PCR (Polymerase Chain Reaction) amplifies template DNA through repeated cycles of denaturation, annealing, and extension using a thermostable DNA polymerase. This protocol uses Q5 High-Fidelity DNA Polymerase (New England Biolabs) for accurate amplification of target sequences.

Materials

Step by Step Instructions

1. Set up the PCR reaction in a PCR tube by adding all components from the materials table to reach a total volume of 25 µL
2. Mix gently by pipetting or flicking the tube, then briefly spin down to collect all liquid at the bottom
3. Place the PCR tube in a thermocycler and run the following program: Initial denaturation at 98°C for 30 s, followed by 25-40 cycles of denaturation (98°C, 10 s), annealing (60-72°C, 10-30 s), and extension (72°C, 30 s/kb), then final extension at 72°C for 5 min, and hold at 4°C
4. Store the PCR product at 4°C until further use

Notes

Annealing temperature should be optimized based on primer Tm values (typically 60-72°C) and can be calculated with the NEB Calculator.

Extension time depends on amplicon length: use 30 seconds per kilobase.

Number of cycles (25-40) should be optimized to minimize off-target amplification while ensuring sufficient product yield. Typically 30 cycles are sufficient.

If regular Q5 PCR is not sufficient to achieve desired DNA amounts (e.g. because of poor Primer design) SuperFi DNA Polymerase can be used. Platinum™ SuperFi™ DNA Polymerase for high-fidelity amplification of DNA templates minimizes non-specific amplification, making it ideal for cloning and mutagenesis applications.

Materials

Step by Step Instructions

1. Prepare the reaction mix by combining all components in a PCR tube to achieve a final volume of 25 µL, adding template DNA and primers last
2. Mix the reaction gently by pipetting or flicking, then use a QuickSpin to briefly collect contents at the tube bottom
3. Transfer tubes to a thermocycler and run the following program: initial denaturation at 98°C for 30 s, followed by 25-35 cycles of denaturation (98°C, 10 s), annealing (60°C regardless, no need for different annealing temperatures, 10 s), and extension (72°C, 30 s/kb), then final extension at 72°C for 5 min, and hold at 4°C

This protocol anneals complementary single-stranded oligonucleotides to form double-stranded DNA fragments. Starting with 100 µM oligo stocks, the procedure typically yields 10 µM annealed products suitable for cloning applications such as Golden Gate assembly.

Materials

Step by Step Instructions

1. Preheat a heating block to 95°C while preparing the reaction mixture
2. Combine all components in a 1.5 mL microcentrifuge tube for a total volume of 50 µL
3. Place the tube in the heating block and incubate at 95°C for 5 minutes to fully denature the oligonucleotides
4. Turn off the heating block and allow the tube to cool slowly inside the block for at least 1 hour, enabling gradual annealing of complementary strands
5. Store the annealed oligonucleotides at -20°C until needed for downstream applications

Notes

Volumes can be scaled proportionally if different final concentrations or volumes are required. As an alternative cooling method, use a thermocycler programmed to cool at 0.5°C per second from 95°C to room temperature for more controlled annealing. The rCutSmart buffer provides optimal salt conditions for efficient annealing of complementary oligonucleotides. Ensure oligonucleotides are designed with perfect complementarity in the regions intended to anneal.

Golden Gate Assembly allows efficient and precise joining of multiple DNA fragments in a single tube (single pot reaction) using type IIS restriction enzymes such as BsaI or Esp3I. These enzymes cut outside their recognition sites, enabling seamless, scar-free assembly of multiple-fragments to one construct.

Materials

Step by Step Instructions

1. Combine all components in a PCR tube, maintaining a 1:3 molar ratio of vector to insert DNA, and adjust the final volume to 25 µL with nuclease-free water
2. Transfer the PCR tube to a thermocycler and execute the following program: 10-30 cycles alternating between restriction enzyme activation (37°C, 5 min) and ligase activation (16°C, 5 min), followed by enzyme inactivation at 65°C for 5 min, then hold at 4°C
3. Store the assembled product at 4°C until transformation or further processing

Notes

The 1:3 vector to insert molar ratio can be adjusted if assembly efficiency is low. Cycle number (10-30) can be optimized depending on fragment complexity and insert number. Ensure that all DNA fragments have compatible overhangs designed for Golden Gate assembly. The type IIS restriction enzyme used (e.g., BsaI, BbsI) must match the recognition sites in your constructs. Add 3-5 basepairs at the ends of linear constructs containing terminal type IIS recognition sides to enhance enzyme activity.

Gibson assembly enables scarless joining of multiple DNA fragments with overlapping ends without requiring restriction sites. This one-step isothermal reaction uses exonuclease, polymerase, and ligase enzymes to assemble fragments in a sequence-independent manner, making it highly versatile for complex cloning projects.

Materials

Step by Step Instructions

1. Design DNA fragments with 15-30 bp overlapping regions. DNA fragments should be >120 bp for this protocol
2. Amplify all DNA fragments by PCR, ensuring primers incorporate the designed overlapping sequences at fragment termini
3. Combine the appropriate amount of DNA fragments with Gibson Assembly Master Mix and nuclease-free water in a PCR tube to reach a final volume of 20 µL
4. Incubate the assembly reaction at 50°C for 15 minutes (2-3 fragments) or 60 minutes (4-6 fragments)
5. Transform 2-5 µL of the assembly reaction into competent cells or store at -20°C until transformation

Notes

For assembling 4-6 DNA fragments 0.2 - 0.5 pmols of total DNA should be used. The recommended Vector:Insert ratio is 1:2 for 2-3 fragments or 1:1 for 4-6 DNA fragments.

KLD (Kinase, Ligase, DpnI) ligation efficiently circularizes blunt-end DNA fragments using a streamlined enzyme mix containing kinase for phosphorylation, ligase for joining, and DpnI restriction endonuclease for digesting template DNA. This rapid protocol is particularly useful for cloning PCR products or assembled fragments.

Materials

Step by Step Instructions

1. Combine all components in a microcentrifuge tube to achieve a total reaction volume of 10 µL
2. Mix thoroughly by gently pipetting the reaction up and down several times
3. Incubate the reaction at room temperature (25°C) for a minimum of 5 minutes to allow complete ligation
4. Use 5 µL of the ligation mixture directly for transformation into 50 µL of competent cells

Notes

The DpnI component digests methylated template DNA, reducing background from PCR templates in downstream transformations. Extended incubation beyond 5 minutes does not significantly improve ligation efficiency for most applications. The reaction can be performed at room temperature without specialized equipment, making it convenient for routine cloning. Store unused ligation mixture at -20°C if transformation will be delayed, though fresh ligations typically yield better results.

Agarose gel electrophoresis separates DNA fragments based on size, using electricity. Smaller fragments migrate faster through the agarose gel matrix towards the Kathode than larger fragments. This technique is essential for visualizing PCR products, verifying digestions, and purifying DNA fragments of specific sizes.

Materials

Step by Step Instructions

1. Pour an agarose gel with the appropriate agarose concentration typically ranging from 1-3% (w/v). Add 3 µL SYBR-Safe directly to the still liquid agarose gel and stir with a pipette tip until no more colored traces are visible
2. Allow the gel to solidify for 15-30 minutes at room temperature until firm
3. Add 6x Loading Dye to your PCR Product volume and mix thoroughly by pipetting up and down
4. Place the solidified gel in an electrophoresis chamber, cover with TAE buffer, load DNA samples and ladder into separate wells
5. Run the gels at 120-135V for 20-30 min for a standard gel
6. Visualize DNA bands using UV or blue light illumination, and document or excise bands as needed

Notes

Higher agarose concentrations provide better resolution for smaller fragments but increase run time. Recommended agarose concentrations for different fragment lengths are given in Table 2. Gels are prepared by mixing the appropriate mass of Agarose powder with TAE buffer in an Erlenmeyer flask, heating it in a microwave until completely dissolved, swirling occasionally and monitoring carefully to prevent boiling over. The lid of the flask must be unscrewed to prevent pressure build-up and bursting of the flask. Once heated the dissolved agarose can be kept liquid in a water bath at 60°C until gel casting. This temperature allows immediate addition of staining agents. SYBR Safe is preferred over ethidium bromide for safety reasons and can be visualized with blue light transilluminators. As a rule of thumb 10 volts per centimeter of gel are recommended. Voltage can be adjusted for faster or slower runs, but higher voltages may cause band distortion or gel heating. The maximum voltage that can be applied without causing issues remains to be determined.

Gel extraction recovers specific DNA fragments from agarose gels following electrophoresis. This technique is essential for isolating correctly sized PCR products, restriction fragments, or assembly products before downstream cloning steps.

Materials

Step by Step Instructions

1. Visualize DNA bands on a transilluminator and identify the fragment of interest based on size comparison with the DNA ladder
2. Using a clean scalpel, carefully excise the gel slice containing the desired DNA band while minimizing excess agarose
3. Weigh the gel slice and follow the manufacturer's protocol provided with the QIAquick Gel Extraction Kit to dissolve the agarose and purify the DNA
4. Elute purified DNA in an appropriate volume of nuclease-free water and measure concentration using a NanoDrop spectrophotometer
5. Use the extracted DNA for subsequent cloning assembly, or other applications

Notes

Minimize UV exposure time during band excision to prevent DNA damage—use a blue light transilluminator when possible. Remove excess agarose around the band to improve extraction efficiency and reduce contaminants. Preheating water for elution to 50°C and performing a second elution can increase DNA yield.

Chemically competent cells acquire the ability to uptake exogenous DNA through treatment with calcium chloride, which destabilizes the bacterial cell membrane. These cells can then be transformed via heat shock, making them essential for molecular cloning workflows.

Materials

*10X transformation buffer: 100 mM HEPES pH 6.7, 150 mM CaCl₂, 550 mM MnCl₂, 2.5 M KCl

Step by Step Instructions

1. Streak DH5α from glycerol stock onto an LB agar plate and incubate overnight at 37°C to obtain isolated colonies
2. Inoculate a single colony into 5 mL LB medium and grow overnight at 37°C with shaking
3. Supplement 1 L of LB medium with 10 mL each of 1 M MgSO₄ and 1 M MgCl₂, then inoculate to an initial OD₆₀₀ of 0.006 using the overnight culture
4. Grow the main culture at 20°C with shaking until OD₆₀₀ reaches approximately 0.1-0.2, then increase temperature to 25°C and continue growth until OD₆₀₀ reaches 0.45
5. Aliquot into 50 mL tubes, and chill in an ice-water bath for 10 minutes
6. Pellet cells by centrifugation at 3000 × g for 15 minutes at 4°C, discard supernatant, and resuspend each pellet in 3 mL ice-cold 1X transformation buffer
7. Pool all suspensions, incubate on ice for 10 minutes, then pellet again at 3000 × g for 15 minutes at 4°C
8. Resuspend the final pellet in 25 mL ice-cold 1X transformation buffer, add 2 mL DMSO, mix gently by inversion, and aliquot into pre-chilled microcentrifuge tubes
9. Store microcentrifuge tubes immediately at -80°C

Notes

This protocol typically requires 3-4 days from initial streaking to frozen competent cells. All manipulations after culture harvest must be performed at 4°C to preserve cell viability. Work on ice whenever possible.

Heat shock transformation introduces plasmid DNA into chemically competent E. coli cells by briefly disrupting the cell membrane at elevated temperature. This rapid method is suitable for routine cloning and enables selection of successfully transformed cells through antibiotic resistance markers.

Materials

Step by Step Instructions

1. Thaw one aliquot of competent cells on ice for 15 minutes, handling gently to avoid premature warming.
2. Add 1-2 µL of plasmid DNA directly to the cells and incubate on ice for 30 minutes to allow DNA binding.
3. Transfer cells to a 42°C heat block for exactly 30 seconds to induce heat shock.
4. Immediately return cells to ice and incubate for 10 minutes to allow membrane recovery.
5. Transfer the full volume onto LB agar plates containing the appropriate selection antibiotic, spread using an inoculation loop and incubate overnight at 37°C.

Notes

Optionally SOC medium can be added to the cells after heat shock and incubated for 45 min before plating. With the exception of the 30 second heat shock step, this protocol proved very robust, reliably transforming bacteria even when cells were kept on ice for longer or shorter periods before or after heatshock.

MiniPrep purification extracts plasmid DNA from bacterial cultures using alkaline lysis followed by column-based purification. This method provides high-quality plasmid DNA suitable for sequencing, downstream cloning applications and transfection into mammalian cells.

Materials

Step by Step Instructions

1. Pick a single colony from a transformation plate and inoculate into 5 mL LB medium containing the appropriate selection antibiotic
2. Incubate the liquid culture at 37°C with shaking for 12-16 hours until turbid
3. Pellet the bacterial cells by centrifugation at maximum speed for 1-10 minutes, then discard the supernatant
4. Purify plasmid DNA from the cell pellet following the manufacturer's protocol provided with the QIAprep Spin Miniprep Kit
5. Elute plasmid DNA in 30-50 µL of nuclease-free water, then measure DNA concentration using a NanoDrop spectrophotometer at 260 nm
6. Analyse purified plasmid DNA with sequencing or restriction digest it to interpret transformation success by gel electrophoresis

Notes

Picking two colonies allows a backup option—the second pellet can be stored at 4°C and processed only if the first clone proves incorrect after sequencing. Preheating water to 50°C and performing a second elution can increase DNA yield. For processing multiple samples simultaneously, a vacuum-manifold instead of a centrifuge can substantially accelerate workflow with the QIAprep Spin columns. Avoid over-incubating cultures beyond 16 hours, as this may reduce plasmid quality due to chromosomal DNA contamination. For rapid DNA preparation (6 hours instead of overnight) or higher yields, inoculate two separate 5 mL cultures using the same colony pick, then combine the resuspended pellets after centrifugation to increase yield from shortened growth time.

This protocol describes how to produce a protein in bacteria and purify it via HisTag - Ni-NTA column.

Buffers and Reagents

Protein Expression

Day 0 - Overnight culture

1. Prepare 20 mL overnight culture (ONC) of transformed bacteria in LB and selection antibiotics

Day 1 - Recombinant protein expression

1. Inoculate 500-2000 mL 2xYT media 1:100 with ONC and selection antibiotics
2. Incubate cells at 37 °C, shaking at 180 rpm until OD₆₀₀ 0.4-0.6 is reached
3. Add inducer for expression (for example 0.1-0.2 mM IPTG)
4. Incubate for 24h at 32 °C, shaking at 180 rpm
5. Pellet cells at 3,750 x g for 45 min at 4 °C and transfer pellet into a 50 mL tube (pellet can be stored at -80 °C)

Cell Lysis

1. Dissolve pellet in 2 mL/g (pellet weight) lysis buffer (NaP10 with lysozyme and protease inhibitor)
2. Sonicate samples on ice for 10 min at 40% amplitude with 20 s on and 30 s off
3. Add DNAse I (1:1000) and incubate for 15 min on ice
4. Centrifuge to clear lysate at 20,000 x g for 30 min at 4 °C (Take 20 µL aliquot)
5. Sterile filter supernatant with 0.22 µm filter (Take 20 µL aliquot)

Purification of His-tagged Protein

Column Preparation

1. Add 1 mL of Ni-NTA resin slurry (500 µL effective Ni-NTA resin) to a gravity-flow column and wait until storage buffer is drained (can also be done at slow speed in a centrifuge)
2. Equilibrate column twice with 2 mL of NaP10

Purification

1. Apply filtrate onto the column and collect flow-through in a 50 mL tube (Take 20 µL aliquot)
2. Apply the flow-through onto the column again twice to increase yield
3. Wash column with 10 mL NaP10 (Take 20 µL aliquot)
4. Wash column with 10 mL NaP20 (Take 20 µL aliquot)
5. Repeat step 4
6. Apply 4 mL NaP250 (Elution buffer)
7. Seal column with cap and incubate at room temperature for 30 min, light shaking
8. Open column and collect purified protein

This protocol describes how to keep Human Embryonic Kidney 293T (HEK293T) cells in culture for use in experiments. It is optimized for culture in T75 flasks, volumes vary for other formats. Cell culture work requires a sterile environment in a biosafety cabinet (BSC) under laminar flow. All liquid handling above 1 mL should be done using serological pipettes and not by pouring.

1. Warm reagents to 37 °C in a water bath and prepare complete DMEM supplemented with 10% fetal calf serum (FCS), Penicillin/Streptomycin (P/S) (final concentration: 100 U/ml penicillin, 100 µg/mL streptomycin) and L-glutamine (final concentration: 2 mM):

2. Take flask (T75) out of the incubator (37°C, 5% CO₂) and inspect cells under a light microscope to assess confluency, then move into BSC. It is recommended to split at a confluence of 70-80%.

3. Aspirate old medium from the flask and aspirate old medium.

4. Wash with 4 mL Dulbecco’s Phosphate Buffered Saline (DPBS), tilt flask and aspirate DPBS.

5. Add 1 mL trypsin-EDTA and let sit until cells visibly detach (takes around 3-4 minutes)

5. Add 9 mL DMEM to neutralize trypsin and thoroughly resuspend cells.

6. Count cells or decide dilution based on confluency. At 80% confluency, a 1:5 split (2 mL of old cells) is ready to split again after 48h, 1:10 after 72h, 1:20 after 96h)

7. Dilute with DMEM to desired cell concentration.

8. Transfer 10 mL of diluted cells into a new T75 flask.

Notes: For splitting at 70-80% confluence, these rules of thumb work for splitting ratios: A 1:5 split (2 mL of cells + 8 mL DMEM) is ready to split again after 48h, 1:10 (1 mL cells + 9 mL DMEM) after 72h, 1:20 (0.5 mL + 9.5 mL DMEM) after 96h. Ratios with less cells than 1:20 are not recommended.

This protocol describes how to seed Human Embryonic Kidney 293T (HEK293T) cells for experiments, mostly involving transfection. Cell culture work requires a sterile environment in a biosafety cabinet (BSC) under laminar flow. All liquid handling above 1 mL should be done using serological pipettes and not by pouring.

1. Split cells until cell collection steps as described in Cell Maintenance protocol.

2. Seed according to the used plate format, plate should be declared as tissue culture treated.

3. Gently shuffle plate to disperse across wells and place cells into the incubator.

This protocol describes how to transfect Human Embryonic Kidney 293T (HEK293T) cells using Lipofectamine 2000 (Thermo Fisher Scientific, Waltham) as a cationic transfection reagent. The goal of transfection is to introduce exogenous DNA into eukaryotic cells.

1. Seed cells in an appropriate well plate and incubate at 37°C and 5% CO₂ for 24h.
2. Dilute DNA to desired concentrations in OptiMEM (serum-reduced medium) (Thermo Fisher Scientific, Waltham).
3. Dilute Lipofectamine 2000 in OptiMEM. Incubate for 5 min at room temperature.
Table 1: Lipofectamine 2000 and DNA amounts per well for different plate formats. The table shows the recommended amounts of DNA and Lipofectamine 2000 transfection reagent for different well plate formats used in transfection experiments.

Component	96-well	24-well	12-well
DNA	100-200 ng	200-400 ng	300-600 ng
Lipofectamine 2000	0.5 µL	1.0 µL	2.0 µL

4. Add diluted Lipofectamine 2000 to DNA solutions, briefly vortex and spin down. Incubate at room temperature for 20 min.
5. Add DNA-Lipo-Mix to cells and incubate them until measurement or induction.

NanoLuc Luciferase (NanoLuc) is a 19 kDa luciferase, derived from the deep sea shrimp Oplophorus gracilirostris that produces bioluminescence in the presence of its substrate furimazine. It’s commonly used as a reporter in mammalian cells as it has a wide dynamic range and high sensitivity. This protocol describes how to measure its activity in a 96 well plate format using reduced amounts of reagent. We tested different amounts of reagent and the difference in results was negligible, which is why we settled for these amounts. This protocol explains both how to measure secreted NanoLuc in supernatant cell culture medium, and intracellularly expressed NanoLuc. For intracellular expression, NanoLuc is commonly normalized by Firefly luciferase (FLuc), which is then co-transfected in a plasmid with a weak constitutive promoter such as the Herpes simplex virus-thymidine kinase (HSV-TK) promoter.

Materials

96 well plate with transfected cells

Black or white non-transparent 96 well plate

NanoGlo Kit or NanoGlo Dual Luciferase Reporter (DLR) Kit (both Promega)

Passive Lysis buffer (Promega)

Phosphate buffered saline (PBS)

Amounts per Well

NanoGlo substrate needs to be prepared freshly before every use. For this the substrate needs to be diluted in the provided buffer. In the normal NanoGlo kit, the substrate is of 50X concentration, for the DLR kit it is 100X. The OneGloEX reagent for measuring FLuc can be prepared all at once and stored at -20 °C. Per well, 10 µL of reagent can be mixed with 25 µL PBS, yielding sufficient volume to cover the bottom of the well. Passive Lysis buffer should be diluted to 1X with WDC water.

Measuring secreted NanoLuc

Prepare diluted NanoGlo reagent as described above Transfer 10 µL of supernatant into opaque plate Add 35 µL of NanoGlo reagent/PBS Wait 10 min Measure Luminescence using a plate reader

Measuring intracellular NanoLuc and FLuc

1. Prepare diluted NanoGlo DLR, OneGloEX and 1X Passive Lysis buffer as described
2. Discard medium from cell culture plate
3. Add 30 µL 1X Passive Lysis buffer to each well
4. Shake plate at 500 rpm for 30 min at room temperature
5. Transfer 10 µL of lysate to opaque plate and add 35 µL OneGloEX reagent
6. Wait 10 min
7. Measure luminescence using a plate reader
8. Add 35 µL of NanoGlo DLR reagent to inhibit FLuc and activate NanoLuc activity
9. Wait 10 min
10. Measure NanoLuc luminescence using plate reader

This protocol describes how to perform flow cytometry for transfected Human Embryonic Kidney 293T (HEK293T) with mCherry reporter and GFP transfection normalization or SplitFast (RspA(N), RspA(C)) reporter with mCherry transfection normalization. In order to measure sufficient cells and accurately depict the cell population, 12 well plates are used. For conditions induced with a ligand, according ligands were included in each step including the final DPBS suspension.

1. Remove cells from the incubator 48h after transfection and wash each well twice with 1 mL DPBS without detaching the cells.
2. Detach cells by incubating them with 5 mM EDTA in DPBS at 37°C for 10 minutes and transfer suspension into 1.5 mL microcentrifuge tubes. From this point on handle samples on ice.
3. Centrifuge suspensions (3 min., 600 g) and discard supernatants. Resuspend the pellets in 500 µL DPBS and centrifuge (3 min., 600 g).
4. Discard the supernatant and resuspend the pellets in 200-400 µL DPBS depending on pellet size.
5. Strain cell suspensions through 40 µm cell strainers into fresh microcentrifuge tubes.
6. For SplitFast as reporter, add 5 µM HMBR to each sample.
7. Proceed with the measurement (Excitation lasers at 488 nm were used for SPlitFast and GFP, with fluorescence measurement at 525 nm. Excitation lasers at 638 nm were used for mCherry, with fluorescence measurement at 610nm.)
8. Subsequent analysis includes size gating, single cell gating and transfection gating (Fig. 1).

The CF2H assay provides a rapid method for validating computationally designed protein binders using a cell-free expression system. Successful binding events reconstitute a functional CI transcription factor dimer, activating sfGFP expression from a pRM promoter. This enables screening of BindCraft-designed binders within 24 hours without requiring protein purification or bacterial transformation.

Materials

Cell-free bacterial extract and cell-free buffer (prepared according to Capin et al., 2025, or commercial systems such as Promega E. coli S30 System for linear DNA)
pRM-sfGFP reporter plasmid (circular DNA)
Linear DNA fragments encoding CI-DBD fused to test binders
Biotinylated, Fc-tagged target protein (e.g., GDF-15)
Streptavidin
Low-binding 384-well plate with black walls and transparent bottom
Nuclease-free water

Preparation of Stock Solutions

Linear DNA fragments: Resuspend to 10 ng/μL in nuclease-free water to create a 20X stock (final concentration: 5 nM).
Note: To reduce synthesis costs, binder sequences can be ordered without CI-DBD and cloned downstream of a reusable CI-DBD construct, followed by PCR amplification using high-fidelity polymerase.

Reporter plasmid: Prepare as 20X stock at 100 nM (final concentration: 5 nM).

Biotinylated target protein: Resuspend to 1.25 μM in ultrapure water with gentle mixing. Do not vortex.
Note: Follow manufacturer guidelines for minimum aliquot volumes and consider adding 0.1% (w/v) BSA for protein stability.

Streptavidin: Prepare working solution at 1.25 μM in ultrapure water.

Protein-streptavidin complex: Mix streptavidin and biotinylated protein in 1:1 molar ratio. Incubate at room temperature for 1 hour.

CF2H Reaction Assembly

Work on ice throughout assembly. Assemble reactions in a total volume of 22 μL according to the following composition:

Bacterial extract: 7.3 μL (1/3 of total volume)
Cell-free buffer: 9.2 μL (5/12 of total volume)
Reporter plasmid (20X stock): 1.1 μL (final concentration: 5 nM)
Linear binder DNA (20X stock): 1.1 μL (final concentration: 5 nM)
Streptavidin-protein complex: 2.6 μL (final concentration: 150 nM)
Nuclease-free water: 0.7 μL

Assembly Steps

1. Combine bacterial extract and cell-free buffer in a PCR tube on ice
2. Add reporter plasmid and linear binder DNA
3. Add streptavidin-protein complex
4. Add nuclease-free water to reach final volume of 22 μL
5. Mix gently by pipetting

Incubation and Measurement

1. Incubate reactions at 37°C for 8 hours (Note: Fluorescence is detectable after 2 hours but reaches maximum intensity at 6-8 hours)
2. Dilute each reaction 1:3 with nuclease-free water and mix thoroughly
3. Transfer three 20 μL aliquots from each diluted reaction to separate wells of a 384-well plate (technical triplicates)
4. Measure sfGFP fluorescence using a plate reader with excitation at 485 nm and emission at 530 nm

Controls

Implement the following controls for data quality:

Blank: Cell-free extract only (no plasmid or DNA). Subtract mean fluorescence from all wells to correct for autofluorescence.

Negative control #1: pRM-sfGFP plasmid without binder DNA to establish leakage expression.

Negative control #2: Each binder without streptavidin-protein complex to detect homodimerization or aggregation artifacts.

Positive control: ALFA epitope tag and anti-ALFA nanobody (NbALFA), each fused to CI-DBD, which bind with low picomolar affinity.

Standard curves: Dilution series of purified GFP or fluorescein measured on the same plate for quantification.

Notes

BindCraft-designed binders with i_pTM scores above 0.8 should be prioritized, though higher scores do not necessarily correlate with superior binding. The assay provides qualitative validation and relative comparisons rather than quantitative binding affinities. This method is particularly useful for pre-screening candidates before biophysical characterization methods such as SPR.

This protocol enables visualization of fluorescently labeled proteins in mammalian cells using fluorescence microscopy. Cells are fixed with paraformaldehyde to preserve cellular structures, stained with DAPI to visualize nuclei, and mounted on microscope slides for imaging. This technique is essential for assessing protein localization, expression levels, and cellular distribution following transfection.

Materials

Step by Step Instructions

1. Seed HEK293T cells or other mammalian cells onto coverslips in a 24-well plate, transfect according to your standard protocol, and incubate for 36 hours.
2. Remove culture medium by vacuum aspiration under a fume hood and immediately add 250 µL of 4% PFA solution per well, then incubate at room temperature for 15-20 minutes in the dark to fix cells.
3. Perform three washing steps with 500 µL PBS per well, incubating for 5 minutes at room temperature with gentle rotation for each wash, disposing of all PFA waste in a designated waste bottle.
4. Add 190 µL of DAPI solution (diluted 1:1000 in PBS) to each well and incubate at room temperature for 5 minutes in the dark to stain nuclei.
5. Wash cells 2-3 times with PBS as before, then carefully remove each coverslip from the well using fine forceps.
6. Gently absorb excess liquid by touching the edge of the coverslip to a Kimwipe, then invert the coverslip and place it cell-side down onto 10-20 µL of Mowiol mounting medium on a labeled microscope slide.
7. Seal the edges of the coverslip with nail polish to prevent drying and store slides upright in the dark until the mounting medium solidifies, then transfer to a slide box.
8. Image samples using a fluorescence microscope with appropriate excitation and emission filters for your fluorophore and DAPI according to the manufacturer's instructions.

Notes

All steps involving PFA must be performed under a fume hood due to toxicity—dispose of all PFA-containing solutions in designated waste containers. Also wear appropriate personal protection equipment. Keep samples in darkness after fixation to prevent photobleaching of fluorescent signals When mounting coverslips, ensure cells face the slide (coverslip must be inverted) for proper imaging distance from the objective. Label slides immediately with date, construct information, and initials for proper sample tracking. Mounted slides can be stored for several weeks at 4°C in the dark, though signal intensity may decrease over time. Colored nail polish is advantageous for sealing as it provides better visualization of where polish has already been applied around the coverslip edges.

⚠️ SAFETY WARNING: Cobalt chloride is classified as a CMR (Carcinogenic, Mutagenic, Reprotoxic) substance. This protocol requires enhanced safety measures including mandatory double gloving, face mask protection, and fume hood work.

Safety Precautions

MANDATORY SAFETY EQUIPMENT:

Double nitrile gloves (change outer gloves after stock preparation)
Face mask (N95 or equivalent respiratory protection)
Safety goggles
Lab coat
Work exclusively under fume hood

WASTE DISPOSAL:

Segregate all cobalt chloride-contaminated waste
Dispose of tips, tubes, and gloves in designated CMR waste containers
Neutralize liquid waste according to institutional guidelines

Stock Solution Preparation (3 mM)

This protocol prepares a 3 mM cobalt chloride stock solution for hypoxia induction experiments.

Materials

Cobalt chloride hexahydrate (CoCl₂·6H₂O, MW: 237.93 g/mol)
Sterile nuclease-free water or cell culture-grade water
15 mL conical tube
Analytical balance
Vortex mixer
0.22 µm sterile filter

Procedure

1. Put on double gloves and face mask. Work under fume hood for all steps
2. Calculate the required mass of cobalt chloride hexahydrate:
   • For 3 mL of 3 mM solution: 2.14 mg CoCl₂·6H₂O
   • For 5 mL of 3 mM solution: 3.57 mg CoCl₂·6H₂O
3. Weigh 1-2 mg of cobalt chloride hexahydrate into a 15 mL conical tube using an analytical balance
4. Add sterile water to achieve the desired final volume for 3 mM concentration:
   • If 1 mg weighed: add 1.4 mL water
   • If 2 mg weighed: add 2.8 mL water
5. Vortex vigorously until completely dissolved (solution should be clear pink/red)
6. Change outer gloves after stock preparation
7. Filter sterilize through a 0.22 µm filter into a sterile tube
8. Label clearly with "3 mM CoCl₂ - CMR HAZARD" and date
9. Store at 4°C protected from light for up to 2 weeks

Hypoxia Induction in 96-Well Plate

This protocol induces hypoxia-like conditions in HEK293 cells using 100 µM final concentration of cobalt chloride.

Materials

3 mM cobalt chloride stock solution
HEK293 cells seeded in 96-well plate
Cell culture medium
Multichannel pipette
Sterile pipette tips

Procedure

1. Ensure double gloving and face mask are in place. Work under fume hood
2. Calculate treatment volume:
   • For 100 µM final concentration in 150 µL well volume: add 5 µL of 3 mM stock
   • For 200 µL well volume: add 6.7 µL of 3 mM stock
3. Remove old medium from cells or prepare for direct addition to existing medium
Table 1: Cobalt chloride treatment setup for 96-well plate.

Component	Volume per well	Final concentration
3 mM CoCl₂ stock	5 µL	100 µM
Cell culture medium	145 µL	-
Total volume	150 µL	-

4. Add 5 µL of 3 mM cobalt chloride stock to each designated well using a pipette
5. Add 145 µL of fresh cell culture medium or gently mix if adding to existing medium
6. Change outer gloves after treatment application
7. Incubate cells at 37°C, 5% CO₂ for desired time period (typically 4-24 hours for hypoxia-inducible factor activation)
8. Dispose of all contaminated materials in CMR waste containers

Controls and Considerations

Negative control: Wells treated with equivalent volume of sterile water
Positive control: Cells under actual hypoxic conditions (1% O₂) if available
Time points: Common treatment durations are 4h, 8h, 16h, and 24h
Viability: Monitor cell viability as cobalt chloride can be cytotoxic at high concentrations or long exposures

Waste Management

All pipette tips, tubes, and gloves must go into designated CMR waste containers
Liquid waste should be collected separately and disposed according to institutional CMR waste protocols
Decontaminate work surfaces with appropriate cleaning agents
Remove and dispose of outer gloves before leaving fume hood area