
7. Weigh out 10 g of agar - add to the solution. This will create a 1% concentration of agar in the final MS media. An improved agar-plate method for studying root growth and response of Arabidopsis thaliana (Agar ID: BP1432-500, from Fischer BioReagents)
8. Cover with tin foil and use autoclave tape to secure it to the flask.
9. Place flask in the "To Be Autoclaved" section. As the agar will begin cooling as soon as it is in the autoclave, ask Nicole to put the flask in the waterbath so that the solution stays liquid.
10. To create individual grow boxes containing MS media, first sterilize the hood by generously spraying with 70% ethanol and wiping down with a paper towel. Do the same with a styrofoam lid and place in the hood.
11. Coat gloves with 70% ethanol and allow to dry. Transfer the autoclaved grow boxes that are to be filled with MS media into the hood. Transfer the flask of autoclaved MS media into the hood and rest on the styrofoam lid.
    1. To create media that contains cycloheximide, transfer the stock solution of cycloheximide (100mg/mL) into the flask of media in order to dilute 1000x; swirl to mix. For example, add 500 uL of cycloheximide to 500 mL of media.
12. Using a serological pipet, transfer 50 mL media to the individual boxes and close the lids back when full. Do not move the boxes as they are solidifying.
13. Once solidified, place boxes on their plastic trays and move to their next location.

Version 4

Note: We returned to this protocol version after testing a 1% agarose concentration for plant media and finding that it is too dense for Arabidopsis roots to penetrate.

All parts of this protocol are to be followed in accordance to the rules of a Biosafety Level 2 lab - lab coat and gloves are worn at all times.

1. Fill a 1 Liter granulated cylinder with 400 mL of MilliQ water - gently drop magnetic stir bar into the cylinder and put the cylinder on a magnetic plate.
2. Open the pouch of MS media powder and pour into the cylinder. Slowly turn on the magnetic plate and allow the media to dissolve. Use a small squirt bottle of MilliQ water to wash some of the leftover MS powder off the sides of the cylinder. (Note: The amount of powder is not written on the pouch, so we will weigh the powder the next time we make media. Based on previous papers, it is likely around 4 g.)
3. Fill the cylinder to 950 mL
4. Adjust pH to 5.7±0.1 using the 5M NaOH and pH meter (caution: very strong base)
5. Add water up to 1000ml with a squirt bottle.
6. Pour solution into clean 2 L erlenmeyer flask.
7. Weigh out 8 g of agar - add to the solution. This will create a 0.8% concentration of agar in the final MS media.

An improved agar-plate method for studying root growth and response of Arabidopsis thaliana (Agar ID: BP1432-500, from Fischer BioReagents)

1. Cover with tin foil and use autoclave tape to secure it to the flask.
2. Place flask in the "To Be Autoclaved" section. As the agar will begin cooling as soon as it is in the autoclave, ask Nicole to put the flask in the waterbath at 55C so that the solution stays liquid.
3. To create individual grow boxes containing MS media, first sterilize the hood by generously spraying with 70% ethanol and wiping down with a paper towel. Do the same with a styrofoam lid and place in the hood.
4. Coat gloves with 70% ethanol and allow to dry. Transfer the autoclaved grow boxes that are to be filled with MS media into the hood. Transfer the autoclaved MS media into the hood and rest on the styrofoam lid.
5. Using a serological pipet, transfer as much media as desired into individual boxes and close the lids back when full. Do not move the boxes as they are solidifying.
6. Once solidified, place boxes on their plastic trays and move to their next location.

Note for future research: Given that grow-boxes with vented lids lead to the desiccation of the MS media over time, further research should be conducted to determine how much media is necessary to sustain A. thaliana growth in grow-boxes with vented lids.