Methods

Plasmid Construction Go to Results Results

For plasmid construction, restriction digestion cloning was employed in all cases except for the integration of mCherry into pAG-304, which was generated using Gateway cloning. For this construct, the original plasmid pENTRY-ymCherry served as the template. During the LR reaction, the mCherry insert from pENTRY-ymCherry was recombined into the destination vector pAG-304. All other DNA fragments were inserted into their respective destination vectors via restriction digestion. The oligonucleotide fragments used in our project were all designed by us using existing BioBricks and other parts found in papers. The oligos were then ordered and synthesised by IDT, GenScript, or Twist Bioscience. The cloning workflow involved digestion of both fragments and vectors with the appropriate restriction enzymes (Protocol 1), followed by gel electrophoresis to prove successful cutting (Protocol 2). For a few restriction enzymes (NOT1, BSU36I, BstX1, XHO1) the restriction digestion initially failed, new respective enzymes were validated by control digestions and gel electrophoresis to confirm their activity. Here 1% agarose gels and 90V/60min running conditions were used. As a marker 1kb marker GeneRuler (NEB) was used. All our gels were analyzed under UV light. Following successful digestion, the fragments and vectors were combined and incubated at 37 °C for 1 h prior to ligation. For a the gRNA inserts for both variants of our Cas13 a Ligation was necessary to combine multiple gRNA oligonucleotides. During our Project we constructed the following Constructs that way:

Fragment Vector Purpose Cutting Enzyme Auxotrophy Marker
GFP-E9-Tetra Im2-MiniCas13.X1 pAG-416 This is the first version of our completed TRAPS System with Cas13 Sac1, Mlu1 URA3
GFP-E9-Tetra Im2-RfxCas13d pAG-416 This is the second version of our completed TRAPS System with Cas13 Sac1, Mlu1 URA3
gRNAsMiniCas13.X1 pAG-305 This Construct contains the gRNAs for the MiniCas13.X1 version of our TRAPS system Mlu1, Sac1 LEU
gRNAsRfxCas13d pAG-305 This Construct contains the gRNAs for the RfxCas13d version of our TRAPS system Mlu1, Sac1 LEU
GALymCherry pAG-304 mChery RNA is the first target for our RNA detection system and is under a conditional promotor to make working with it easy Gateway Cloning TRP
Pumby1 pAG-416 Pumby is a RNA binding protein that binds to a specific 8nt sequence. We use it to bind our target RNA In this Construct half of the TRAPS system with Pumbys is present Sac1, Mlu1 URA3
Pumby2 pAG-305 In this Construct half of the TRAPS system with Pumbys is present. Together with the pAG416-Pumby1 Construct they form the completed TRAPS system with Pumbys instead of Cas13. Mlu1, Sac1 LEU
GFP-E9-Tetra pAG-416 This Construct is part of the TRAPS system and used to test the interaction of the Tetramerization domains Sac1, Sph2 URA3
Im2-MiniCas13.X1 pAG-416 This Construct is part of the TRAPS system and used to test if the Im2-MiniCas13.X1 form aggregates. Mlu1, Sph2 URA3

Resulting in the following strains:

Strain Genotype Selection
E. coli DH5α fhuA2 Δ(argF-lacZ)U169 phoA glnV44 Φ80 Δ(lacZ)M15 gyrA96 recA1 relA1 endA1 thi-1 hsdR17 -
E.coli DH5α pAG304-GALymCherry Ampicillin
E.coli DH5α pAG416-GFP-E9-Tetra-Im2-MiniCas13.X1 Ampicillin
E.coli DH5α pAG305-gRNAsMiniCas13.X1 Ampicillin
E.coli DH5α pAG416-GFP-E9-Tetra-Im2-RfxCas13d Ampicillin
E.coli DH5α pAG305-gRNAsRfxCas13d Ampicillin
E.coli DH5α pAG416-Pumby1 Ampicillin
E.coli DH5α pAG305-Pumby2 Ampicillin
E.coli DH5α pAG416-GFP-E9-Tetra Ampicillin
E.coli DH5α pAG416-Im2-MiniCas13.X1 Ampicillin
E.coli DH5α pAG416-Im2-RfxCas13d Ampicillin
S. cerevisiae W303 MATa/MATα {leu2-3,112 trp1-1 can1-100 ura3-1 ade2-1 his3-11,15} [phi+] -
S. cerevisiae W303 pAG304-GALymCherry TRP
S. cerevisiae W303 pAG416-GFP-E9-Tetra-Im2-MiniCas13.X1 + pAG305-gRNAsMiniCas13.X1 + pAG304-GALymCherry URA3, LEU, TRP
S. cerevisiae W303 pAG416-GFP-E9-Tetra-Im2-RfxCas13d + pAG305-gRNAsRfxCas13d + pAG304-GALymCherry URA3, LEU, TRP
S. cerevisiae W303 pAG416-Pumby1 + pAG305-Pumby2 + pAG304-GALymCherry URA3, LEU, TRP
S. cerevisiae W303 pAG416-GFP-E9-Tetra + pAG304-GALymCherry URA3, TRP
S. cerevisiae W303 pAG416-Im2-MiniCas13.X1 + pAG304-GALymCherry URA3, TRP

Transformations Go to Results Results

All plasmids were initially transformed into E. coli DH5α cells for propagation using the chemical transformation method (see Protocol E. coli Transformation) and grown on selective plates with ampicillin as a selective marker. Single colonies were grown in overnight cultures (4mL 100µg/ml ampicillin) and plasmids were then isolated from those cultures of the transformed strains using the Monarch Spin Plasmid Miniprep Kit from NEB. The DNA concentration was afterwards measured with a NanoPhotometer and to confirm the correct size a test restriction of the plasmid was performed and subjected to an agarose gel-electrophoresis. An empty vector sample was used as negative control and loaded as well to the agarose gel.
The purified plasmids intended for further use were subsequently prepared and sent to Microsynth for sanger sequencing. After confirming the desired sequences, the plasmids were introduced into our S. cerevisiae strain W303 (see Protocol Yeast Transformation). The final strains were kept as glycerol (25 % (v/v)) stocks at -80°C.
Over the course of the project, we optimized the yeast transformation procedure. Initially, transformations were carried out without the use of sorbitol using a standard yeast transformation protocol, which resulted in low success rates. Further investigation led us to adopt a sorbitol-containing protocol, which we refined throughout the project, ultimately achieving high transformation efficiencies. The final protocol is detailed in the Protocols section.
The vectors pAG304 and pAG305 are genomically integrating vectors and were therefor genomically integrated into the Tryptophan (pAG304) and Leucin (pAG305) locus of the W303 yeast cells. Therefore, the Plasmid was first linearized with the appropriate restriction enzyme and then used in the transformation reaction. Successful transformation was verified by plating the cells on selective media lacking the appropriate auxotrophic markers corresponding to the plasmids used (see Table 1 for details on auxotrophic markers).

Microscopy Go to Results Results

Before microscopy we prepared the cells (see protocol Microscopy). We either microscoped the samples from a 384 Well Plate, a sliding dish or fixed the cells for long time measurements and FRAP. All microscopies were performed with a Nikon ECLIPSE Ti2 microscope, equipped with a photometrics prime 95B camera (Teledyne Technologies Incorporated, CA, USA), at a temperature of 25 °C. For imaging the yeast cells, a 1000x magnification was used. Imaging always included Bright Field and Fluorophore images. Imaging of the fluorophores was enabled by exciting RFP and GFP.
All our created strains were microscoped always induced cells expressing mCherry RNA compared to uninduced cells as a control. (see Protocol Inducing of the GAL1 Promoter). All measurements were done on at least three times on at least 4 different spots per sample.
For fluorescence microscopy image analysis, we used a Python pipeline (see Software). We first employed Cellpose 2.0 and their cyto3 model in the brightfield channel to obtain the segmentation labels of all the yeast cells. Then, we extracted the fluorescence intensities within these segmentation labels from the fluorescence channels. For quantification, we calculated the total cell fluorescence by multiplying the corrected mean fluorescence intensity of a cell with the total amount of pixels in the cell region. The corrected mean is calculated as mean intensity of the region minus the mean fluorescence intensity of all the non-cell pixels.

Protein Analysis Go to Results Results

Yeast cell culture of cell lines W303-mCherry-Cas13X.1, W303-mCherry-Cas13X.1-gRNAs, W303-RfxCas13.d, W303-mCherry-305-Pumby1 and W303-mCherry-416-Pumby were grown in SD -TRP -URA Medium, while W303 wild type (WT) culture was grown in YPD medium. All cultures were grown at 30°C with shaking at 180 rpm until they reached an OD₆₀₀ of 0.5 to ensure the cells had reached the exponential phase. The cultures were harvested at a volume equivalent to 10 OD₆₀₀₀ units, calculated from OD₆₀₀ measurements of a 1 ml sample. The 10 OD₆₀₀ samples were then spun down at 2,000 rcf for 4 min. The cells of every sample were washed in 1 ml of ice cold water and centrifuged at 3,000 rpm for 2 min at 4°C. The cell pellets were resuspended in 800 µl lysis buffer and lysed by bead beating with 500 µl silica beads shaken at 30 oscillations/s for 20 min at 4°C. Afterwards the lysate was centrifuged for 1 min at 500 rpm at 4 °C to reduce foam. 300 µl of lysate were taken and centrifuged at 4°C for 30 sec at 3,000 rcf to get the crude extract. 61 µl were taken from the crude extract and stored as the total fraction (T). 200 µl of the crude extract were further centrifuged at 4°C for 5 min at 20,000 rcf. 121 µl were taken as the Supernatant (S).
The protein concentration of the T and S samples were measured with Detergent Compatible (DC) Assay. Bovine Serum Albumin (BSA) was used as the standard protein. A standard curve was generated from a dilution series of BSA covering the concentrations 2.58-0.00 mg/ml in a consecutive 7 step 1:2 dilution. In a 96-well plate 1 µl of the T and S sample as well as each dilution step of the BSA standard was diluted in 4 µl water and mixed with 25 µl SA-mix (50:1 ratio of A to S). After adding 200 µl reagent B to each sample, they were incubated at room temperature in the dark for 15 min. Absorbance was measured at 750 nm using the TECAN Spark® microplate reader.
The T and S samples of the different constructs were split in half and one half was heated at 95°C for 5 minutes (Heated), the other half was not heated (Non-Heated). An SDS-PAGE of the Cas13- and Pumby-constructs was performed with a 12 % SDS-Gel each. An equal amount of protein of each sample and prestained protein marker VI (10-245 kDa) was loaded on the respective Cas13 and Pumby SDS gels in duplicates, so after the blotting the membranes could be cut in half and each half would include a duplicate of the samples and a ladder. The gels were run at a constant voltage of 200 V for 40 min. A scan of the gels was performed using the Cy2 525BP20 and IRshort 720BP20 emission filters of the Amersham Typhoon to detect GFP and the ladder. Proteins were transferred to nitrocellulose membranes at 200 mA for two hours, followed by blocking with 5% (w/v) in PBS-T for 1h. The membranes were washed four times for 5 min in PBS-T. An antibody staining with the primary antibodies fitting the different tags of our target constructs was performed overnight at 4°C. For the Cas13-constructs an α-myc-tag antibody was used. For the Pumby constructs an α-FLAG-tag antibody and an α-HA-tag antibody were used. Each respective primary antibody were mouse host and added to the membranes in a 1:5000 dilution in 2% skim milk (w/v) PBS-T and incubated overnight at 4°C. The membranes were washed four times for 5 min in PBS-T. Both the Cas13 and the Pumby membranes were then cut at the duplicate line so each half contained the target samples and a ladder. The secondary antibody Alexa-Flurophor 488 anti-mouse diluted 1:10,000 in 5% skim milk was added and incubated for 1h at room temperature. After conducting a first fluorescent measurement of the membrane, the immunostaining protocol was repeated with a second primary antibody against the loading control PGK1 (mouse, 1:5000 dilution in 2% skim milk (w/v) PBS-T) followed by the incubation with secondary antibody Alexa-Flurophor 488 anti-mouse (1:10,000 in 5% skim milk).
The membranes were washed four times for 5 minutes in PBS-T. They were then scanned using the Cy2 525BP20 and IRshort 720BP20 emission filters of the Amersham Typhoon. Densitometric analysis was performed using the Gel Analysis tool in Fiji (doi:10.1038/nmeth.2019). Band intensities were quantified by selecting rectangular regions of interest (ROIs) around each lane and subtracting the background signal. The intensity of the target protein construct bands was normalized to the corresponding loading control (PGK1).

Protocols

Restriction Digestion

Abstract

Restriction Digestion is used to create defined DNA fragments by cutting plasmids and other fragments with restriction endonucleases. Here we use a protocol adapted for HF-Endonucleases.

Material

  • Plasmid DNA or DNA fragment
  • Restriction enzyme(s) (e.g., HF-Endonucleases)
  • rCutSmart Buffer (10X)
  • Nuclease-free water
  • Microcentrifuge tubes
  • Pipettes and tips
  • Thermoblock or water bath (37 °C)

Protocol

Composition of restriction digestion reaction:

Component Volume/Amount
DNA 1 µg
Restriction enzyme 1 µl
rCutSmart Buffer 10X 5 µl
Nuclease free water up to 50 µl

Mix and incubate at 37 °C for 1 h.

PCR

Abstract

PCR (Polymerase Chain Reaction) is a method to amplify DNA fragments. Here we use Q5 High-Fidelity DNA Polymerase for high accuracy.

Material

  • Template DNA
  • Forward and reverse primers (10 µM)
  • dNTP mix (10 mM each)
  • Q5 High-Fidelity DNA Polymerase
  • Q5 Reaction Buffer (5X)
  • Nuclease-free water
  • Microcentrifuge tubes
  • Pipettes and tips
  • Thermocycler

Protocol

Composition of PCR reaction:

Component Volume/Amount
Template DNA 1 µl
Forward Primer (10 µM) 0.5 µl
Reverse Primer (10 µM) 0.5 µl
dNTPs (10 mM each) 1 µl
Q5 High-Fidelity DNA Polymerase 0.5 µl
Q5 Reaction Buffer (5X) 5 µl
Nuclease free water up to 25 µl

Mix and run the following program in a thermocycler:
1. Initial denaturation: 98°C for 30s
2. 25-35 cycles of:
a. Denaturation: 98°C for 10s
b. Annealing: X°C for 20s (X = Tm of primers - 3°C)
c. Extension: 72°C for Y min (Y = 30s per kb of target length)
3. Final extension: 72°C for 2min
4. Hold at 4°C

Ligation

Abstract

Ligation is used to join DNA fragments together. Here we use T4 DNA Ligase to ligate fragments with compatible ends.

Material

  • Linearized vector DNA
  • Insert DNA fragment
  • T4 DNA Ligase
  • T4 DNA Ligase Buffer (10X)
  • Nuclease-free water
  • Microcentrifuge tubes
  • Pipettes and tips
  • Thermoblock or water bath (37 °C)

Protocol

Composition of ligation reaction:

Component Volume/Amount
Vector 50 ng
Insert (3:1 molar ratio) X ng
T4 DNA Ligase 1 µl
T4 DNA Ligase Buffer 10X 2 µl
Nuclease free water up to 20 µl

Mix and incubate at 37 °C for 1.5 h.

Agarose Gel electrophoresis

Abstract

Agarose Gel electrophoresis is a simple method for analysing DNA Fragments. It is used to separate different DNA fragments with different lengths from each other. Here we show Gel electrophoresis with agarose gels stained with Ethidium Bromide.

Material

  • Agarose
  • 1x TAE buffer
  • Ethidium Bromide (or alternative DNA stain)
  • Gel casting tray and comb
  • DNA samples
  • Purple gel loading dye (6x)
  • 1kb extend DNA Ladder (NEB)
  • Electrophoresis chamber and power supply
  • Gel imager with UV light
  • Pipettes and tips

Protocol

1. Prepare 1% agarose gel:
• Weigh 1g agarose and add it to 100ml 1x TAE buffer.
• Heat the solution in a microwave until the agarose is completely dissolved.
• Cool the solution to about 60°C and add 5µl of Ethidium Bromide.
• Pour the solution into a gel casting tray with a comb and let it solidify for at least 30 minutes.
• After solidification pour 1x TAE Buffer into the tray.
2. Prepare samples:
• For sample Preparation add purple gel loading dye 6x at a final concentration of 1x to each sample.
• Load each sample in a separate well and alongside 5µl 1kb extend DNA Ladder (NEB).
3. Run the gel:
• Let the gel run at 90V for 60-90min depending on the loaded samples and agarose concentration (1-2%).
• Image the gel in a gel imager under UV light

E. coliTransformation

Abstract

E. coli Transformation is a method to introduce foreign DNA into E. coli cells. Here we use chemically competent cells and a heat shock protocol.

Material

  • Chemically competent E. coli (e.g., DH5α)
  • Plasmid DNA
  • ddH2O
  • 5x KCM buffer
  • LB medium
  • Microcentrifuge tubes
  • Pipettes and tips
  • Incubator (37 °C)
  • Shaker
  • Water bath (42 °C)
  • Agar plates with appropriate antibiotic

Protocol

Mix:

Component Volume/Amount
dH2O 7 µl
5x KCM 2 µl
DNA 1 µl

•Set the mixture on ice for 10min
•Add 10µl chemical competent E. coli (DH5α)
•Set it on ice for another 20min
•Heat shock the cells at 42°C for 1min, 300 RPM
•Add 900µl LB for regeneration
•Incubate the cells for 45min at 37°C
•Spin the cells down and resuspend them in ddH2O before plating them

Yeast Transformation

Abstract

Yeast transformation is a widely used method to introduce plasmid DNA into Saccharomyces cerevisiae. This protocol is based on the lithium acetate/PEG method, which relies on chemical permeabilization of the yeast cell wall.

Material

  • S. cerevisiae cells
  • Linearized plasmid DNA
  • Salmon sperm DNA (ssDNA)
  • Sorbitol/LiAc solution (1M/1M)
  • PEG
  • DMSO
  • YPD medium
  • ddH2O
  • Selective agar plates
  • Microcentrifuge tubes
  • Pipettes and tips
  • Incubator (30 °C)
  • Shaker
  • Centrifuge

Protocol

1. Preparation of yeast culture:
• Grow 5 ml overnight pre-culture to saturation
• Dilute to OD₆₀₀ = 0.1 in 50 ml of fresh medium in 250 ml flask
• Incubate at 30 °C for 3–4 h to reach OD₆₀₀ ≈ 0.5-0.7
2. Spin cells at 2000x g for 3 min
3. Wash once with 25 ml of Sorbitol/LiAc (1M/1M) solution,
4. Spin cells at 2000x g for 3min and discard supernatant.
5. Resuspend cells in the rest drops of liquid (usually it is 400-500 microliters)

6. Assemble transformation reaction:

Component Volume/Amount
Linearized plasmid ~25 µl (1 µg)
ssDNA (salmon sperm DNA) 15 µl
Yeast cells (resuspended in rest drops Sorbitol/LiAc) 100 µl
PEG 280 µl

7. Assemble transformation reaction for a negative control with everything excluding the linearized plasmid
8. Mix well and incubate at room temperature for 45-60min
9. Add 40µl DMSO to the mixture
10. Incubate at 42°C for 15min, 300RPM
11. Add 1ml of YPD and mix well 11. Add 1ml of YPD and mix well
12. Allowing the cells to recover for 1-2h at 30°C with shaking at 200rpm is optional.
13. Spin at 4000rpm for 5min and discard the
14. Resuspend the cell pellet in 200µl ddh3O
15. Plate on the appropriate selective media.
16. Incubate plate at 30°C for 2-3 days until colonies appear.

Inducing of the GAL1 Promoter

Abstract

Conditional promoters enable precise regulation of gene expression in response to defined stimuli. In our case the GAL1 promotor is used, which is a very strong promoter that is activated through galactose

Material

  • Yeast strains with GAL1 promoter constructs
  • SC medium with glucose
  • SC medium with 2% raffinose
  • SC medium with 2% galactose (or raffinose + galactose)
  • Sterile water
  • Microcentrifuge tubes
  • Pipettes and tips
  • Incubator (30 °C)
  • Shaker
  • Spectrophotometer (for OD600 measurement)

Protocol

Pick a single colony and bring it into the appropriate media containing Glucose.
1. Incubate overnight at 30 °C with shaking at 200 rpm.
2. Removal of Glucose:
• Pellet 1–5 mL of the overnight culture by centrifugation at 3,000 × g for 5 minutes.
• Wash the cell pellet twice with sterile water or SC medium lacking a carbon source to eliminate residual glucose.
3. Pre-induction in Raffinose:
• Resuspend the washed cells in SC medium supplemented with 2% raffinose.
• Adjust the cell density to OD₆₀₀ = 0.2–0.4.
• Incubate at 30 °C with shaking (200 rpm) for 4–6 hours, or until cells reach mid-log phase (OD₆₀₀ ~0.6–0.8).
• This step allows for derepression of GAL promoters without activation.
4. Induction with Galactose:
• Add galactose to the raffinose culture to a final concentration of 2% (w/v), or alternatively, pellet and resuspend the cells in fresh SC medium containing 2% raffinose and 2% galactose.
• Continue incubation at 30 °C with shaking.
• Induction times may vary

Microscopy

Abstract

Microscopy is used to visualize protein expression and localization in yeast cells. This protocol describes the preparation of yeast samples and the use of fluorescence microscopy to analyze induced and control cultures.

Material

  • Yeast cultures (induced and control)
  • Microscope slides and cover slips
  • Fluorescence microscope with appropriate filters
  • Well plates (optional, for multiple samples)
  • Pipettes and tips

Protocol

Preparation of Yeast cells:
•Set the OD₆₀₀ of your yeast culture to ~0.3
•Take 20µl of the induced culture and put it on a microscope slide and add a cover slip.
•Optional: If you want to analyze multiple samples at the same time add 20µl of each sample to a well of a appropriate well plate.
Image the cells with a fluorescence microscope using the appropriate filters for your fluorophore.
Analyze the images using image analysis software.

FRAP

Abstract

Fluorescence Recovery After Photobleaching (FRAP) is a technique used to study the dynamics of fluorescently labeled molecules within living cells. This protocol outlines the steps for performing FRAP experiments on yeast cells expressing fluorescent proteins. For FRAP it is important to have immobilized cells. Therefore, we show a immobilization protocol using Concanavalin A.

Material

  • Yeast cultures expressing fluorescent proteins
  • Fluorescence microscope with FRAP capabilities
  • Microscope slides and cover slips
  • NaOH
  • Concanavalin A solution
  • Imaging software
  • Pipettes and tips

Protocol

1. Preparation of Yeast cells: •Set the OD₆₀₀ of your yeast culture to ~0.3 2. Immobilize the yeast cells to the cover slip: •Wash dish with 1M NaOH.
•Add Concanavalin A solution to the dish and coat for 20min while gently shaking.
•2.2ml 1x PBS
•125µl 0.2M K2HPO4
•2.5µl 1M CaCl2
•5µl 10% NaN3
•250µl 5mg/ml Concanavalin A
•Rinse dish 3 times with ddh3O
•Do not let your dish dry after the coating as this will give you a high background
•Add your yeast suspension to the dish and let it sit for 10min
3. Image the cells: Image the cells with a fluorescence microscope using the appropriate filters for your fluorophore.
Analyze the images using image analysis software.

Yeast Lysate Preparation & Protein Quantification

Abstract

This protocol describes the preparation of yeast lysates for protein analysis and subsequent quantification using the DC Protein Assay (Bio-Rad). The procedure includes cell growth, lysis by bead beating, and quantification against a BSA standard curve.

Protocol

  1. Grow yeast culture: Inoculate an overnight culture. Measure OD600 (dilute 1:10 if necessary). Dilute to OD600 ≈ 0.2 and incubate until OD600 ≈ 0.5–0.7 (typically ~4 h in SD -TRP -URA medium).
  2. Harvest cells: Collect enough culture for a total OD of 10 (e.g., for OD 0.52: 10/0.52 = 19.2 ml). Spin down at 2000g, 4 min.
  3. Wash cells: Discard supernatant, resuspend pellet in 4°C autoclaved water, transfer to 1.5 ml tube, spin at 3000 rpm, 2 min, 4°C.
  4. Prepare for lysis: Prepare a 1.5 ml tube with 500 µl silica beads. Remove supernatant from washed cells.
  5. Cell lysis: Resuspend pellet in 800 µl complete lysis buffer (with DTT and protease inhibitor), transfer to bead tube. Bead-beat at 30 oscillations/s for 20 min at 4°C.
  6. Clarify lysate: Spin at 500 rpm, 1 min, 4°C to reduce foam. Transfer 200 µl lysate to new tube (avoid beads/foam). Spin at 3000g, 30 sec, 4°C to obtain crude extract.
  7. Prepare samples: For "Total" (T): Take 61 µl crude extract into new tube (1 µl for quantification).
    For "Supernatant" (S): Take 120 µl crude extract, spin at 20,000g, 5 min, 4°C, then take 61 µl supernatant (1 µl for quantification).
  8. Add sample buffer: Add 20 µl sample buffer (with 10% β-mercaptoethanol) to T and S after taking 1 µl for quantification. Freeze samples for later SDS-PAGE.
  9. Protein quantification (DC Protein Assay, Bio-Rad):
    • Prepare BSA standard curve: 2.58 mg/ml, 1.29, 0.64, 0.32, 0.16, 0.08, 0.04, 0.00 mg/ml (serial 1:2 dilutions in lysis buffer).
    • Prepare S and A mixture (50:1 ratio of A to S).
    • Dilute 1 µl sample in 4 µl water.
    • Load samples and standards into a 96-well plate.
    • Add 25 µl SA mixture to each well.
    • Add 200 µl B reagent to each well, mix gently (avoid bubbles).
    • Incubate 15 min at room temperature in the dark (color changes from orange to blue).
    • Measure absorbance at 750 nm (e.g., with Tecan plate reader).
    • Determine protein concentration using the BSA standard curve.
  10. Notes:
    • Quantification and freezing/loading the gel should be done ASAP to avoid protein degradation.

SDS-PAGE

Abstract

Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE) is a technique used to separate proteins based on their molecular weight. This protocol describes the steps for preparing and running SDS-PAGE gels to analyze protein expression and purity.

Protocol

1. Gel Casting

If using pre-cast gels, skip this section.
Note: In our experiments, self-cast gels with 10–12.5% acrylamide were used (Mini-PROTEAN Tetra Cell system).

Separating Gel (10 ml, 12.5% acrylamide, 1 mm thickness preferred):
Component Volume
H₂O3.7 ml
1.5 M Tris-HCl pH 8.83.0 ml
10% SDS178 µl
30% Acrylamide5.0 ml
10% APS79 µl*
TEMED8 µl*

* Important: Add APS and TEMED last, mix quickly, and pour immediately.
Allow separating gel to polymerize (check solidification with leftover mix in Falcon tube).

Stacking Gel (4%, 1 gel):
Component Volume
H₂O1.8 ml
1 M Tris-HCl pH 6.50.8 ml
10% SDS96 µl
30% Acrylamide0.5 ml
10% APS32 µl*
TEMED4 µl*

* Important: Add APS and TEMED last, mix quickly, insert comb, and allow to polymerize.
Cast gels can be stored in SDS running buffer at 4 °C or used directly.

2. Sample Preparation

•Load 10µg protein per lane (calculated from quantification).
•Load 4µl prestained protein ladder per gel.
•Load samples directly without additional denaturation steps.

3. Sample Loading

•Insert gels into the running chamber. Push the glass plates to the rim to avoid leaks.
•If running only one gel, insert a dummy plate in the second slot.
•Fill inner chamber with fresh SDS running buffer. Once confirmed tight, fill the outer chamber halfway with used buffer.
•After removing the comb, rinse wells with running buffer.
•Load marker and samples as indicated above.

4. Running the Gel

•Run the gel at a Constant voltage: 200 V, 40 min
•Run until dye front reaches the end of the gel (slightly running out).
•After run: recycle buffer into used buffer flask.

5. Post-Run

•Open cassette carefully with spatula under running water.
•Transfer gel into container with water.
Scanning (Typhoon): •Select appropriate channels for scanning.
•Perform prescan: if signal appears pink = oversaturated → reduce voltage.
•Otherwise, lower voltage slightly before final scan.
•Save scan file!
•Clean glass plates carefully, ensure no lint remains. If organic solvents were used, do not place gel directly on glass → use foil support instead.

Western Blot

Abstract

Western blotting is a technique used to detect specific proteins in a sample after separation by SDS-PAGE and transfer to a membrane.

Protocol

  1. Run SDS-PAGE: Perform SDS-PAGE as described in the "SDS-Page Protocol".
  2. Prepare transfer sandwich: Prepare a container with transfer buffer.
    Assemble the sandwich in the following order (from bottom to top):
    • Black side of clamping cassette (down)
    • Sponge
    • Whatman paper
    • Gel
    • Nitrocellulose membrane (roll flat with roller!)
    • Whatman paper
    • Sponge
    Note: Ensure the gel is oriented so that the ladder will be on the left side after transfer.
  3. Close cassette: Place the white side of the clamping cassette on top and close it.
  4. Transfer proteins: Place the sandwich in the container with transfer buffer.
    Run the transfer for 2 hours at 200 mA.
    Note: For proteins >300 kDa, extend transfer time beyond 2 hours.
  5. Blocking: Incubate the membrane for 1 hour in blocking buffer.
  6. Primary antibody incubation: Dilute the primary antibody in blocking buffer.
    Incubate the membrane overnight at 4 °C with gentle shaking.
  7. Wash membrane: Wash the membrane 3 times for 10 minutes each with TBST.
  8. Secondary antibody incubation: Dilute the secondary antibody in blocking buffer.
    Incubate the membrane for 1 hour at room temperature with gentle shaking.
  9. Wash membrane: Wash the membrane 3 times for 10 minutes each with TBST.
  10. Imaging: Image the membrane using the appropriate imaging technique.

Materials

Chemical compounds used in this study were purchased from Merck (Darmstadt, Germany), Roche Diagnostics (Basel, Switzerland), Carl Roth (Karlsruhe, Germany) and Qiagen (Venlo, Netherlands). Restriction enzymes were purchased from New England Biolabs (NEB, Ipswitch, USA). Water used for buffers, solutions and media was filtered using Milli-Q Ultrapure water systems (Merck)

Chemicals and Reagents

  • 10% APS
  • 10% SDS
  • 10x TAE Buffer
  • 1.5 M Tris-HCl pH 8.8
  • 1 kb DNA ladder Generuler (NEB)
  • 1 kb extend DNA Ladder (NEB)
  • 1M Tris-HCl pH 6.5
  • 30% Acrylamide
  • 5x KCM buffer
  • Agarose
  • Agar plates with antibiotic selection
  • Alexa-Fluor 488 anti-mouse secondary antibody (1:10,000)
  • Antibiotics: Ampicillin, Kanamycin
  • Blocking Buffer: 5% Milk powder in PBST
  • Bovine Serum Albumin (BSA)
  • Complete lysis buffer (with DTT and protease inhibitor)
  • β-Mercaptoethanol
  • Coating solution:
    • 1x PBS
    • 0.2M K2HPO4
    • 1M CaCl2
    • 10% NaN3
    • 5mg/ml Concanavalin A
  • Competent E. coli cells (DH5α)
  • ddh3O
  • DMSO
  • dNTPs
  • DTT
  • EDT
  • Ethanol
  • Ethidium bromide
  • Forward and reverse primers (10 µM)
  • Galactose, Glucose, Raffinose
  • Gel casting tray and comb
  • Gel loading dye, Purple (NEB)
  • HEPES (pH 7.6)
  • High-Fidelity Buffer
  • Ice cold water
  • Insert DNA fragment
  • KCM
  • LB medium
  • LiAc
  • Ligation Buffer
  • Linearized plasmid DNA
  • Linearized vector DNA
  • Lysis buffer
  • Magnesium acetate
  • NaOH
  • Nitrocellulose membranes
  • Nuclease-free water
  • PBS
  • PBS-T (PBS with Tween-20)
  • PEG 3350
  • Plasmid DNA
  • Potassium acetate
  • Purple gel loading dye (6x)
  • Q5 High-Fidelity DNA Polymerase
  • Q5 Reaction Buffer (5x)
  • Primary antibodies:
    • 9E10 α-myc mouse
    • α-FLAG M2 mouse
    • α-HA.11 mouse
    • α-PGK1 mouse
  • Protein Marker VI (10 - 245 kDa)
  • Protease inhibitors:
    • 0.1M PMSF
    • 0.5M Benzamidine
    • Aprotinin
    • Chymostatin
    • E-64
    • Leupeptin
    • Pepstatin
  • rCutSmart Buffer (NEB)
  • Restriction enzymes (HF): BstXI, EcoRI, MfeI, MluI, SacI, SphI, XhoI (NEB)
  • Running buffer (1x):
    • 0.025 M Tris base
    • 0.192 M Glycine
    • 0.1% SDS
  • S. cerevisiae cells (strain W303)
  • Salmon sperm DNA (ssDNA)
  • Sample buffer (with 10% β-mercaptoethanol)
  • SC medium with glucose/raffinose/galactose
  • Selective agar plates
  • Skim milk (2% and 5% w/v in PBS-T)
  • Sorbitol
  • Sorbitol/LiAc solution (1M/1M)
  • Sterile water
  • T4 DNA Ligase (NEB)
  • T4 DNA Ligase Buffer (10X)
  • Taq polymerase (Thermofisher)
  • TBST
  • TEMED
  • Template DNA
  • Transfer Buffer:
    • 0.025 M Tris base
    • 0.192 M Glycine
    • 20% Methanol
  • Triton-X
  • Whatman paper
  • Yeast cultures (induced and control)
  • Yeast synthetic dropout media (-LEU, -TRP, -URA)
  • YPD medium

Kits

  • DC Protein Assay Kit (Bio-Rad)
  • Gel Extraction Kit (Qiagen, NEB)
  • Monarch Spin Plasmid Miniprep Kit (NEB)
  • PCR Clean Up Kit (Qiagen, NEB)

Equipment & Instruments

Laboratory Equipment

  • 96-well plate
  • Amersham Typhoon laser scanner
  • Bead beating device (30 oscillations/s)
  • Cover slips
  • Culture tubes
  • Electrophoresis chamber and power supply
  • Eppendorf centrifuge 5425 R
  • Erlenmeyer Flask
  • Falcon Tubes (15 mL, 50 mL)
  • Gel documentation system (Bio-Rad Gel Doc XR+)
  • Gloves (nitrile)
  • Heidolph Rotamax 120
  • Incubator (30 °C and 37 °C)
  • Microcentrifuge tubes (1.5 mL, 2 mL)
  • Microscope slides
  • Microwave
  • Mini-PROTEAN Tetra Cell system (Bio-Rad)
  • NanoPhotometer® NP80
  • Nikon ECLIPSE Ti2 microscope
  • Parafilm
  • Petri dishes
  • Pipettes and tips (sterile)
  • PCR tubes (0.2 mL)
  • Shaker
  • Silica beads (0.1 mm, 500 µL)
  • Spectrophotometer (for OD₆₀₀ measurement)
  • TECAN Spark® microplate reader
  • Thermoblock
  • Thermocycler (Bio-Rad T100)
  • Thermo Scientific Multifuge X Pro series centrifuge
  • UV transilluminator
  • Vortex mixer
  • VWR Perfect Blue Power Supply Universal
  • Water bath (42 °C)
  • Well plates (384-well and general)

References

Preibisch, S., Rueden, C., Saalfeld, S., Schmid, B., Tinevez, J.-Y., White, D. J., Hartenstein, V., Eliceiri, K., Tomancak, P., & Cardona, A. (2012a). Fiji: An open-source platform for biological-image analysis. Nature Methods, 9(7), 676–682. https://doi.org/10.1038/nmeth.2019