Team:Rochester/Experiments

Loading...

Project

Description Results Proof of Concept Modeling Hardware Engineering Success Contribution Software Measurement

Lab

Notebook Experiments Safety Parts

Practices

Education Inclusivity Human Practices Entrepreneurship Implementation Communication

Collaboration

Teams Partnership Sponsors

Team

Team Attributions Public Relations

Menu

Top

Stop Animations

Menu

Top

Stop Animations

Experiments

Introduction

The goal of our project is to develop a way to continuously monitor the concentration of biomarkers (small molecules that are associated with a disease or condition), specifically to help with the diagnoses of sepsis. We designed a biosensor that uses aptamers (single stranded pieces of DNA that can bind to proteins and similar molecules) to detect the presence of sepsis biomarkers. These aptamers are attached to reduced graphene oxide (rGO), and the conformational change that results from aptamer binding alters the electrical properties of the rGO. By measuring these property changes, we can track the concentration of biomarkers with our sensor. You can read more about the hardware component here Aptasensor hardware. To maximize our efficiency, our wetlab team split into two subteams: one to design, produce, and test our aptamers, and one to engineer Shewanella oneidensis MR-1 to synthesize the rGO for our project. Click the buttons below to learn more about each part of our project.

Aptamers rGO

Aptamer Experiments

We ordered the aptamers DNA from IDT as small synthetic genes, so they came inside a vector with ampicillin resistance, pUCIDT - Amp Golden Gate. To make sure we will always have a supply of the template DNA for our asymmetric PCR experiments, the first step was to introduce the plasmid in Escherichia coli (E. coli). We also prepared our own chemically competent cells.

Protocol¹

Protocol

1. Streak strain on LB and grow overnight.
2. Inoculate 500 ml SOC (recipe below) to OD600 of no more than 0.04 in the evening.
3. Grow overnight at 18°C with vigorous shaking. Grow until OD600 is approximately 0.6 (0.5-0.6 is best).
4. Incubate cells for at least 10 minutes on ice. Centrifuge 10 minutes at 4°C, 2500 rpm.
5. Discard the supernatant and resuspend in 8ml TB (recipe below). Spin again.
6. Discard supernatant and resuspend in 4ml TB + 0.3ml DMSO.
7. Incubate on ice for 30 minutes and freeze 100µl aliquots in liquid nitrogen and store at -80°C.

SOC Recipe (makes 500mL)

Combine the following in a 1000 mL flask.

• Tryptone - 10g
• Yeast extract - 2g
• NaCl - 0.25g
• 2M KCl - 0.4ml
• Adjust to pH 7 with NaOH. Autoclave, then add:
• 1M MgCl2 - 5ml
• 1M glucose - 5ml. Add water to 500 mL

TB Recipe (Makes 100 mL)

• Make 0.5M PIPES and adjust to pH 6.7 with KOH. Filter sterilize.
• For 100ml of TB mix: 1.1 g MnCl2 4H2O, 0.2 g CaCl2 2H2O, 1.9g KCl, 2ml of 0.5M PIPES and water to 100ml.

Close

Transformation Protocol

Protocol

1. Mix DNA (1-5 µl) with 100µl cells.
2. Incubate on ice 30 minutes.
3. Heat shock at 42°C for 30 seconds and return to ice for 2 minutes.
4. Add 500µl LB and grow at 37°C for 1 hour at 200rpm.
5. Plate appropriate dilutions and leave overnight at 37°C.

Close

To prepare a frozen stock culture of the E. coli cells that contain our plasmid of interest, we first have to plate them on media with ampicillin (Amp) resistance. We prepared our own LB-Amp plates.

LB-Amp Plates Preparation

PROTOCOL:

1. Place magnetic bar in Erlenmaeyer flask for stirring.
2. Weigh all powders (see recipe).
3. Add appropriate amount of water, following the recipe.
4. Once dissolved, Cover with foil, place in proper liquid autoclave bin and autoclave.
5. Take out of the autoclave, let it cool down but DO NOT LET IT SOLIDIFY. Place it on hot plate, at 50 C and stirring.
6. When ready to add Amp, add 1 mL Amp (50 mg/mL) by quickly removing the foil and pipetting the 1 mL. (such that final Amp concentration is 50 µg/mL).
7. Let it stir briefly.
8. Once it's cooled such that you can comfortably hold the flask, pour on agar plates. Pour such that the plate is about 1/2 full or less.
9. Let the plates solidify and then place in the fridge, labelled with the proper color code.

Broth Recipe (1 Liter)

• DI water (700mL)
• Tryptone - 10 g
• NaCl - 10 g
• Yeast extract - 5 g
• Add water up to 1 L.

Close

After the plates were in the incubator at 37°C overnight, we are ready to prepare the frozen stock.

Frozen Stock

Protocol

Materials

• Plate with single colonies
• Liquid media (no agar) with right selection marker
• Shaking incubator at required temperature
• Next day: 50% glycerol - media (see recipe below)

Steps

1. Add 3 mL of sterile liquid media with the right selection marker to a sterile tube.
2. Pick one single colony of bacteria from the plate, using a tip of a 1000 uL pipette. Move the tip inside the media for about 30 sec, to ensure that the colony went all in solution.
3. Place the tube in a shaking incubator overnight.
4. Next day: take 1500 µL liquid culture and place in a freezer-friendly microcentrifuge tube (cryogenic screw-cap vial).
5. Spin down for 60 sec at the highest microcentrifuge speed.
6. Remove the supernatant and add 1000 µL liquid culture.
7. Spin the tube down again, and remove the supernatant.
8. Add 750 µL of the glycerol-media solution to the tube. Place in a freezer box, in the -80 C freezer.

Glycerol Recipe

1. Dilute 100% glycerol solution to 50% glycerol in a microcentrifuge tube (with water).
2. Add 5mL TSB or LB using Powerpette plus to a newly labeled 15mL tube.
3. Add 5mL 50% glycerol to the same tube using a syringe.
4. Vortex and mix the solution well.
5. Set up Corning™ Disposable Vacuum Filter with the neck connected to the tube that is connected to vacuum faucet.
6. Pour the mixed solution in a glass container, turn on vacuum faucet, start filtration.
7. After all liquid has come down, discard the top part of the filtration device and cap the glass container.
8. Store in the fridge

Close

Once we have the frozen culture, it was always streaked on a plate first, and then a liquid culture was grown from a single colony. The liquid culture was used to miniprep the plasmid DNA, using Monarch® Plasmid DNA Miniprep Kit Protocol (NEB #T1010).

Plasmid Miniprep

Protocol

Note: Buffers mentioned in the protocol are included in the kit.

1. Pellet 1.5 ml bacterial culture (or more, but not to exceed 15 OD units) by centrifugation for 30 seconds. Discard supernatant.
2. Resuspend pellet in 200 μl Plasmid Resuspension Buffer (B1). Vortex or pipet to ensure cells are completely resuspended. There should be no visible clumps.
3. Lyse cells by adding 200 μl Plasmid Lysis Buffer (B2). Invert tube immediately and gently for 5–6 times until color changes to dark pink and the solution is clear, uniform and viscous. Do not vortex! Incubate for one minute.

Note: Care should be taken not to handle the sample roughly and risk shearing chromosomal DNA, which will co-purify as a contaminant. Avoid incubating longer than one minute to prevent irreversible plasmid denaturation.

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

Note: Be careful not to shear chromosomal DNA by vortexing or vigorous shaking. Firmly inverting the tube promotes good mixing, important for full neutralization.

5. Clarify the lysate by spinning for 2–5 minutes at 16,000 x g. For culture volumes > 1 mL, a 5 minutes spin is recommended to ensure efficient RNA removal by RNase A.
6. Carefully transfer supernatant to the spin column and centrifuge for 1 minute. Discard flow-through.
7. Re-insert column in the collection tube and add 200 μl of Plasmid Wash Buffer 1. Plasmid Wash Buffer 1 removes RNA, protein and endotoxin. (Add a 5 minute incubation step before centrifugation if the DNA will be used in transfection.) Centrifuge for 1 minute. Discarding the flow-through is optional.

Note: Empty the tube whenever necessary to ensure the column tip and flow-though do not make contact.

8. Add 400 μl of Plasmid Wash Buffer 2 and centrifuge for 1 minute.
9. Transfer column to a clean 1.5 ml microfuge tube. Use care to ensure that the tip of the column has not come into contact with the flow-through. If there is any doubt, re-spin the column for 1 minute before inserting it into the clean microfuge tube.
10. Add ≥ 30 μl DNA Elution Buffer to the center of the matrix. Wait for 1 minute, then spin for 1 minute to elute DNA.

Note: Nuclease-free water (pH 7–8.5) can also be used to elute the DNA. Delivery of the Monarch DNA Elution Buffer should be made directly to the center of the column to ensure the matrix is completely covered for maximal efficiency of elution. Additionally, yield may slightly increase if a larger volume of DNA Elution Buffer is used, but the DNA will be less concentrated as a result of dilution. For larger plasmids (≥ 10 kb), heating the DNA Elution Buffer to 50°C prior to eluting and extending the incubation time after buffer addition to 5 minutes can improve yield.

Note: We used nuclease-free water for all elutions to ensure the buffer does not interfere with downstream applications.

Close

The miniprepped DNA was quantified using the Nanodrop, and then we prepared a working concentration of 10 ng/μl to be used in asymmetric PCR.

Asymmetric PCR

Materials

• Taq DNA polymerase - 500 units at 5U/μL
• PCR reaction buffer 1X - stock 1.25 mL 10X without MgCl₂
• dNTPs mix 200 uM each - stock 10 mM each.
• Template DNA, primers
• Forward primer: 100 μM (FWD)
• Reverse primer: 10 μM (REV)

Prepare master mix with all reagents, with Taq DNA polymerase being added last.

At first, these are the volumes (in uL) we used in order to run 8 PCR reactions of 20 μL each, for gradient PCR.

	10x buffer	MgCl2	dNTPs	Taq	DNA	FWD	REV	H2O	Total
IL-1β	18	5.4	3.6	4.5	18	4.5	2.25	123.75	180
IL6	18	5.4	3.6	4.5	18	4.5	2.25	123.75	180
TNF-α	18	5.4	3.6	4.5	18	4.5	2.25	123.75	180
Lactoferrin	18	5.4	3.6	4.5	18	4.5	2.25	123.75	180
CRP	18	5.4	3.6	4.5	18	4.5	2.25	123.75	180

After optimizing our conditions, these are the volumes we used:

	10x buffer	MgCl2	dNTPs	Taq	DNA	FWD	REV	H2O	Total
IL-1β	15	6	3	1.5	6	1.5	0.5	116	150*
IL6	44	0	4.4	2.2	4.4	2.2	1.47	161.33	220***
TNF-α	44	0	4.4	2.2	4.4	2.2	1.1	161.7	220***
Lactoferrin	12	4.8	2.4	1.2	4.8	1.2	0.6	93	120*
CRP	11	4.4	2.2	2.2	4.4	2.2	1.47	82.13	110**

* - for two PCR reactions of 50 μL

** - for four PCR reactions of 25 μL

*** - these “final” conditions were with Q5 polymerase, for which the buffer is 5X and already has MgCl₂

The PCR cycle followed was:

Initial Denaturation	95°C	1 min
Denaturation	94°C	45 sec (15-20x)
Annealing Temp	Specific Temperature	1 minute (15-20x)
Elongation	68°C	1 minute (15-20x)
Final Elongation	68°C	5 minutes

Close

We purified the PCR products using the PCR & DNA cleanup kit from NEB (NEB #T1030).

DNA Cleanup

Protocol

1. A starting sample volume of 50 μl is recommended. For smaller samples, nuclease-free water can be used to adjust the volume. Larger sample up to 100 μl work well too.
2. Add 100 μl DNA Cleanup Binding Buffer (ensure that isopropanol has been added, as indicated on the bottle label)* to the 50 μl sample.

*- if needed

3. Add 300 μl ethanol (≥ 95%). Mix well by pipetting up and down or flicking the tube. Do not vortex.
4. Insert column into collection tube, load sample onto column and close the cap. Spin for 1 minute, then discard flow-through.
5. Re-insert column into collection tube. Add 500 μl DNA Wash Buffer and spin for 1 minute. Discard flow-through.
6. Repeat Step 5 (Optional). This step is recommended for removal of enzymes that may interfere with downstream applications (e.g., Proteinase K).
7. Transfer column to a clean 1.5 ml microfuge tube. Use care to ensure that the tip of the column does not come into contact with the flow-through. If in doubt, re-spin for 1 minute to ensure traces of salt and ethanol are not carried over to the next step.
8. Add ≥ 6 μl of DNA Elution Buffer to the center of the matrix. Wait for 1 minute, then spin for 1 minute to elute the DNA. Typical elution volumes are 6–20 μl. Nuclease-free water (pH 7–8.5) can also be used to elute the DNA. Yield may slightly increase if a larger volume of DNA Elution Buffer is used, but the DNA will be less concentrated.

We used nuclease-free water to make sure the buffer doesn’t interfere with downstream purposes.

Close

We determined the concentration of our products using Quantifluor kit, which has a specific fluorescence dye for ssDNA and a different dye for dsDNA.

Quantification

Protocol

Quantifluor Protocol (Catalog #E3190 - ssDNA and #E2670 - dsDNA)

1. Prepare 1X TE buffer: Dilute the 20X TE Buffer 20-fold with nuclease-free water (not provided).
2. For example, add 1ml of 20X TE Buffer to 19ml of Nuclease-Free Water (Cat.# P1195), and mix.
3. Prepare Working Dye Solution: Prepare enough QuantiFluor® ssDNA Dye working solution to quantitate blank, standard and unknown samples. Protect the working solution from light by covering with foil or placing in the dark.
4. High-Concentration Standard Curve (for samples 10–400ng/μl): Dilute the QuantiFluor® ssDNA Dye 1:400 in 1X TE buffer to make the QuantiFluor® ssDNA Dye working solution. For example, add 10μl of QuantiFluor® ssDNA Dye to 3,990μl of 1X TE buffer, and mix thoroughly.
5. Low-Concentration Standard Curve (for samples 0.2–10ng/μl): Dilute the QuantiFluor® ssDNA Dye 1:2,000 in 1X TE buffer to make the QuantiFluor® ssDNA Dye working solution. For example, add 2μl of QuantiFluor® ssDNA Dye to 3,998μl of 1X TE buffer, and mix thoroughly.

Prepare Standard Curve

High-Concentration Standard Curve: The following recommended standards result in 6.25-400ng/well and are designed for optimal pipetting accuracy, transferring 10μl of standard to each well.

1. Prepare seven 1.5ml tubes labeled: 400, 200, 100, 50, 25, 12.5, 6.25 (for ssDNA) and 200, 50, 12.5, 3.1, 0.78, 0.2, 0.05 (for dsDNA)
2. b. Prepare ssDNA or dsDNA standards by serially diluting QuantiFluor® ssDNA Standard (100ng/μl) as shown in Table 1. Do not introduce air bubbles.


Table 2. Preparing Recommended dsDNA Standard Curve Samples.

3. Pipet 200μl of QuantiFluor® ssDNA Dye working solution into each well that is intended for an unknown, blank or standard sample.
4. Dispense 10μl of the high-concentration ssDNA standards prepared in Table 1 (labeled Standards A–G) to rows A–G of the 96-well plate (Figure 3, Panel A). Alternatively, dispense 10μl of the low-concentration ssDNA standards prepared in Table 2 (labeled Standards A–G) to rows A–G of the 96-well plate (Figure 3, Panel B). We recommend pipetting duplicates or triplicates of the standards.
5. For the blank, pipet 10μl of 1X TE Buffer into row H.
6. Add 1–20μl of unknown sample to the remaining wells.
7. Mix the plate thoroughly using a plate shaker or by pipetting the contents of each well up and down.
8. Incubate assays for 5 minutes at room temperature, protected from light.
9. Measure fluorescence (492nmEx/528nmEm - for ssDNA and 504nmEx/531nmEm - for dsDNA) using your plate reader. If using the GloMax® Discover System, select the preloaded protocol: “QuantiFluor ssDNA System.” or QuantiFluor dsDNA System.”
10. Calculate concentration by removing the blank fluorescence value from each measurement (all standard and unknown samples) and create a standard curve. Use the equation to determine the concentration. Example of obtained standard curves:

Close

After the concentration is known, we attached the optimized aptamers to reduced graphene oxide (rGO), which was then drop-casted on the electrode (Drop casting) and tested with different biomarker solutions using our low-cost potentiostat.

Attaching aptamers to rGO

Protocol

1. Sonicate rGO for 1 hour
2. Use 10 mL rGO (4g/L) and 2 mL aptamer solution.
• dilute aptamer solution in aptamer binding buffer*, such that the aptamer concentration is 10 nM in the 2 mL final solution.
3. Put on end over end rotator for 2 hours at r.t. If not working with it right away, place in fridge.
4. Filter via gravity filtration, washing with 20 mL aptamer binding buffer. Leave the rGO to dry on paper and then resuspend in 10 mL water. Shake very well until fully resuspended.
5. Drop cast on electrode.

*The aptamer binding buffer is different for each aptamer, according to the literature. (can add references from crp and il1b papers linked in results/design page)

CRP buffer: 5mM NaH2PO4/Na2HPO4 at pH = 7, 0.05 M NaCl and 100 mM MgCl₂

IL-1β buffer: 10 mM Tris, 50 mM KCl, 3 mM MgCl2, 0.1 mM EDTA and 1.0 mM DTT

Lactoferrin buffer: pH = 7.4: 20 mM HEPES, 2 mM MgCl2, 150 mM NaCl, 2 mM CaCl2, and 2 mM KCl

Close

We showed that the aptamer successfully attached to the rGO by placing the solution on glass slide, incubating with Quantifluor (QF) ssDNA dye and taking confocal microscope images.

Microscopy

Protocol

1. Plate 500 μL of each aptamer-rGO (or buffer-rGO as control) on glass slide and let it dry at r.t. Overnight.
2. Place one slide in a small box and add 500 uL of QF ssDNA dye solution directly on top of slide (only the black rGO surface), and let it incubate for 5 minutes at room temperature, in the dark.
3. After 5 minutes, wash with 1x TE buffer to wash away any unbound dye. For washing, tilt the slide and place on a Kimkwipe. Gently add 500 uL 1x TE buffer to rGO slide, collecting with Kimwipe. Some rGO might move off the slide, and that's ok as long as not all of it leaves. It is crucial to add the 1x TE buffer slowly such that rGO doesn't move.
4. Place the slide on the table (with kimwipe underneath if the slide is wet). Place the lid on top and cover with foil. Let it air dry for 30 mins before imaging.

Close

Read about our work with rGO here:

rGO

rGO Experiments

Background Information

Microbial Synthesis of rGO and O.D.₆₀₀ measurements

Materials

• 15ml conical tubes
• TSB + antibiotic (only use antibiotic if desired strain is resistant)
• serological pipette
• GO solution (Graphenea, 4g/L)
• 1M IPTG (or other gene inducer, depending on strain)
• Bacterial overnights
• plate reader
• Disposable plastic cuvettes

Protocol

1. Set up 5ml overnight cultures of desired Shewanella oneidensis MR-1 strains in the appropriate TSB+antibiotic media including wildtype (no antibiotic).

Note: After this step, IPTG can be added directly to the overnight cultures to allow for gene expression to be induced for a specified amount of time before initiating reduction (see step 4 for adding IPTG) . Otherwise immediately continue to the next step.

2. Prepare a TSB +antibiotic medium with a GO concentration of .5 mg/mL.
3. Dilute the overnight cultures of S. oneidensis MR-1 strains to an O.D.₆₀₀ of 0.1 within the GO and TSB solution.
4. Add IPTG to the TSB+antibiotic and GO solution to obtain the final desired concentration (for example: 7.5ul added to 5ml GO solution for a concentration of 1500uM IPTG from a 1M stock).
5. Set up separate tubes with just TSB+antibiotic (no GO) and repeat steps 2-3 for each condition.
6. Pipette 150ul of each sample into the well of a 96-well plate and repeat until each sample has been pipetted in triplicates. Then pipette 150ul of TSB only into the wells in triplicates , GO only (no TSB) into the wells in triplicates and TSB+antibiotic and GO only into the wells in triplicates as the controls and blank, respectively.
7. Incubate all the samples in the tubes at room temperature under continuous shaking of 200 rpm. Incubate the 96 well plate in a plate reader at room temperature under continuous shaking at 200 rpm. The plate reader should be set to read O.D.₆₀₀ of the plate every hour for a 48 hour period.

Close

As a control, we also prepared rGO that was reduced by ascorbic acid.

Chemical Reduction

Materials

• Hot plate
• Stir bar
• GO solution (4g/L)
• Ascorbic Acid
• Water

Protocol

1. Create an aqueous solution of ascorbic acid (0.1g/mL).
2. Heat an equal volume of GO solution to 95 degrees °C.
3. Slowly add the ascorbic acid solution to the GO.
4. Stir for one hour.

Note: Solution should become much darker

Before and after reduction

Close

After synthesizing rGO, it has to be washed to remove impurities.

Washing rGO

Note: Washed rGO tends to aggregate. Sonicate the rGO for one hour before using it.

Washing Bacterially Reduced rGO

1. Centrifuge the solution at 8000 rpm for 8 minutes.
2. Dispose of the supernatant and replace with DI water.
3. Repeat steps 1-2 three times for a total of 4 rounds of centrifugation.
4. Pour off as much supernatant as possible. Freeze the rGO at -80 degrees °C overnight. This will help with Raman sample preparation and will also kill any remaining bacteria.

Note: Bacterially reduced rGO is too fine to filter with standard filter paper.

Washing Chemically Reduced rGO

1. Let the solution cool to room temperature.
2. Filter it via gravity filtration.
3. Wash the rGO with 10 mL 1M HCl

Note: Use a glass pipette for this step. It is much less likely to melt than a plastic one.

4. Wash the rGO with water until the filtrate has a neutral pH.
5. Resuspend the rGO in water to create a solution of the desired concentration. Freeze ~1 mL for Raman spectroscopy.

Close

After washing the rGO, it needs to be characterized. We used Raman spectroscopy for this.

Taking Raman

Sample Preparation

1. Prepare a concentrated sample of rGO or GO. If the sample is frozen or very concentrated (4mg/mL), it will be less likely to spread and you are more likely to get a better spectrum.
2. Deposit the rGO (or other sample) on a glass or quartz slide. Quartz has lower background noise and is reusable, but is significantly more expensive.

Note: The sample can also be deposited between two slides if drop casting it is not possible. Do note that very strong glass or quartz peaks are expected in this case.

3. Allow the rGO to dry. A dessicator or dehydrator can be used, or it can be left at room temperature for a few days.

Taking the Spectra

Note: It is HIGHLY reccomended that you get help from whichever department's Raman you use.

1. Set the Raman to 532 nm.

Note: As rGO and GO are both very dark, setting the laser to a high intensity may be needed to get a spectrum. Be careful not to let your sample burn.

2. Focus the machine. This process will vary depending on the Raman setup. The goal is to have a flat baseline and distinct peaks at ~1400 and ~1600 cm^-1.

Close

In order to transform our bacteria, we needed to amplify our genes of interest to clone them into a plasmid. Unfortunately, we did not have access to standard PCR conditions. In order to find our ideal annealing temperature, we used touchdown PCR to amplify our genes.

Touchdown PCR

Materials

• Q5 DNA polymerase from neb (.25 uL, 0.02U/uL)
• Q5 rxn buffer (5uL)
• dNTPs (.5 uL)
• Forward primer (10 uM, 1:10 dilution, 1.25 uL)
• Reverse primer (10 uM, 1:10 dilution, 1.25 uL)
• DNA (1 uL)
• Water (15.75 uL)

Protocol

1. Combine all reagents in a pcr tube. Add the enzyme last, and make sure to keep it and the DNA cold.
2. Run the following cycle in a thermocycler

Step	Temperature ()	Duration
Initial denaturation	98 °C	30s
20 cycles	98°C	10s
	64°C (-1°C per cycle)	30s
	72°C	4 minutes
25 cycles	98°C	10s
	54°C	30s
	72°C	4 minutes
Final step	72°C	2 minutes
Hold	23°C	Indefinite

Close

After transforming our genes, we needed to clone them into a plasmid. We chose the pcD8 plasmid because it is compatible with S. oneidensis and is inducible with IPTG. In order to get our gene into the plasmid, we used neb's golden gate cloning.

Golden Gate Cloning

Protocol

1. Two heat blocks were heated; one to 37°C and one to 60°C
2. Calculations were made using NEBioCalculator® to achieve a molar ratio of 2:1 insert: vector for each cloning reaction and their respective inserts.
3. The golden gate reaction mixtures were set up using the NEB® Golden Gate Assembly Kit (BsaI-HF®v2) as follows:

Component	Quantity (uL)
Plasmid (pcD8 19.1ng/ul - 75ng)	3.93
Gene insert (19.8 ng/ul)	2.17
T4 DNA Ligase Buffer (10X)	2
NEB Golden Gate Assembly Mix	1
Nuclease- free water	10.9
Plasmid (pcD8 28.7ng/ul - 75ng)	2.61

4. The negative control was set up using the ydeH mixture without the ydeH insert and making up the rest of the reaction in nuclease free water.
5. The mixtures were placed on the 37C heating block for 5 minutes and then on the 60C heating block for 5 minutes
6. The cloned mixtures were then placed at 4C.

Close

Now that we had our cloned plasmids, we needed to transform them into bacteria. We used electrocompetent S. oneidensis.

Transformation

Materials

• Plasmid DNA(100ng)
• Shewanella oneidensis liquid culture( about 1 ml)
• 10%v/v glycerol
• LB
• Eppendorf tubes
• Centrifuge
• Electroporation cuvette (Bio-Rad 0.1)
• LB+agar+antibiotic plates

Protocol

1. Culture a colony overnight in LB liquid medium at 30 °C with shaking (~200 rpm)
2. Make 10% v/v glycerol
3. Obtain purified plasmid DNA
4. DNA concentration should be ~100 ng/μL. Use a Nanodrop if available; if 260/230 and 280/260 ratios are not ~2 and ~1.8, respectively, the efficiency may be lower.
5. Prepare ~100 ng plasmid DNA in a 2 mL Eppendorf tube. Perform a control with no DNA.
6. Collect 1 mL of an overnight culture in an Eppendorf tube, spin at 7,906 rcf for 1 min
7. Wash cells three times with 1 mL 10% glycerol, spinning at 7,906 rcf for 2 min each time
8. After the third wash, decant the glycerol and leave ~70 μL for suspending the cells
9. When decanting the glycerol after spins, use care to not lose the pellet; gently tap the tube 3 times.
10. Suspend cells gently by shaking the tube and pipetting with P100, then mix cells with the DNA (~3ul) by swirling around with the tip

Note:A 0.1 cm electroporation cuvette fits ~100 μL max. A 0.2 cm cuvette requires higher voltages.

11. Transfer the mixture immediately into a 0.1 cm cuvette without making bubbles
12. Electroporate at 1.2 kV. Time constant (tc) should be ~5
13. Quickly add 1 mL LB into the cuvette and mix gently while pipetting up and down
14. Transfer the cells into the same 2 mL tube you had the DNA in
15. Let the cells recover at 30 °C for 2 h with shaking ~200 rpm
16. Make 10x serial dilutions in LB.
17. Plate several different dilutions spreading out LB-agar-antibiotic plates.
18. Incubate plates at 30 °C for ~24-36 h until colonies appear.

Close

After transformation, we wanted to make sure that our cloning was successful. In order to do that, we ran a digest to cut our plasmid into pieces. We could then compare the resulting gel with computer simulations and find out if we had been successful in transforming our bacteria.

Digest

Materials

• NEB KpnI (10U/uL)
• Nuclease free water
• 10X NEBuffer r1.1
• DNA
• Heat block
• 1.5 mL microcentrifuge tube

Protocol

1. Set a heat block to 37 C.
2. Ensure that the reagents are kept on ice while setting up the reaction.
3. In a 1.5mL microcentrifuge tube set up the digest mixture in the following order:

Component	25 uL reaction
Nuclease-free Water	17 uL
10X NEBuffer r1.1	2.5 uL
DNA	5 uL
NEB KpnI	5 uL

4. Ensure the reaction mixture is well mixed and incubate in the heat block for 1 hour at 37 C.
5. Immediately remove from the heat block after 1 hour and perform gel electrophoresis to visualize the digest products.

Close

Read about our work with aptamers here:

Aptamers