Team:Washington/Protocols

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Protocols

Aptamer Protocols

• Auxin Protocols
• Aptamer Protocols
• Paper Protocols

Yeast Chromosome Integration

Digest E. Coli plasmid using PmeI restriction enzyme

• 1 ug of DNA
• 5 uL of 10x NEB CutSmart buffer
• 1 uL of restriction enzyme
• Fill to 50 uL with water
• Incubate at 37 C for 15 minutes (1 hour if not using TimeSaver buffer)
• Heat inactivate at 65 C for 20 minutes
• Agarose gel purify (optional)

Salmon Sperm Transformation

• Grow a yeast overnight
• Check OD of culture. 0.5-0.6 are the preferred readings, if the reading is lower, wait for longer growth, if the reading is higher, dilute the sample.
• Spin down 10 ml of cells per transformation.
• Decant supernatant and wash with 10 ml ddH2O. Vortex to resuspend and spin down.
• Remove the supernatant.
• Resuspend cells in 300 uL .1 M LiOAc. Transfer to a 1.5 mL tube.
• Incubate at 30 C for 15 min
• Put salmon sperm DNA in boiling water for 5 minutes. Cool immediately on ice.
• Spin down cells and remove supernatant.
• Add the following in order:
1. 240 uL 50% PEG
2. 36 uL 1.0 M LioAc
3. 10 uL salmon sperm DNA
4. 34 uL DNA
5. 40 uL ddH2O
6. Final volume: 360 uL

Theophylline Stock

50 mM Theophylline dissolved by DMSO

Replace occasionally due to possible interactions between theophylline & DMSO

Fluorescence Reading

• Put yeast plate to a blue light imager.
• Note differences in brightness between yeast colonies

Assay:

• Fill 96 well plate with 250 uL of cell cultures

Flow Cytometer:

• Set a bottom cutoff of 10,000 units
• Excitation: 515 nm
• Emission: 530 nm

X-GAL ASSAY

1. Prepare a solution of 50 mg/mL X-Gal DMSO.
2. Take one ml of cells (OD 500+) and spin down.
3. Remove supernatant as you please and add 1ml of water.
4. Resuspend cells, spin down, and remove supernatant again.
5. Add 5ul X-Gal solution, 25 ml 2% SDS and 70ml water. Mix well.
6. Incubate @ 37C someplace dark for 30+ min.

Auxin Assay.

1. Prepare a solution of 100 mMol auxin in ethanol.
2. Add 3ul auxin per ml of cell culture you are using.
3. Wrap your culture container in foil and put in the shake incubator @ 30C for 4+ hours.
4. Finish by doing the X-Gal assay described above. Alternatively, just add the SDS and X-Gal directly.

Transformation of E.Coli Competent Cell :

1. Thaw competent E.coli cells on ice (XL1-Blue for cloning)
2. Add 50 uL of competent cells to sterile 14 mL Falcon culture tube.
3. Add 1 uL of the miniprep to each culture tube
4. Equilibrate the cells on ice for 10 min
5. Heat shock the cells at 42C for 30-45 second.
6. Immediately place the cells back on ice for 3 min
7. Add 250 uL LB media and shake at 250 rpm and 37C for 30 min
8. Plate 10 ul and 290 ul of the recovered cells onto LB-agar plates supplemented with appropriate antibiotics (spread with ~150uL DiH2O)
9. Invert and incubate at 37C overnight

DIGESTION:

• Buffers are in 10X. Upper limit 120ng/uL for Plasmids. 50uL Reaction Volume
• DNA/Plasmid: minimum 1ug - 5ug max (more or less depending on the amount from the miniprep)
• DO NOT EXCEED 120ng/uL
• 10x Buffer: 5uL of NEBuffer 2.1
• Enzyme: 1uL of EcoRI and 1uL of Nhel (or other restriction enzyme)
• *Add enzyme last
• ADD water to 50uL total volume

Incubate at 37 C for 1 Hour

High Efficiency Yeast Transformation Protocol

(Modified from Agatep, R. et al. (1998) Technical Tips Online)

1. Inoculate a 50 ml ypd liquid culture (with a single yeast colony). This volume depends on how many transformations are needed. Allow about 5-10ml of culture per sample. However, never start a culture less than 50ml.
2. Grow overnight with shaking at 30° C.
3. In the morning, read the OD of the culture. A culture in mid-log with an OD between 0.5 and 0.6 is ideal (which represents approximately 2.0 x 107total yeast cells per ml), but cultures ranging from 0.4 to 0.8 are also acceptable. If the culture is overgrown, dilute to approximately 0.2 and grow for 2 generations (2-3 hours). If efficiency is not essential, it is acceptable to use 10 ml of a culture at 0.3 OD.
   1. How to use the spectrometer. Cuvettes are located in a Styrofoam box on the right of the computer.
   2. Make sure the reading is at 600 nm
   3. Blank with 900 uL of ypd
   4. Add 100 uL of culture to make a 1:10 dilution
   5. Measure the OD. Keep in mind that the actual OD is 10x larger
   6. Use vacuum line to empty the cuvette, cuvettes can be reused
   7. Dilute or concentrate and repeat measurements until OD is in the range
4. Spin down 10 ml of cells per transformation (2000 rpm / 2 min) in a 15 ml Falcon tube (which equals 2 x 108 total cells). Multiple transformations with the same strain can be combined into larger volumes up until step 7. Please note that these volumes are for haploid strains. A transformation with a diploid strain would require 5 ml of culture per transformation (equaling 1 x 108 total cells).
5. Decant supernatant and wash with 10 ml ddH2O. Vortex briefly to resuspend cells. Spin down cells (2000 rpm / 2 min). Completely remove supernatant.
6. Resuspend cells (by vortexing) in 300 µl 0.1 M LiOAc. Transfer to a sterile 1.5 ml eppendorf tube. (This step and those that follow list volume amounts that are PER TRANSFORMATION).
7. Incubate at 30° C for 15 min.
8. Begin boiling water to denature Salmon Sperm DNA. Boil 5 min. Cool immediately on ice to prevent re-annealing of single stranded DNA.
9. Spin down cells (2500 rpm/ 1 min). Remove supernatant with a pipetman.
10. Add the following components of the transformation mix in order:
1. 240 µl 50% PEG
2. 36 µl 1.0 M LiOAc
3. 10 µl Salmon Sperm DNA (10 mg/ml)
4. 34 µl DNA (plasmid, PCR or digest)
5. 40 µl ddH2O

FINAL VOLUME: 360 µl

Please note: a mastermix at this point is acceptable if transforming multiple samples into the same yeast strain. Add 326 µl of mastermix to each pellet, then the DNA sample.

11. Vortex thoroughly to resuspend cells (approximately 30 sec). Be sure all cells are in suspension.
12. Incubate at 30° C for 2 hours. Mix cells gently every hour. Incubation time can range anywhere from 1.5 hours to 3 hours, resulting in the same transformation efficiency.
13. Add 36 µl of DMSO and mix by inverting tubes. (DO NOT VORTEX)
14. Heat shock for 5 min at 42° C.
15. Spin down (2500 rpm / 1 min). Remove supernatant with a pipetman.
16. Wash cells by gently pipeting cells in 1 ml ypd liquid (or selective media). Spin down (2500 rpm / 1 min). Remove supernatant.
17. Resuspend cells by gently pipeting in 1 ml liquid media. Dilute if necessary.
18. Incubate culture with shaking at 30° C overnight if expression is necessary, otherwise skip this step. Often cells expressing certain selectable markers require an additional incubation at this step to allow phenotypic expression of the antibiotic resistance marker.
19. Titer on selective plates (10-1,10-2,10-3 are the standard dilutions).
1. If you re suspend the cells in 1 ml of -ura then plate 100ul on a -Ura plate, that is a 10-1 dilution.
2. If you plate 100 or 200ul on a plate, you'll get colonies for sure
20. Plate on selective media.

Making 1mm-Thick Sheets of PDMS (Polydimethyl Siloxane)

• Obtain Sylgard 184 Silicone Elastomer Kit from Dow Corning
• Obtain cell culture plates to use as templates
• Calculate surface area of plate in order to determine the mass needed of PDMS
• Make a 1:10 mixture of activator to PDMS
• Stir well using a wood or plastic stick
• Pour polymer onto the top of the lid or bottom of the culture plate. Pour into the middle and work out towards the edges. Tilt plate to let PDMS
• flow to the edges of the plate.
• Remove bubbles, either by putting the sample in a vacuum or by popping the bubbles with the stirring stick
• Let polymer cure for 2 days in a flat place to ensure even distribution.

Making the Paper Device

• Cut strips of Whatman 114 filter paper to dimensions 3cm x 4cm
• Cut 2cm x 2cm pieces of PDMS sheet
• Use heat-resistant tape (such as electrical tape or 3M heat-resistant packing tape) to tape polymer squares to either side of the paper strip. On one side, tape around all four sides of the PDMS. On the other, leave the top open in order to insert media and cells.
• Autoclave devices in a sterile container on a dry cycle

Inserting cells and media

• Make either liquid or gel media with conditions specific for the intended yeast strain (note that liquid media may dry out fast)
• For liquid media, pipette no more than 100uL through the open side of the device, between the paper and the PDMS window
• For gel media, pipette no more than 300uL between the paper and PDMS
• Using a pipette tip or toothpick, insert a clump of cells and deposit them in the middle of the PDMS window. For gel media, it may be necessary to use a toothpick to poke a pathway through the gel in order to ensure cells can be deposited easily.
• Tape over the remaining side of the device to seal the cells and media inside

Running detection assays

• The molecule of interest can either be pipetted directly between the paper and PDMS along with the yeast media, or the device can be stood up in a solution of the molecule, which can travel to the yeast via wicking.

Yeast lysis on paper to view beta-Gal production

• Place yeast on liquid media inside a device.
• Pipette 30uL of 0.2% SDS and 10uL of 50mM X-gal directly to the cells, or place end of device in solution of 300uL of 0.2% SDS and 100uL X-gal
• When SDS is in contact with cells, massage PDMS window briefly to mix
• Allow devices to sit at 37C for 30 minutes to 1 hour

Observe blue color

• Theophylline detection on paper (assuming inducible Gal promoter)
• Load device with theophylline-detecting strain and c-ura media, with galactose as the sugar*
• Seal device
• Allow device to sit in a beaker of 3mL of 50mM theophylline solution for 6-10 hours.
• Observe fluorescence

*Alternatively, use media with no sugar and add galactose the same way as theophylline

Possible controls

• Strain that doesn’t produce theo aptamer with theophylline (negative control)
• Theo detection strain without theophylline (negative control)
• Strain that constitutively produces YFP with theophylline (positive control)
• Theo detection strain on arafinose or glucose c-ura media
• Theo detection strain on galactose c-ura with caffeine (no theophylline)

Auxin detection on paper

Possible controls

• Strain without auxin detection pathway with auxin (negative control)
• Strain with auxin detection pathway without auxin (negative control)
• Strain without gRNA, which constitutively produces beta-Gal/blue chromoprotein with auxin (positive control)

Insert text

For commercial shellfish farmers and recreational hunters alike, marine biotoxins pose a significant threat to health and welfare. With this project, we aim to create an inexpensive and easy-to-use test kit for the detection of the shellfish toxin okadaic acid using engineered yeast strains and DNA aptamers on a paper device. We also hope that this project paves the way for a new class of biosensors capable of detecting a wide range of small molecules.