Experiments

Lac Z Buffer

For the beta-galactosidase assay utilized in the in-cell assay [8].

Reagents

• 2.13 g Na2HPO4 (dibasic)
• 1.20 g NaH2PO4 or 1.38 g NaH2PO4•H2O (monobasic)
• 23.8 mg MgCl2 (corresponds to 9.99 mL of a 25 mM MgCl2 solution)
• 186.4 mg KCl
• 0.5 g N-lauroylsarcosine sodium salt
• 100 mg ONPG
• Deionized water to a final volume of 250 mL

Consumables

• Weighing boats

Equipment

• 1 L flask
• Decimal scale

Procedure

1. Mix everything to a final volume of 250 mL.
2. Store the buffer in 20 mL aliquots at -20°C (sufficient for one 96 well plate in the in-cell assay assay).

Glucose/Galactose Media

Minimal defined yeast media without tryptophan. Allows for selection of successful transformants using the TRP1 marker.

Reagents

• 1.675 g Yeast Nitrogen Base without amino acids (YNB) (Should be 0.67%)
• 25 ml 20% glucose & galactose
• 0.48 g Yeast Synthetic Drop-out Medium Supplements without Tryptophan (Should be 0.192%)
• 225 ml distilled water
• 5 g Bacto agar

Consumables

• Weighing boats

Equipment

• Autoclave
• 500 ml bluecap bottle
• Scale

Procedure

1. Add YNB, Drop-out Medium Supplements to a 500 ml bluecap bottle.
2. Add 450 ml water and mix with a stirring bar until dissolved.
3. Add 5 g Bacto agar.
4. Autoclave.
5. Add 25 ml 20% glucose/galactose to each bottle (25 ml).

LB amp agar

Ensures a 100ug/mL ampicillin conc. applicable with E. coli with amp resistance gene

Reagents

• 10 g LB broth with agar (miller)
• 250 ml ddH2O
• 100 mg/ml ampicillin solution

Consumables

• Weighing boats

Equipment

• 500 ml bluecap bottles

Procedure

1. Add 10 g LB broth with agar (miller) to a 500 mL bluecap bottle.
2. Add 250 ml water and mix with a stirring bar while heating until dissolved.
3. Autoclave.
4. Add 250 µL 100 mg/mL amp stock solution.
5. NOTE: Ensure that the media is cooled to around 50°C before adding ampicillin. Higher temperatures degrade it.

LB Amp Broth Preparation

This protocol ensures a 100 µg/mL ampicillin concentration, applicable for use with E. coli strains carrying the ampicillin resistance gene.

Materials:

• Reagents:
• 5 g LB broth
• 250 mL ddH₂O
• 100 mg/mL ampicillin solution
• Consumables:
• Weighing boats
• Equipment:
• 500 mL bluecap bottles

Procedure:

1. Add 5 g of LB broth to a 500 mL bluecap bottle.
2. Add 250 mL ddH₂O and mix with a stirring bar while heating until completely dissolved.
3. Autoclave the solution to sterilize.
4. Once cooled to approximately 50°C, add 250 µL of 100 mg/mL ampicillin stock solution.
5. Note: Ensure the media is cooled before adding ampicillin, as higher temperatures degrade the antibiotic.

Hormone Dilution Standards

Protocol for diluting the desired hormone standards for both the assays, In-cell and Cell-free. CAUTION: These hormones are harmful if inhaled, swallowed, or absorbed through the skin. Use appropriate personal protection (gloves, fume hood, dust mask) and avoid exposure during pregnancy and lactation. Dispose as hazardous waste and do not release into the environment.

Materials

• Reagents:
• Hormone powders
• Ethanol
• 50% ethanol
• Consumables:
• Eppendorf tubes
• Parafilm
• Equipment:
• Scale
• 1000 μl pipette
• 100 μl pipette
• 10 μl pipette

Procedure

Dilution of Estradiol to Working Conditions

1. Mix 1 mg estradiol powder with 1 ml ethanol and let it dissolve. This yields a 3.67 mM temporary stock #1.
2. Dilute 10 μl of stock #1 in 990 μl ethanol. This yields a 36.71 μM temporary stock #2.
3. Dilute 100 μl of stock #2 in 900 μl 50% ethanol. This yields a 3.6713 μM temporary stock #3.
4. Dilute 619.7 μl of stock #3 in 380.3 μl 50% ethanol. This yields the 2.275 μM working stock for the testrun (E1).
5. Dilute 100 μl of stock #3 in 900 μl 50% ethanol. This yields a 367.13 nM temporary stock #4.
6. Dilute 619.7 μl of stock #4 in 380.3 μl 50% ethanol. This yields the 227.5 nM working stock for the ERα receptor (E2).
7. Dilute 26.56 μl of stock #4 in 973.44 μl 50% ethanol. This yields the 9.75 nM working stock for the ERβ receptor (E3).
8. Seal tubes with parafilm and store at -20°C or -70°C.

Dilution of Testosterone to Working Conditions

1. Mix 22.6 mg testosterone powder with 1 ml ethanol and let it dissolve. This yields a 78.4 mM temporary stock #1.
2. Dilute 10 μl of stock #1 in 990 μl 50% ethanol. This yields a 784 μM temporary stock #2.
3. Dilute 128 μl of stock #2 in 872 μl 50% ethanol. This yields the 100 μM working stock for the AR.
4. Seal tubes with parafilm and store at -20°C or -70°C.

Dilution of Aldosterone to Working Conditions

1. Mix 5 mg aldosterone powder with 1 ml ethanol and let it dissolve. This yields a 13.9 mM temporary stock #1.
2. Dilute 72.3 μl of stock #1 in 927.7 μl 50% ethanol. This yields the 1 mM working stock for the MR.
3. Seal tubes with parafilm and store at -20°C or -70°C.

Dilution of Dexamethasone to Working Conditions

1. Mix 18.5 mg dexamethasone powder with 1 ml ethanol and let it dissolve. This yields a 47.1 mM temporary stock #1.
2. Dilute 21.3 μl of stock #1 in 978.7 μl 50% ethanol. This yields the 1 mM working stock for the GR.
3. Seal tubes with parafilm and store at -20°C or -70°C.

Plasmid Transformation of E. coli

Introduction: Transformation protocol for DH5alpha or BL21 E. coli cells that were prepared using the Mix&Go E. coli Transformation Kit developed by [2]

Materials

Reagents:
• 100 μL x (N+2) LB (use SOC for better results)
• MQ water
• N+2 competent E. coli tubes (N+1 if no positive control)
• N amount of cloning products
• Negative control (NC) from cloning (can use water instead)
• Positive control (PC) (optional but suggested)
Consumables:
• N+2 LB antibiotic plates (N+1 if no positive control). Can multiply this amount if plating different amounts (step 10).
• Pipettes
• 200 μL tips
• 10 μL tips

Procedure

Prepare

1. Take competent cell tubes and thaw them on ice.
2. If competent cells are in larger volumes, split them into tubes of 50 μL each.
3. Label each tube, remembering NC (Negative Control) and PC (Positive Control).
4. Take plates and prewarm them in the incubator.

Transform

1. While cells are still on ice, add 2 μL of cloning product or controls. Mix gently with a pipette. Note: Quantify the DNA! This protocol is optimal with ~100 ng/μL of DNA, so a total of 200~250 ng of DNA.
2. The optimal protocol states that no more than 5% of volume (of DNA) should be added to the competent cells. So, 5% of 50 μL = 2.5 μL.
3. Incubate on ice for 5 minutes.
4. Add 100 μL of SOC medium to each tube.
5. Incubate at 37°C for 1 hour. This can be done in an incubator or shaker (preferably sideways for better mixing; tape the tubes so they don't fall out).
6. Label the plates while waiting.

Plate

1. Plate the appropriate amount of media. It could be 50 μL, 100 μL, or 200 μL. You can plate different volumes on different plates for each tube.
2. Incubate plates at 37°C overnight. If no results appear, incubate longer. Note: Beware of satellite colonies if using ampicillin and incubating for a long period.

Yeast Transformation

This protocol is used to make competent yeast cells and then transform them with the desired plasmid

Reagents

• 1 ml Frozen yeast glycerol stock
• 150 ml YPD media
• ca. 30 ml Sterile H2O
• 1.5 ml 0.1M lithium acetate (LiAc)
• 0.5 ml 1M lithium acetate (LiAc)
• 2.4 ml PEG 3350 (50% w/v)
• 0.5 ml Pre-denatured 2.0mg/ml single-stranded carrier DNA (salmon sperm derived). To prepare, boil 1 ml of the ss DNA for 5 min and quickly chill it on ice. Aliquote it and store at -20°C.

Consumables

• Spectrophotometer cuvettes
• Sterile 50 ml falcon tube
• Sterile 1.5 ul eppendorf tubes

Equipment

• Incubator set at 30°C and 200 rpm
• Spectrophotometer set at 660 nm
• Centrifuge for 50 ml falcon tubes set at 3000g
• Centrifuge for 1.5 ml eppendorf tubes set at 6000-8000rpm
• Water bath / heating block set at 42°C
• Pipettes
• Vortex
• Culture flask

Procedure

Day 1

Inoculate 100 ml YPD with 1 ml of a glycerol yeast stock. Grow O/N at 30°C with 200 rpm.

Day 2

After 12-16h determine the titer of the yeast culture using a spectrophotometer: Pipette 10 μl of cells into 1.0 ml of water in a spectrophotometer cuvette, mix thoroughly by pipetting and measure the OD at 660 nm.

1. Determine the titer in the cell culture (remember the dilution factor). A suspension containing 1*10⁶ cells ml^-1 will give an OD₆₆₀ of 0.1.
2. Add 2.5*10⁸ cells to 50 ml of pre-warmed YPD in a pre-warmed culture flask: this will give a final solution of 5*10⁶ cells ml^-1.
3. Incubate the flask in the shaking incubator at 30°C with 200 rpm until the cell titer is at least 2*10⁷ cells ml^-1. This should take about 4 hours. Follow the OD closely toward the end.
4. Transfer the culture to a sterile 50 ml falcon tube and centrifuge at 3000g for 5 minutes.
5. Pour off the medium, resuspend the cells in 25 ml of sterile H₂O, and centrifuge again.
6. Pour off the H₂O, resuspend the cells in 1.0 ml of 0.1M lithium acetate (LiAc), and transfer the suspension to a sterile 1.5 ml microfuge tube.
7. Centrifuge the cells at top speed for 5 seconds and remove the supernatant with a pipette.
8. Resuspend the cells in 0.1M LiAc to a final volume of 500 μl (by adding around 400 μl of 0.1M LiAc). Aliquot 50 μl per transformation.
9. Spin down the cell suspension and remove the supernatant.
10. Add the following transformation mix to each of the eppendorf tubes in the specified order:
    • 240 μl PEG 3350 (50% w/v)
    • 36 μl 1.0M LiAc
    • 10 μl pre-denatured single-stranded carrier DNA (5.0 mg/ml) (thaw on ice)
    • 65 μl total: H₂O + plasmid DNA (0.1-10 μg)
    Note: The order is important! The PEG, which shields the cells from the detrimental effects of the high concentration of LiAc, should go in first.
    Note 2: You can also make a premix of all the ingredients without the plasmid DNA. Add this to the cells first, followed by the DNA. This mix can be stored at -80°C before adding the DNA for future use.
11. Mix by pipetting up and down slowly until the cell pellet is completely mixed (about 1 minute).
12. Incubate for 30 minutes at 30°C.
13. Heat shock for 30 minutes in a water bath at 42°C.
14. Centrifuge the tubes at 6000-8000 rpm for 15 seconds and remove the transformation mix with a pipette.
15. Add 0.2-1.0 ml of sterile H₂O to each tube and resuspend the pellet by pipetting up and down gently.

Plating Procedure:

1. If using a dominant marker for transformation, add 1 ml YPD to each tube, resuspend the pellet by pipetting up and down gently, and incubate for 1-2 hours at 30°C.
2. If performing a plasmid transformation, plate both a small amount (10 μl + 100 μl H₂O) and a larger amount (200 μl) of the transformation mix onto selective plates.
3. For integration transformations, plate the entire amount of the transformation mix onto selective plates.
4. Incubate the plates at 30°C for 3 days.

Preparation of Glycerol Stocks

Preparation of glycerol stocks for long-term storage at -80°C, suitable for both E. coli and yeast strains. The primary difference is the choice of media and incubation temperature.

Materials

Reagents

• 50% glycerol (Add 500 mL glycerol to a 1L glass bottle. Fill up to 1L with distilled water. Autoclave)
• Broth for the O/N culture (e.g., YPD)

Consumables

• Inoculation needles
• 1000 μL pipette tips
• Freezing vials

Equipment

• Incubator (set to 30°C and 200 rpm for yeast, or 37°C and 200 rpm for E. coli)
• 1000 μL pipette
• Glass test tube

Procedure

Day 1

1. Inoculate 5 mL of broth (e.g., YPD for yeast, or LB for E. coli) with the strain in a glass test tube.
2. Incubate overnight in an incubator (set to 30°C and 200 rpm for yeast, or 37°C and 200 rpm for E. coli).

Day 2

1. Label cryotubes and record corresponding information in Benchling (name of culture, name, and date).
2. Pre-fill the cryotube with 750 µL of 50% glycerol solution.
3. Add 750 µL of the overnight culture to the cryotubes.
4. Invert the tubes several times to ensure proper mixing.
5. Store the tubes in the -80°C freezer.

PCR with Phusion U Hot Start Polymerase

Protocol adapted from [3]. We use it to amplify the G-blocks attached with primers compatible for the USER reaction in the following steps.

Reagents

• 10 mM Primers
• 10 mM dNTPs mix
• PCR grade water
• Template DNA
• Phusion U DNA Polymerase
• 5X Phusion HF Buffer

Consumables

• desired number of PCR tubes
• Pipettes
• 20 µL or 10 µL tips
• 200 µL tips

Procedure

Preparation

1. Label PCR tubes, including negative and positive controls. Thaw the reagents. Only take the Phusion U Polymerase out right before use.
2. Calculate the template concentration (using gel or Nanodrop) to determine the amount of template to add.
3. Prepare the Master Mix without the template/primers if these differ between reactions.
4. Remember to dilute primers to 10 µM - they are 100 µM when resuspended from the supplier.

Master Mix Composition

Component	50 µL Reaction	Final Concentration in Reaction
H2O	32.5 µL
5X HF Buffer	10 µL	1x
10 mM dNTPs	1 µL	200 µM (each)
Primer FWD (10 µM)	2.5 µL	0.5 µM
Primer RV (10 µM)	2.5 µL	0.5 µM
Phusion U Polymerase	0.5 µL	0.02 U/µL
Template	1 µL	1 pg - 10 ng DNA per 50 µL

Reaction Setup

Add the DNA template/primers to each tube. For the negative control tube, add water (or nothing) instead of the DNA template.

Thermocycler Program

Step	Temperature (°C)	Time	Repeats
Initial denaturation	98	2 min
Denaturation	98	15 sec	25-30x
Annealing	Tm - 5	20 sec
Elongation	72	kb x 15-30 sec
Final elongation	72	5 min
Hold	4 or 12	infinite

Massive PCR for Cell-Free Assay Template

We use the AQ97 High Fidelity DNA Polymerase 2x Master Mix to amplify massive amounts of the templates for our cell-free biosensor [4].

Process

Split the 1000 µL into 2 PCR strips of 8 wells each, resulting in 16 wells, with 62.5 µL in each well.

Reaction Preparation

Component	Volume (µL)	Final Conc.	Initial Conc.
H2O	up to 1000
2X Master Mix Buffer	500	1x
Primer FWD (10 µM)	30	0.3 µM	10 µM
Primer RV (10 µM)	30	0.3 µM	10 µM
Template	Vary		20 ng DNA
Total	1000

Thermocycler Program

Step	Temp (°C)	Time	Cycles
Initial Denaturation	98	2 min
Denaturation	98	15 sec
Annealing	60	20 sec	37 cycles
Elongation	72	20 sec
Final Elongation	72	5 min
Hold	4 or 12	Infinite

PCR Clean-up (NucleoSpin®)

The following protocol (NucleoSpin®) is suitable for PCR clean-up, DNA concentration, and removal of salts, enzymes, etc. from enzymatic reactions (SDS < 0.1%)NucleoSpin Gel and PCR Clean-up [5]

Materials

• DNA binding buffer
• Elution buffer
• Column

Procedure

Adjust DNA binding condition

For very small sample volumes < 30 μL, adjust the volume of the reaction mixture to 50–100 μL with water. It is not necessary to remove mineral oil.

Mix 1 volume of sample with 2 volumes of Buffer NTI (e.g., mix 100 μL PCR reaction and 200 μL Buffer NTI).

Note: For removal of small fragments like primer dimers, dilutions of Buffer NTI can be used instead of 100% Buffer NTI.

Bind DNA

Place a NucleoSpin® Gel and PCR Clean-up Column into a Collection Tube (2 mL) and load up to 700 μL sample. Centrifuge for 30 s at 11,000 x g. Discard flow-through and place the column back into the collection tube.

Note: Load remaining sample if necessary and repeat the centrifugation step.

Wash silica membrane

Add 700 μL Buffer NT3 to the NucleoSpin® Gel and PCR Clean-up Column. Centrifuge for 30 s at 11,000 x g. Discard flow-through and place the column back into the collection tube.

Recommended: Repeat the previous washing step to minimize chaotropic salt carry-over and improve A260/A230 values.

Dry silica membrane

Centrifuge for 1 min at 11,000 x g to remove Buffer NT3 completely. Make sure the spin column does not come in contact with the flow-through while removing it from the centrifuge and the collection tube.

Note: Residual ethanol from Buffer NT3 might inhibit enzymatic reactions. Total removal of ethanol can be achieved by incubating the columns for 2–5 min at 70°C prior to elution.

Elute DNA

Place the NucleoSpin® Gel and PCR Clean-up Column into a new 1.5 mL microcentrifuge tube (not provided). Add 15–30 μL Buffer NE and incubate at room temperature (15–25°C) for 1 min. Centrifuge for 1 min at 11,000 x g.

Note: DNA recovery of larger fragments (> 1000 bp) can be increased by multiple elution steps with fresh buffer, heating to 70°C, and incubation for 5 min.

USER Cloning

Using the specific design primers, we can perform User cloning attached parts in a customize manner [6].

Materials

Reagents:

• USER reaction targets (PCR products)
• CutSmart buffer
• USER enzyme (keep on ice)
• DpnI enzyme (keep on ice)
• MQ water

Procedure

Prepare the mix:

Reagent	Volume (μL)
MQ water	11 μL
PCR product	2 μL each
CutSmart buffer	1 μL
USER enzyme	1 μL
DpnI enzyme	1 μL
Total	20 μL

Thermocycler Settings

Temperature	Time
37°C	20 min
25°C	20 min
4-12°C	Hold

Monarch Plasmid Miniprep Kit (For Purification of Plasmids from E. coli)

Follow this protocol to purify the desired plasmid which was inserted in the E.colicells. For more detailed instructions, please refer to the protocol [7].

Reagents

• Plasmid Resuspension Buffer (B1) (pink)
• Plasmid Lysis Buffer (B2) (blue/green)
• Plasmid Neutralization Buffer (B3) (yellow)
• Plasmid Wash Buffer 1
• Plasmid Wash Buffer 2
• DNA Elution Buffer or Nuclease-Free Water

Consumables

• Centrifuge tubes from the kit
• 1.5 mL Eppendorf tubes
• Maybe 2 mL Eppendorf tubes

Equipment

• Microcentrifuge

Procedure

Pellet Bacterial Culture

Pellet 4 mL bacterial culture by centrifugation for 30 seconds. Discard supernatant.

Resuspend Pellet

Resuspend pellet in 200 μL Plasmid Resuspension Buffer (B1). Vortex or pipet to ensure cells are completely resuspended.

Lyse Cells

Lyse cells by adding 200 μL Plasmid Lysis Buffer (B2). 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.

Neutralize Lysate

Neutralize the lysate by adding 400 μL of Plasmid Neutralization Buffer (B3) (yellow). Gently invert the tube until the color is uniformly yellow and a precipitate forms. Do not vortex! Incubate for 2 minutes.

Clarify Lysate

Clarify the lysate by spinning for 2–5 minutes at 16,000 x g.

Transfer Supernatant

Carefully transfer supernatant to the spin column and centrifuge for 1 minute. Discard flow-through.

Wash Column

Re-insert column in the collection tube and add 200 μL of Plasmid Wash Buffer 1. Centrifuge for 1 minute. Discarding the flow-through is optional.

Add 400 μL of Plasmid Wash Buffer 2 and centrifuge for 1 minute.

Elute DNA

Transfer column to a clean 1.5 mL microfuge tube. Add ≥ 30 μL DNA Elution Buffer to the center of the matrix. Wait for 1 minute, then spin for 1 minute to elute DNA.

DNA Digestion with Restriction Enzyme

In order to validate the correct assembly we digest our constructs with Restriction Enzymes.For restriction enzyme analysis and to visualize cutting sites, use the [8].

Materials

• DNA template
• Restriction Enzyme(s)
• PCR grade water (Nuclease-free water)
• Tubes (PCR tubes can work)
• Buffer (depends on the Enzyme(s) used)

Procedure

According to the restriction enzyme used, different buffers are required. Check the NEBcloner Restriction Enzyme Digestion for specific buffer recommendations.

Reaction Setup

Component	Amount
DNA	0.5 µg
Buffer	2.5 µL (1X)
Enzyme 1	1.0 µL
Enzyme 2 (optional)	1.0 µL
Nuclease-free Water	to 25 µL

Incubation

Incubate at 37°C for 1 hour.

In-Cell β-galactosidase Assay

This assay is based on a paper about a YES/YAS assay [1].Though this, we can testify the presence or not of the EDCs using yeast cells are scaffolds. The protocol reference also describes how to calculate the LacZ values, creating the standard curves, and calculating EEQ values.

Reagents

• Yeast strain carrying the pRR plasmids
• Glucose media broth (trp-)
• Galactose media broth (trp-)
• Sodium carbonate (105.9 g sodium carbonate in a final volume of 1 L)
• 1 M dithiothreitol (DTT). Should be frozen in 25 µL aliquots. CAUTION: use gloves and fume hood.
• 9.75 nM estradiol standard
• 227.5 nM estradiol standard
• LacZ buffer with ONPG
• 50% ethanol

Consumables

• Filters for sterilization of media
• Sterilized 96-well polypropylene plates (done by autoclaving stacks wrapped in foil)
• Sterile, adhesive, porous film for sealing the plates
• Non-sterilized 96-well polypropylene plates
• Sterile pipette tips (all sizes)
• Gloves

Equipment

• 250 ml Erlenmeyer flask
• Fume hood
• Incubator (set to 30 °C and 200 rpm)
• Spectrophotometer
• Plate spectrophotometer
• Sterile beakers
• 1000 µL multipipette
• 100 µL multipipette
• 1000 µL pipette
• 100 µL pipette
• 10 µL pipette

Procedure

Before Assay

1. Maintain active yeast cultures (with the pRR plasmids) by incubating yeast in filter-sterilized glucose media (trp-) at 30 °C. Subculture yeast weekly in fresh filter-sterilized glucose media.
2. Two nights (about 42 h) prior to preparing the 96-well assay plates, subculture yeast by adding 0.1 mL of active yeast cultures to 10 mL of filter-sterilized glucose media using sterile technique. Incubate at 30 °C for two nights. Shaking is not required if yeast are grown as shallow cultures in sterile 250 mL Erlenmeyer flasks.

Day 1 (Prepare the Assay Plates)

1. In a sterile beaker and using sterile technique, dilute yeast from step 3.2 to an optical density of 0.065 ±0.005 at 610 nm (OD610) in filter-sterilized galactose media.
2. Confirm optical density by pipetting 120 μL of each yeast sample in triplicate along with triplicate galactose blanks into a clear polystyrene plate (the plate does not need to be sterile). Use a plate spectrophotometer to verify that the diluted yeast culture has a net OD610 of 0.065 ±0.005. Subtract OD610 readings of galactose blanks from yeast OD610 readings to determine the net OD610 of the yeast. Prepare a final volume of at least 30 mL diluted yeast for each 96-well plate. NOTE: Spectrophotometer filters measuring absorbance between 595 and 610 nm can be used for this measurement. An approximate 1:6 v:v ratio of yeast:media typically yields an OD610 close to 0.065, so start by adding 4.5 mL yeast to 30 mL media and add more yeast or media as appropriate to achieve a net OD610 between 0.060 and 0.070.
3. Using sterile technique, add this diluted yeast suspension to the wells of a sterile, polypropylene, 96-well microplate according to the plate layout (see figure below). White wells are for the standard curve and light gray wells are for the controls.
4. During pipetting, keep the source yeast suspended by continuously swirling the container or mixing with the pipette. For this step, it is helpful to use a repeating pipettor with a sterile syringe tip.
5. Add 120 µL filter-sterilized galactose media to 3 additional wells (D4 - D6) according to the plate layout. These wells serve as media controls to account for absorbance contributed by the media alone.
6. Add 5 µL of vehicle (50% ethanol) to vehicle control wells containing the yeast suspension (D1 - D3) according to the plate layout. NOTE: Vehicle control wells are used to quantify background absorbance associated with yeast cells and media. Additionally, cytotoxicity or growth-promoting effects of test samples can be detected by comparing the OD610 values of test wells with those of vehicle control wells.
7. Construct a standard curve in each plate by serially diluting triplicates of hormone working standards in wells containing the yeast suspension according to the plate layout:
8. Begin by adding 5 µL of hormone standards to wells A1 through C1. Mix microwell contents by pipetting, and then serially dilute this suspension by moving 205 µL from wells A1 - C1 into wells A2 - C2. Repeat the mixing and transfer of 205 µL through column 12. Note: After the serial dilution is completed, discard 205 µL from wells A12 - C12. At the end of this step, all standard wells should contain 120 µL. For the serial dilution steps, it is helpful to use a multichannel pipettor with sterile tips.
9. Mix the contents of the sample, extraction control, and vehicle control wells by pipetting. Adjust volumes of these wells to 120 µL by removing 205 µL from each as described in the legend for. This can be done using a multichannel pipette.
10. Seal the plate(s) with a sterile, adhesive, porous film. Label plates and incubate for 17 h at 30 °C. The plate does not need to be shaken during incubation.

Day 2 (Processing of the Assay Plates)

1. After the 17 h incubation at 30 °C, remove plates from the incubator. Use a multichannel pipettor to mix the contents of each row of wells, and then transfer 50 µL of yeast suspension from each well to the corresponding well of a clear, polystyrene, 96-well microplate. Note: Polystyrene plates do not need to be sterile but should have a lid. Use new pipet tips for each row of wells to avoid cross-contaminating wells, although if pipetting across triplicates with an 8-tip multichannel pipette, tips only need to be changed between triplicate sets.
2. Add 20 µL of 1 M DTT to 20 mL of thawed, room-temperature LacZ buffer for a final concentration of 1 mM DTT. Mix this LacZ buffer well. CAUTION: Wear gloves and work with DTT in a hood.
3. Using either a multichannel pipette or repeating pipettor, add 200 µL of LacZ buffer containing 1 mM DTT to all wells of the polystyrene plate, and immediately measure and record the OD610 of all wells using a plate spectrophotometer.
4. Cover plates with a lid. Incubate plates containing yeast expressing ERα at 30 °C for 40 min and plates containing yeast expressing ERβ at 30 °C for 70 min. You can use another plate as the lid if you don't have one.
5. After incubating the plates, use a multichannel pipette or repeating pipettor to add 100 µL of sodium carbonate to each well. Sodium carbonate raises the pH and halts the β-galactosidase reaction.
6. Measure and record the OD405 (for ONPG) of all wells using a plate spectrophotometer.

This protocol is based on the Rosalind System Manual [9]. It is used to validate that our DNA constructs can be transcribed into an aptamer that in complex with the fluorophore DFHBI-IT emits fluorescence at the appropriate wavelength.

Materials

• 10x IVT buffer
• 100 mM Tris-buffered NTPs
• 2 U/uL inorganic pyrophosphatase
• 1 M IVT linear DNA template
• 40 mM DFHBI-IT
• H₂O
• 1 mg/mL T7 RNAP

Equipment

• 384 plate reader
• Nuclease-free PCR strips

Procedure

Setting the Microplate Reader

• the temperature to 37°C. The wavelengths should be set to 488 nm for excitation and 530 nm for emission.
• kinetic measurement, take reads every 3 minutes over the course of a 4-hour reaction.

Preparing the Master Mix

Mix the components to make a 15 µL master mix for each reaction:

Component	Final Conc.	Volume per reaction (µL)
10x IVT buffer	1x	2
100 mM Tris-buffered NTPs	11.4 mM	2.28
0.1 U/uL inorganic pyrophosphatase	0.015 U/uL	3
1 M IVT linear DNA template	5-50 nM	Vary (max is 7.68 µL)
40 mM DFHBI-IT	0.2 mM	0.1
H2O	-	Up to 15 µL

Note: The parameters should be optimized, as the concentration of DFHBI-IT greatly impacts transcription efficiency.

Preparing the Activator Mix

Incubate the master mix at 37°C while preparing the activator mix. Mix the components listed in a separate tube:

Component	Final Conc.	Amount per reaction (µL)
1 mg/mL T7 RNAP	10 ng/uL	2 µL
H2O	-	2.8 µL (up to 5 µL)

Reaction Setup

Ensure the microplate reader is ready, then add 5 µL of the activator mix to 15 µL of the master mix. Gently pipette up and down several times to mix, avoiding the introduction of air bubbles.

If setting up multiple reactions simultaneously, use a multichannel pipette to add the activator mix to the master mix to ensure that each reaction activates at the same time.

Immediately transfer the reaction onto a 384-well plate, seal the plate to prevent evaporation, and start the microplate reader run.

Protein Concentration

Because the protein concentration is low, we use protein concentrators [10] with a cutoff of 10K. Since our protein is 72 kD, we expect everything smaller than this size to pass through the filter in the column. This process allows us to reduce the volume of the protein sample used in our assay and switch to a buffer that is more compatible with the assay.

Materials

• PBS 1X
• Column[10]
• Centrifuge

Process

1. Place the sample into the concentrator sample chamber.
2. Cap, then place the concentrator assembly into the rotor with proper counterbalance.
3. Centrifuge the sample at 15,000 g for up to 7 minutes. A good strategy is to load the column with up to 500 μL of sample, then centrifuge until the volume is reduced to 50 μL.
4. Load an additional 450 μL of the desired buffer (PBS in this case) and centrifuge again.
5. Repeat this process as many times as possible to increase the efficiency.
6. Use a pipette tip to gently aspirate the retentate from the bottom and center of the sample chamber.

Cell Free assay with Receptor

In this experiment we test the capabilty of our desing to sence the presence of hormone in the solution

Materials

• Androgen receptor (0.14 μg/μl), MW = 72kDa
• IVT buffer
• NTPs
• 2 U/μl inorganic pyrophosphatase
• 40 mM DFHBI-IT
• Nuclease-free Water
• 1 mg/mL T7 RNAP

Procedure

1. Setting the microplate reader:
• Set the temperature to 37°C.
• Wavelengths should be set to 488 nm for excitation and 530 nm for emission.
• For kinetic measurement, take reads every 1 min over the course of a 2-h reaction.
2. Mix the components to make a 15 µL master mix for each reaction:
• Note: The parameters should be optimized, as the concentration of DFHBI-1T greatly impacts the transcription efficiency.
3. Incubate the master mix at 37°C for 1h 30 mins while preparing the activating reaction.
4. Mix the components listed below in a separate tube to make an activator mix:

Component	Concentration	Volume
1 mg/mL T7 RNAP	10 ng/μL	2 μL
H₂O	-	up to 5 μL

5. Ensure the microplate reader is ready to read, then add 5 μL of the activator mix to 15 μL of the master mix.
6. Gently pipette up and down several times to mix, avoiding the introduction of air bubbles.
7. If setting up multiple reactions simultaneously, use a multichannel pipette to ensure each reaction activates at the same time.
8. Immediately transfer the reaction onto a 384-well plate, seal the plate to prevent evaporation, and start the microplate reader run.

SDS-PAGE

This protocol describes the SDS-PAGE method for visualizing protein expression and purification, particularly for the Androgen Receptor. The method applies to other SDS-PAGE samples as well. Note: Acrylamide is carcinogenic, so always wear gloves when handling gels or buffers.

Materials:

Reagents:

• 4X Laemmli loading buffer
• 500 mM DTT
• Precision Plus Standard (Unstained)
• Pre-cast SDS-PAGE gel (12% acrylamide)
• 1X Tris/Glycine/SDS running buffer
• Bio-Safe Coomassie stain

Consumables:

• ThermoMixer at 95°C
• Staining tray

Sample Preparation:

Sample	Sample Volume/Amount	4X Laemmli Loading Buffer	500 mM DTT
Culture	500 μL	200 μL	2 μL
Crude protein (after centrifugation)	50 μL	200 μL	2 μL
Pellet (after centrifugation)	Approx. 2 μL	100 μL	2 μL

Procedure:

1. Prepare the SDS-PAGE sample mixes according to the table above.
2. Boil the samples at 95°C and 300 RPM for 10 minutes to ensure full protein denaturation.
3. Prepare a pre-cast gel.
4. Pour 1X SDS running buffer in the tank.
5. Load the gel: 5 μL unstained standard and 10 μL of each SDS sample.
6. Run the gel at 180V for 45 minutes.
7. Once the run is complete, open the gel casing with a gel opener and transfer the gel to a staining tray.
8. Wash the gel 3 times for 5 minutes each with demineralized water, using slow agitation.
9. Stain the gel for 1 hour with Coomassie stain.
10. Destain for approximately 1 hour with frequent water changes.
11. Once destained, place the gel in a plastic sheet and scan it.

Western Blotting with Turbo Blotter

Introduction:

This protocol describes Western blotting using a Turbo blotter. While other methods can be used, this is the fastest and simplest method.

Materials:

• SDS-PAGE gel (unstained) using Dual color protein standard.
• Mini transfer pack.
• Turbo blotter.
• Skim milk buffer (2% skim milk powder, 1X TBS).
• Anti-His antibodies (HRP-conjugated).
• 15 mL Falcon tube.
• AEC staining kit.

Procedure:

Blotting:

1. Load the gel as with SDS-PAGE but use Dual color protein standard. Run the gel until the blue color exits completely.
2. Place the bottom stack (with membrane) on the cassette base, followed by the gel, then the top stack.
3. Close and lock the cassette lid. Insert cassette into the instrument and start transfer.
4. Run the block that is loaded.

Washing the Membrane:

1. Transfer the membrane to a fitting container without color stains and block it in skim milk buffer for 1 hour with slow agitation.
2. Discard the buffer and incubate the membrane in skim milk buffer with antibodies for 1 hour, again with slow agitation.
3. Discard the buffer and wash the membrane 3 times for 15 minutes each with slow agitation in skim milk buffer.
4. Rinse the membrane briefly in demineralized water.

Staining:

1. Prepare the AEC staining solution using the AEC staining kit:
• Add 4 mL demineralized water to a 15 mL Falcon tube.
• Add 2 drops (60 μL) of acetate buffer.
• Add 1 drop (30 μL) of AEC chromogen.
• Add 1 drop (30 μL) of hydrogen peroxide solution.
2. Mix gently by inverting a few times.
3. Pour the staining solution over the membrane, ensuring it fully covers the upward side.
4. Leave the membrane to stain for 5-10 minutes (or until sufficient staining is achieved) under tinfoil in the fume hood.
5. Pour the solution out and rinse with excess demineralized water.
6. Scan the membrane in the same way as for gels.