Experiments

Introduction:

This protocol is for the creation of LB-ampicillin plates for use in cloning with E. coli.

Materials:

  • LB-agar powder
  • Double-distilled water (ddH2O)
  • Wooden sticks
  • Aluminum foil
  • Ampicillin
  • Culture plates/petri dishes

Procedure:

  1. Measure 17-20 g of LB-Agar powder on a scale, and add it to a glass container.
  2. Measure 500 mL of ddH2O, and add it to the same container.
  3. Stir this mixture gently until homogenous.
  4. Place aluminum foil over the lid of the container and label with autoclave tape.
  5. Place the container in the autoclave for 20-30 minutes.
  6. Remove the container from the autoclave and let the mixture cool until it reaches 55 ˚C.
  7. Add ampicillin to the container (100 µg/mL) and stir gently for 30 seconds.
  8. Pour a thin layer (about 20 mL) of the LB-ampicillin mixture into culture plates.
  9. Let the plates cool for one hour, or until solidified.
  10. Store the plates inverted overnight.
  11. Label the plates with the correct antibiotic and store in plastic bags at 4 ˚C.

Introduction:

Tris-Acetate-Phosphate (TAP) is a rich medium used for the suspension of cultures of Chlamydomonas reinhardtii. This protocol makes for 1 liter of 1x TAP media.

Materials:

  • 25 mL of 40x Beij
  • 25 mL of 40x TAP
  • 1 mL of 1000x trace elements
    • EDTA-Na2, 25 mM
    • (NH4)6Mo7O24, 28.5 µM
    • Na2SeO3, 0.1 mM
    • Zn·EDTA, 2.5 mM
    • Mn·EDTA, 6 mM
    • Fe·EDTA, 22 mM
    • Cu·EDTA, 2 mM
  • DI water
  • Autoclave

Procedure:

  1. Combine all reagents and top up to 1 liter of DI water.
  2. Autoclave for 30 minutes.
  3. Allow to cool before use.

Introduction:

TAP-paromomycin is a rich medium used for the growth of colonies of Chlamydomonas reinhardtii.

Materials:

  • 1x TAP media
  • 10 µg/mL Paromomycin
  • 15 g/L Agar powder
  • Sterile plates

Procedure:

  1. Follow the “TAP Media” protocol but add 15 g/L of agar before autoclaving.
  2. Add 10 µg/mL of Paromomycin after autocalving 1 liter of TAP and letting cool sufficiently for about 30 minutes.
  3. Pour warm TAP+Paromomycin in a thin layer enough to cover the whole surface of each plate.
  4. Allow media to solidify before keeping in 4C until use.

Introduction:

Miniprep kit is used to elute DNA from a sample. The handbook dictates that this protocol is designed for the purification of up to 20 μg of high-copy plasmid DNA from 1–5 ml overnight cultures of Escherichia coli in LB medium, but we are using 3mL specifically.

Materials:

  • Miniprep kit (QIAprep Spin Miniprep Kit (250) PN: 27106)
  • LB amp bacterial plate
  • LB amp liquid media
  • 37°C incubator
  • 37°C shaker incubator
  • Microcentrifuge
  • Vortex mixer
  • 100% ethanol
  • Microcentrifuge tubes 2ml sterile
  • Microcentrifuge tubes 1.5ml sterile (DNase free)
  • Pipettes (1000 uL and 200 μL) and tips

Procedure:

Streak out bacterial plate (Day 1)

  1. Streak out bacteria on LB amp plate using the pattern below with three sterile pipette tips:

Start overnight (O/N) culture (Day 2)

  1. Label culture tube with plasmid names.
  2. Add 6 ml of LB amp media.
  3. Pick a single colony with a pipette tip from a freshly streaked selective plate and drop the pipette tip into a correspondingly labeled tube.
  4. Incubate for 12–16h/overnight at 37°C on a shaking incubator.

Collect cell pellet (Day 3)

  1. Pellet 1.5 ml bacterial culture (not to exceed 15 OD units) by centrifugation at 16,000 x g (~13,000 RPM) for 30 seconds. Discard supernatants.

Qiagen Miniprep Kit

Done once per kit:

NOTE: Store Plasmid Neutralization Buffer (B3) at 4°C after opening, as it contains RNase A.

NOTE: Check if precipitate has formed in Lysis Buffer (B2), incubate at 30–37°C, inverting periodically to dissolve.

  1. Resuspend pellet in 200 μl Plasmid Resuspension Buffer (B1) (pink). Vortex or pipet to ensure cells are completely resuspended. There should be no visible clumps.
  2. Lyse cells by adding 200 μl Plasmid Lysis Buffer (B2) (blue/green). Invert the tube immediately and gently 5–6 times until the color changes to dark pink and the solution is clear and viscous. Do not vortex! Incubate for one minute or less.
  3. Neutralize the lysate by adding 400 μl of Plasmid Neutralization Buffer (B3) (yellow). Gently invert the tube until color is uniformly yellow and a precipitate forms. Do not vortex! Incubate for 2 minutes.
  4. Clarify the lysate by spinning for 5 minutes at 16,000 x g.
  5. Carefully transfer supernatant to the spin column and centrifuge for 1 minute. Discard flow-through.
  6. Re-insert column in the collection tube and add 200 μl of Plasmid Wash Buffer 1. Plasmid Wash Buffer 1 removes RNA, protein and endotoxin. (Add a 5 minute incubation step before centrifugation if the DNA will be used in transfection.) Centrifuge for 1 minute.
  7. Add 400 μl of Plasmid Wash Buffer 2 and centrifuge for 1 minute.
  8. Transfer column to a clean 1.5 ml microfuge tube. Use care to ensure that the tip of the column has not come into contact with the flow-through. If there is any doubt, re-spin the column for 1 minute before inserting it into the clean microfuge tube.
  9. Add ≥ 30 μl DNA Elution Buffer to the center of the matrix. Wait for 1 minute, then spin for 1 minute to elute DNA.

Introduction:

This protocol is intended for use with the Type IIS restriction enzyme SapI. Golden Gate Assembly is a method of molecular cloning that can easily combine multiple fragments of DNA inserts. Using Type IIS restriction enzymes, the resulting product is "scarless" and does not retain the recognition sites, making it essentially irreversible. This protocol was taken from the DTU Denmark 2019 iGEM team.

Materials:

  • T4 DNA Ligase
  • 10X T4 DNA ligase buffer
  • SapI Enzyme
  • Reciever Plasmid
  • DNA Fragment with SapI Overhangs
  • ddH2O
  • PCR Tubes

Procedure:

Reaction Set-Up

  1. Add the following reagents to a PCR tube:
    • 0.5 uL of T4 DNA ligase
    • 2 uL of 10X T4 DNA ligase buffer
    • 0.5 uL of SapI enzyme
    • 100 ng of receiver plasmid
    • Equimolar amounts of inserts
    • ddH2O for a total volume of 20 uL
  2. Mix the contents of the PCR tube gently by flicking.
  3. Place the tube in a thermocycler.

Thermocycler Program

Step Temperature Time
1. Activation of Sapl 37°C 5 minutes
2: Activation of T4 ligase 16°C 5 minutes
Repeat steps 1 & 2 for 25 cycles
3: Inactivation of SapI 65°C 20 minutes
4: Inactivation of T4 ligase 85°C 10 minutes
5: Hold 4°C Hold
  1. Cleanup any tubes or micropipette tips by disposing in autoclave safe disposable bags and autoclave according to autoclave brand guidelines but cycle time must be set for a minimum of 30 minutes @ 121C, 15 psi.

Introduction:

Escherichia coli (E. coli) is an optimal host for transformation of DNA into cells. Its growth allows for high expression of desired DNA or genes of interest. The following protocol is from New England BioLabs® High Efficiency Transformation Protocol using NEB® 10-beta Competent E. coli (High Efficiency) (C3019H/C3019I) protocol.

Materials:

  • NEB 10-beta Competent E. coli cells
  • SOC media
  • LB+ampR plates

Procedure:

  1. Thaw a tube of NEB 10-beta Competent E. coli cells on ice for 10 minutes.
  2. Add 2 µl containing 1 pg-100 ng of plasmid DNA to the cell mixture (25ml). Carefully flick the tube 4-5 times to mix cells and DNA. Do not vortex.
  3. Place the mixture on ice for 30 minutes. Do not mix.
  4. Heat shock at exactly 42°C for exactly 30 seconds. Do not mix.
  5. Place on ice for 5 minutes. Do not mix.
  6. Pipette 950 µl of room temperature SOC into the mixture.
  7. Place at 37°C for 60 minutes. Shake vigorously (250 rpm) or rotate.
  8. Warm selection plates to 37°C.
  9. Mix the cells thoroughly by flicking the tube and inverting, then perform several 10-fold serial dilutions in NEB 10-beta/Stable Outgrowth Medium.
  10. Spread 50-100 µl of each dilution onto a selection plate and incubate overnight at 37°C. Alternatively, incubate at 30°C for 24-36 hours or 25°C for 48 hours.

Introduction:

This protocol delineates the design of the polymerase reaction and thermal cycles for the production of many copies of amplified DNA fragments.

Materials:

  • Forward/Upstream Primer
  • Reverse/Downstream Primer
  • Template DNA
  • Polymerase Master Mix
  • Nuclease Free Water

Procedure:

PCR Reaction Mix

  1. Determine size of region of interest
    1. Identify gene of interest and record sequence in Benchling including any modifications (i.e., terminator sequence)
    2. Determine primer size and location including any modifications (i.e., homology regions or restriction sites)
    3. Add the gene of interest fragment size and primer fragment sizes together to estimate total length (or just use benchling tool)
  1. Pick a polymerase; Taq polymerase for non-cloning amplification, or high fidelity polymerase for cloning
    1. Different polymerases have different temperature ranges and times for the reaction.
  1. Set up reaction mixture in PCR tube with template DNA, upstream and downstream primers, polymerase master mix and nuclease free water
    1. Ex. Taq 2x Quick Load PCR Components
Component 25 µl reaction 50 µl reaction Final Conc.
10 µM Forward Primer 0.5 µl 1 µl 0.2 µM (0.05–1 µM)
10 µM Reverse Primer 0.5 µl 1 µl 0.2 µM (0.05–1 µM)
Template DNA variable variable < 1,000 ng
Taq 2X Master Mix 12.5 µl 25 µl 1X
Nuclease-free water to 25 µl to 50 µl

PCR Reaction Mix

  1. Calculate annealing temperature and extension times and determine thermal cycles
    1. Determine GC content and primer concentration to calculate annealing temperature (https://tmcalculator.neb.com/#!/main)
    2. For extension time, add approximately 30-60 seconds of time per kilobase in amplified region of interest
    3. For all other times, use higher recommended time
    4. Ex. Taq 2x thermal cycler conditions
Step Temp Time
Initial Denaturation 95°C 3 minutes
30 cycles 95°C (Denature) 15-30 seconds 15-60 seconds 1 minutes per kb
45-68°C (Annealing) 15-60 seconds
68°C (Extension) 1 minute per kb
Final Extension 68°C 5 minutes
Hold 4-10°C Can stay as long as necessary. Store in 4°C fridge or -20°C for long-term

Thermal Cycler Conditions

  1. Store samples in 4C fridge or freeze in -20C firdge for long-term
  2. Add loading dye to sample and run on .7% - 1% agarose gel to confirm amplification

Introduction:

To genetically engineer C. reinhardtii, we used the glass bead transformation method, which has been well-documented as being successful in transforming genes into the C. reinhardtii nuclear genome. Multiple variations of this protocol exist in the literature, but the protocol below is a mixture of two protocols that gave our team many transformants.

Procedure:

  1. Right before transformation, prepare linearized plasmid DNA via restriction enzyme digest using EcoRI to allow for greater chances of transformation in comparison to using circular plasmid.
  2. Grow C. reinhardtii (strain CC-400, cw-) cell cultures in a baffled Erlenmeyer flask (60 mL of TAP medium in 125 mL flasks) for 4-6 days prior to transformation or until the cell count reaches around 3 x 10^6 cells/mL.
  3. Transfer 30-50 mL of cells in TAP into 50 mL falcon tubes and centrifuge them at 3500 x g for 10 minutes. The resulting pellet should be easily visible and fully condensed to the bottom of the falcon tube.
  4. After centrifugation, discard the resulting supernatant in a waste beaker. A dark green pellet of C. reinhardtii cells should remain at the bottom of the tube.
  5. Re-suspended the pellet with 1/100 volume of TAP gently (if using 50 mL of cells, resuspend in 500 microliters TAP = 0.5 milliliters TAP). Avoid excessively agitating the cells by slowly pipetting up and down.
  6. In pre-autoclaved glass tubes with 300 mg of sterilized glass beads (425-600 micrometers in diameter), add the following in order:
    1. 2 micrograms of linearized plasmid DNA.
    2. The previously resuspended 500 microliters of cells.
    3. 5% wt/vol of 20% PEG (polyethylene glycol) (this is ~25 uL if using 50 mL of cells).
  7. Add the PEG immediately before vortexing.
  8. Vortex the tube at the highest speed for 15 seconds.
  9. Transfer the vortexed cells to a 125 mL flask containing 30 mL of TAP medium. Ensure that no glass beads are taken up alongside the transformed cells.
  10. Place the flask on a shaker at room temperature for 24 hours in the dark to allow the cells to recover post-transformation.
  11. After the 24 hour cell recovery period, centrifuge the contents of the shaking flask down into a pellet.
  12. Resuspend the pellet with 50 uL of TAP and spread onto a TAP agar plate with paromomycin (or other antibiotic selection marker).
  13. Place the plates under direct light. Colonies should appear within 5-14 days post-transformation.

Introduction:

This protocol allows for making many copies of the amplified and transformed C. Reinhardtii colonies.

Materials:

Chelex-100 Solutions Preparation

To prepare 5 mL of 5% Chelex-100 Solution:

  • Measure out 5 mL water
  • Add 0.25 g Chelex-100

Procedure:

  1. Label PCR tubes (individual, 8-strip, or 96-well plate depending on throughput) with serial numbers corresponding to the colonies being tested.
  2. Dispense 50 µL of the prepared 5% Chelex-100 solution into each of the PCR tubes.
  3. Touch each transformant colony with a pipette tip and resuspend it in its respective PCR tube.
  4. Vigorously vortex the mixture for 10s.
  5. Incubate the mixture at 95 °C (in the PCR thermocycler) for 10 min.
  6. Cool the plate on ice for 1 minute.
  7. Vigorously vortex the mixture for 10s.
  8. Centrifuge the mixture at 3500× g for 5 min at room temperature.
  9. Nanodrop supernatant.
  10. Perform a PCR reaction (using OneTaq/GoTaq) using 2 µL of the supernatant as the DNA template.

Introduction:

Part of the process of purifying recombinant proteins, Western Blot checks the tagged constructs to verify expression of our target genes.

Materials:

  • 50 mL falcon tubes
  • Wash buffer (1 -10 mL)
    • 50mM Tris-HCL pH 7.5
    • 10mM EDTA
  • 10x Protease Inhibitors (PI)
    • 100mM EDTA
    • 10mM benzzmidine- HCl
    • 500 ug/ml pepstatin A
    • 200 ug/ml leupeptin
    • 100 uM E64
    • 100 mM e-amino caproic acid, freeze
  • XCell SureLock Mini-Cell chamber
  • Pre-Cast Nupage 4-12% gel
  • 1x MES-TBST Buffer
  • CAPS Transfer buffer
    • 200 mL of ethanol
    • 4.43 g CAPS
    • 0.84 g sodium hydroxide pellets
    • Up to 2 L with DI Water
  • Owl VEP-2 Electrotransfer Chamber
    • Sandwich + Clip + Sponges
  • Immobilin-P transfer membrane (0.2um)
  • Whatman paper
  • Spatula to crack open pre-cast gel case
  • Orbital Shaker
  • 5% Milk
    • 100 mL 1XTBST
    • 5 g powdered milk
  • 5% Milk and 0.02% Sodium Azide
    • 10mL 5% milk
    • 40 uL Sodium Azide
  • Primary Antibodies
    • Dilute into sodium azide solution at appropriate conccentration
    • Can be re-used
  • Secondary Antibodies
    • Dilute in 5% milk/TBST solution to appropriate concentration
    • Cannot be re-used
  • Film Development Room:
    • Red light
    • Developer machine
  • Saran wrap
  • Film (Do NOT open box in the light)
  • Film Cassette
  • WesternBright Sirius HRP Substrate Kit
    • WesternBright SiriusTMLuminol/enhancer solution
    • WesternBright Peroxide Chemiluminescent Detection Reagent
  • Scissors

Procedure:

  1. Inocluate a Culture of C. reinhardtii
    1. Fill a beaker with 40 mL of TAP.
    2. User a sterile toothpick to smear a few colonies.
    3. Wet the side of the beaker by gently sloshing the TAP, and spin the stick against the wet side of the beaker to transfer the colonies into the media.
    4. Gently stir and allow to shake at 200 rpm with light until it reaches confluency.
  2. Total Cell Lystate
    1. Transfer 40 mL of cells into a falcon tube.
    2. Spin cells down for 5 minutes at 5000 rpm.
    3. Resuspend cells in 1 mL wash buffer.
    4. Spin the cells again for 5 minutes in a microcentrifuge tube at 4C.
    5. Aspirate the supernatant, removing as much as possible.
    6. Resuspend cells to around 10^8 cells/mL with fresh 1x PI. Keep on ice.
    7. Freeze in liquid nitrogen overnight.
    8. Thaw samples the next day on ice.
    9. BCA ASSAY
    10. Add 1 volume (=volume of cells in 1x PI) of 2x Laemmli gel sample buffer. Keep on ice until all are resuspended.
    11. Use pipet tip and then vortex. It will be viscous.
    12. Heat at 40C for 30 minutes.
    13. Vortex and spin for 5 minutes at full speed in microfuge.
    14. Transfer the supernatant to a fresh tube and discard the pellet. Keep on ice until ready to load on a gel.
  3. Western Blot - Gels
    1. Rinse XCell SureLock Mini-Cell chamber with DI water.
    2. Rinse pre-cast gel (Nupage 4-12%) with DI water prior to taking it out of the bag.
    3. Once pre-cast gel is taken out of the bag, make sure to take the tape off.
    4. Place gel into buffer core slot with the front side facing forward.
    5. Labeling should be in readable orientation.
    6. If not running two gels, place dummy gel or remove slot for second gel.
    7. Lock buffer core using gel tension wedge.
    8. Fill buffer core with 1X MES-SDS buffer.
      1. Wait for ~30 seconds to make sure that gel box is locked properly and buffer is not leaking.
      2. Continue to fill lower chamber until it is half full.
    9. Take out comb from pre-cast gel.
    10. Wash out wells using p200 (~100 ul)
    11. Load ladder using gel-loading 200 uL pipette tips
      1. Load 7-15 ul of sample using gel-loading 200 uL pipette tips
      2. Do not leave empy wells next to samples: Fill with Laemmli
      3. Only load 7ul of ladder and Laemmli, can load more of sample based on concentration of protein (~7 ug)
      4. Be careful with the final dispensing bubble
    12. Run gel at 120 V *buffer should fill up the core and cover the electrodes outside of the core
      1. When starting the electrophoresis, ensure that there are no bubbles
      2. Make sure the gel actually starts running
    13. Stop the gel once the samples have moved sufficiently down gel (~45 mins. Less if smaller proteins)
    14. While Gel is Running:
    15. Make 2 L of transfer buffer
      1. 200 mL ethanol
      2. 4.43 g CAPS
      3. 0.84 g sodium hydroxide pellets
      4. DI water up to 2L
      5. Mix using a stir bar
    16. After Gel is done running: Rinse Owl VEP-2 and sandwich with DI water
    17. Fill Owl VEP-2 with transfer buffer (CAPS)
    18. Cut Immobilin-P transfer membrane to appropriate size
    19. Cut Whatman paper to appropriate size
    20. Fill a bin with transfer buffer to soak both sides of the sandwich
    21. Fill a bin with ethanol (large petri dish)
    22. When electrophoresis is done, unlock buffer core
    23. Remove pre-cast gel from buffer core
    24. Rinse pre-cast gel with DI water
    25. Use spatula to crack open pre-cast gel case
    26. Cut off excess gel (bottom of gel, sides, and top)
    27. Set Up Sandwich as follows:
      1. 1. Lay down Sandwich with BLACK on bottom
      2. 2. Sponge
      3. 3. whatman paper
        • (1) Roll out whatman paper with serological pipet so that there are no bubbles
      4. Place gel (already cut) so that PROTEINS FACE RED.
      5. Place transfer membrane on gel, facing proteins.
        • (1) Activate transfer membrane prior to using by soaking it in ethanol for ~10 seconds.
        • (2) Use non-serrated forceps to handle transfer membrane
        • (3) Make sure it is evenly coated and there are no bubbles (roll out0.
      6. Equilibrate transfer membrane with the transfer buffer in the Owl VEP-2
      7. Whatman paper
        • (1) Roll out whatman paper with serological pipet so that there are no bubbles
        • (2) Add some buffer to wet paper
      8. Sponge
      9. RED on top
    28. Lock Sandwich with plastic clip
    29. Place sandwich in Owl VEP-2
      1. Rubber clip should be facing the top
      2. Black side of the sandwich should be facing forward when the electrodes are on the right (black side of the sandwich should align with the black elecrtodue, red should face red electrode).
      3. If possible, place sandwich as close to positive electrode.
    30. Top off Owl VEP-2 with transfer buffer
    31. Run the electrotransfer at 45 V for 3 hours at 4 C.
    32. Remove the sandwich and membrane from sandwich
    33. Equilibrate the membrane by placing it in dish with 1X TBST on orbital shaker at 40 rpm for ~5 min.
    34. Make 5% milk.
      1. 100 mL TBST
      2. 5 g milk
    35. Western Blot - Blocking and Tagging
      1. Transport samples to film developer room, and bring the following:
        1. waste container
        2. saran wrap
        3. film
        4. cassette
        5. 1.5 mL or 5 mL microcentrifuge tubes
        6. WesternBright Sirius HRP Substrate Kit
          • (1) WesternBright SiriusTMLuminol/enhancer solution
          • (2) WesternBright Peroxide Chemiluminescent Detection Reagent
        7. p1000
        8. scissors
        9. timer
        10. 1X TBST
        11. Marker
      2. Turn on developer (should take ~15 minutes to be ready to use)
        1. Wait until red "ready" light illuminates
        2. When ready to use, place used film in developer to wet rollers
      3. Change out 1X TBST and let sit for 5 minutes (second wash)
      4. Change out 1x TBST and let sit for 5 minutes (third wash)
      5. Prepare enough HRP substrate (1:1) to cover samples.
      6. Place membrane on saran wrap and cover samples with HRP substrate for 1 minute.
      7. Place membrane on cassette (protein side up).
      8. IN THE DARK:
        1. cut film in half
        2. Cut one corner off of film piece: place this corner in the corner of the cassette
        3. Place film in cassette
        4. do not shift film once placed, place directlyh on top
        5. close cassette for appropriate time according to antibody used
        6. place in developer
        7. Wait until film is entirely done going through developer
        8. TURN LIGHTS BACK ON
      9. Mark film
        1. Membrane edges
        2. Relevant ladder bands
        3. Date
    36. Stripping/Re-Blocking/Re-Tagging
      1. Rinse Membrane 3x in TBST for approx 5 mins each rinse
      2. Cover membrane in small amount of antibody stripping solution
      3. Place on shaker @ 40 rpm for 5 mins
      4. Pour off stripping solution
      5. Re-rinse membrane 3 times in TBST for approx 5 mins each rinse
      6. Re-block exposing to 5% milk in TBST solution for 1hr or overnight
      7. Repeat blocking/tagging section of protocol

Introduction:

Inductively coupled plasma mass spectrometry allows for the analysis and quantification of elements, and in this case arsenic, at trace levels in biological samples.

Materials:

  • 7.5% sodium bicarbonate solution
    • 3 g sodium bicardonate
    • 37 mL DI water
    • Filtered to sterlize resulting solution
  • 1000x Arsenic stock solution
    • 4.16 g sodium arsenate dibasic heptahydrate
    • 900 mL DI water
    • Mixed to produce resulting solution
    • To be made in chemical fume hood
  • Cultures of (3) mutant strains (8622.1.1, 8622.1.2, 8622.3.10) and control CC-400 control strain
  • Orbital shaker
  • Hemocytometer
  • 50 mL falcon tubes
  • 15 mL falcon tubes
  • TP media
  • 125 mL baffled flasks

    • Procedure:

    Sample Preparation and Day 0 collection

    1. Gather cultures of three mutant strains (8622.1.1, 8622.1.2, 8622.3.10) and CC-400 control strain.
    2. Count cells in each culture using a hemocytometer.
    3. Normalize each culture to have a standard cell density of 106 cells/mL.
    4. Add 50 mL of culture to 4 50-mL falcon tubes each (label the corresponding strain on each tube).
    5. Spin down cultures at 7100 rpm for 5 minutes at room temperature (21˚C).
    6. Discard the supernatant and resuspend the pelleted cells in 1 mL of TP media.
    7. Assemble the 12 cultures in 125 mL baffled flasks as follows (there should be 4 samples at each arsenic concentration, one for each strain tested):
    Sample concentration (ppb Arsenic) Culture added (mL) TP added (mL) Bicarbonate added (mL) 1000 x Arsenic stock solution added (uL) Total volume (mL)
    50 ppb 1 mL 43.2 mL 0.8 mL 2 uL 45 mL
    250 ppb 1 mL 43.2 mL 0.8 mL 10 uL 45 mL
    500 ppb 1 mL 43.2 mL 0.8 mL 20 uL 45 mL
    1. The cultures should be labeled as follows:
    CC-400 (50 ppb) CC-400 (250 ppb) CC-400 (500 ppb)
    8622.1.1 (50 ppb) 8622.1.1 (250 ppb) 8622.1.1 (500 ppb)
    8622.1.2 (50 ppb) 8622.1.2 (250 ppb) 8622.1.2 (500 ppb)
    8622.3.10 (50 ppb) 8622.3.10 (250 ppb) 8622.3.10 (500 ppb)
    1. Immediately aliquot 10 mL of solution from each of the samples into 15 mL falcon tubes to obtain arsenic uptake levels at t=0.
    2. Record the time at which these samples were removed from culture.
    3. Count cells in each of these samples using a hemocytometer and record cell densities.
    4. Spin down the 10 mL samples in a centrifuge at 7100 rpm for 5 minutes at room temperature.
    5. Decant the supernatant into a separate, clearly labeled falcon tube and store safely at room temperature.

    Day 1 Collection

    1. 24 hours after collecting t=0 samples, aliquot 10 mL of solution from each of the samples into 15 mL falcon tubes to obtain arsenic uptake levels at t=24 hr.
    2. Record the time at which these samples were removed from culture.
    3. Count cells in each of these samples using a hemocytometer and record cell densities.
    4. Spin down the 10 mL samples in a centrifuge at 7100 rpm for 5 minutes at room temperature.
    5. Decant the supernatant into a separate, clearly labeled falcon tube and store safely at room temperature.
    6. Repeat these steps for day 2 and 3 collection.

    Safety and Decontamination

    1. Arsenic stock solution creation will be done in the Bartelle lab chemical fume hood.
    2. All flasks will be thoroughly cleaned and sterilized to remove any residual arsenic contamination.
    3. All falcon tubes and other disposables that came in contact with arsenic will be disposed of in a clearly labeled bin for arsenic waste.