Team:St Andrews/Experiments

Template

Experiments

Cell culturing

1. Making chemically competent cells

Materials

Cells
0.1 M CaCl2
10% Glycerol
LB media
250 mL conical flasks
50 mL Falcon tubes
Ice bucket
Pipette / Pipette gun

Protocol

Prepare 0.1 M CaCl2, filter-sterilize or autoclave and pre-chill to = or <4deg; you will need ~30 ml of that per 50ml cells so prepare 100-200 ml
In a sterile tube (e.g. a Falcon or a white-capped tube) mix the above solution with sterile glycerol to obtain final glycerol concentration of 10% (e.g. 8ml CaCl2 and 2ml sterile 50% glycerol), keep that chilled as well. This is the solution for storing competent cells.
In the afternoon/evening inoculate a small (5-10ml) LB media vial with cells from a colony or a glycerol stock (don’t thaw the glycerol stock, just scrape some frozen cells off the surface) and incubate overnight in the shaking incubator at 37deg C overnight
Next day use the culture to inoculate 50-100 ml LB in a 250ml conical flask (~1 ml per 50ml media) and keep the flask in the shaking incubator at 37 deg
After about an hour start measuring the optical density at 600nm, if it is less than 0.2 check again in half an hour, if it is getting closer to 0.3 then check every 15-20 min after that (they are entering exponential phase so will accelerate!)
When the optical density is 0.5-0.8 transfer the cells into a fresh 50ml Falcon tube, or several tubes if you have more than 50ml, and spin for 10 min at >3000g (if they go above 1 they are entering stationary phase, either abandon ship or dilute them in a fresh flask with fresh media 10x and then keep checking OD again)
Pour off the media from the Falcon (ideally back in the conical flask) and you will see a pellet at the bottom. From now on KEEP CELLS ON ICE.
Resuspend the cells in pre-chilled 0.1M CaCl2 using the long sterile pipette and pipette gun, use ~30 ml per Falcon tube
Incubate the resuspended cells on ice for 1-2 hours
Spin the cells at 4deg, >3000g for 10 min
Resuspend the cells in 0.1M CaCl2/10%glycerol solution by gently pipetting them up and down and occasionally swirling them in the tube, use ~1-2 ml of the storage solution per Falcon tube (see comment below)
Keeping all the tubes and cells on ice, aliquot the cells into sterile eppendorfs and freeze the cells at -80deg C
For BL21/soluBL21 cells I would use 2ml of storage solution to resuspend make lots of 50-100μl aliquots, plus a few larger volumes (e.g. 200 or 300 or 500 μl). You will need 25-50 μl cells per transformation of a plasmid and these are the “easy” transformations. Larger volumes can save you space in the freezer boxes, and if you are transforming several plasmids in one go you can defrost 1 larger aliquot and transfer smaller aliquots from that into separate tubes.
For cloning strains like DH5alpha I resuspend cells in 1 ml storage solution and aliquot 100μl (or multiples of that, as explained for BL21 cells) into sterile tubes. For each ligation reaction/mutagenesis reaction transformation I would use 100μl cells. You can also make a few smaller aliquots, say 50μl, which can be used for plasmid transformations -e.g. when you want to make more of that plasmid for further cloning.

2. Transformation

Materials

Cells
Plasmid
LB media
Ice bucket
Eppendorf tubes 1.5 mL
Pipette
Water bath

Protocol

Thaw an appropriate volume of cells on ice, it can take 5-20 min. you can speed it up by occasionally flicking the tube. After they thaw they may look a bit clumpy – then stir them with pipette tip and if necessary (see above), aliquot them into separate sterile tubes.
For standard plasmid transformation of either cell type (clean plasmid, nothing difficult) I would add 0.5-1μl plasmid per cell aliquot, for the more complex things I would add 10μl of ligation/mutagenesis reaction per 100μl DH5alpha cells
After adding plasmid/reaction, flick the tube a couple of times to ensure mixing
Incubate the cells on ice for at least 10 min (for the challenging transformations perhaps 20 or so), it won’t hurt cells if you leave them much longer than that
Heat shock by putting the cells in 42deg C water bath for 30 sec (you can use a floater if you have many tubes)
Put the cells back on ice for a couple of minutes; for standard transformations you can shorten or even skip this step completely
Add ~200-300μl fresh LB media per tube for standard transformation; for challenging ones use 0.9-1 ml, preferably pre-warmed to 37degC, and you can use SOC or other rich media which would enhance the cell growth. Some people even use 14 ml round-bottom Falcon tubes, not eppendorfs, for these transformations to ensure good aeration…
Recovery step: incubate the cells in the shaking incubator at 37deg for 30-60 min (standard transformation) or 90-120min (challenging ones); I either put the tubes at the bottom of the beaker or tape them down to the bottom of the incubator. Importantly, for better aeration they should be lying flat rather than vertically.
For plating standard transformations I plate 50-100μl cells on a plate. For challenging ones I spin the tubes for a couple of minutes at ~6000g, discard most of the supernatant, and resuspend the pellet in whatever remains (100-200μl) or just add some more fresh media. That maximises the cell concentration. To further maximise the chances of success you can pre-warm the plates to 37degC and use a couple of plates per transformation (each covered with 50-100μl cells).
Incubate the cells overnight at 37degC INVERTED to prevent condensation on the actual plate.
Next day (preferably don’t leave them growing for longer than 16hours) inspect the plates and put them in the fridge or keep on the bench (inverted!) if you are going to use them later in the day to inoculate
If it is Friday and you want to transform then instead of 37deg C you can leave the plates on the bench, at room temp for 2-3 days and they should grow at the same rate as overnight at 37deg. I would then normally inoculate cultures on Sunday evening. For expression strains I would never use a plate older than a few days to inoculate an overnight culture for overexpression as cells may be slowly mutating even in the cold!

3. Measuring bacterial transformation efficiency

Protocol:

1 Transform 1 aliquot of competent cells with 1 ng of a control plasmid
Plate 5 µl, 50 µl and 300 µl of in plates with the appropriate antibiotic
Incubate overnight at 37ºC
Count the colonies on the plate where you can find 30 to 300 isolated colonies
Use this equation to calculate the number of CFUs per micrograms of DNA:

µg CFU = Volume plated Colonies in the plate·(Volume of cells+Volume of SOC +Volume of DNA)

CFU/g=Colonies in the plate(Volume of cells+Volume of SOC +Volume of DNA)/ Volume plated +1000

4. Bacterial glycerol stock

For storing different strains or cells that contain plasmids; instead of transforming fresh cells every time to re-prep plasmids you can just scrape some cells from the glycerol stock to inoculate an overnight culture. Glycerol stocks are made of expression strains (e.g. soluBL21 DE3) containing different expression plasmids (e.g. pHisTevM3110C) so that they can be directly inoculated overnights for expression. Some people are opposed to that idea, and always do a fresh transformation.

Materials:

Overnight cell culture
Glycerol
MilliQ water

Protocol:

As glycerol is very viscous, make 100 ml of 50% glycerol (mixed with Milli Q water) and autoclave that in advance.
To prepare a glycerol stock, mix ~0.5 ml of cells from an overnight culture with 0.5 ml of 50% glycerol in a sterile Eppendorf, invert a few times to ensure proper mixing and put the tube in the -80deg freezer.

5. Growing up E.Coli essential procedures

Liquid medium preparation

Materials:

Agar
LB broth salt-free
NaCl
Distilled water
500 ml autoclavable bottle
Autoclave
Magnetic stirrer

Protocol:

Add 10g of Agar, 7.5 g of yeast extract and 5g of NaCL.
Mix them.
Add the magnetic stirrer.
Add distilled water until liquid reaches 500 mL.
Stir solution.
Put bottle into autoclave.

Solid medium preparation

For 500 ml of medium for 20 plates:

Materials:

Agar
LB broth salt-free
NaCl
Distilled water
500 ml autoclavable bottle
Autoclave
Magnetic stirrer
Petri dishes
1000x antibiotic

Protocol:

Add 10g of Agar, 7.5 g of yeast extract and 5g of NaCL.
Mix them.
Add the magnetic stirrer.
Add distilled water until liquid reaches 500 mL.
Stir solution.
Put bottle into autoclave.
Set LB medium to cool to 50 degrees while stirred
Pour LB media into plates.
Set plates to cool.
After1 hour set plates upside down and leave on the bench overnight
Store plates in a cold room.

Liquid medium culture (inoculation)

After plating transformed competent cells with a ligation product, single colonies are picked and inoculated into liquid medium.

Materials

Shaker
Vials
LB broth liquid medium
Antibiotic (1000x)

Protocol:

Add 10 ml of LB broth and 10 µl of the antibiotic into a vial per colony that will be picked. A single colony is picked up and inoculated into the liquid medium with a pipette tip and this culture is then incubated overnight (12-16 h) at 37 ºC with vigorous shaking (220 rpm).

Petri dish culture

Cells are grown in solid agar medium in order to isolate specific colonies. Agar petri dishes must contain the antibiotic for the resistance carried by the transformed strain.

Materials:

LB agar plates
Sterile spreaders

Protocol:

Spread the bacteria culture all over the plate using the spreader.
Invert plates and incubate overnight at 37ºC.

PCR, agarose gels and DNA purifcation

1. Re-suspension of iGEM DNA

Re-suspending dried DNA from the iGEM distribution kit. Protocol taken from the iGEM website.

Protocol:

Note: There is an estimated 2-3ng of DNA in each well. When following this protocol, assume that you are transforming with 200-300pg/µL

With a pipette tip, punch a hole through the foil cover into the corresponding well of the part that you want. Make sure you have properly oriented the plate. Do not remove the foil cover, as it could lead to cross contamination between the wells.
Pipette 10µL of dH2O (distilled water) into the well. Pipette up and down a few times and let sit for 5 minutes to make sure the dried DNA is fully resuspended. The resuspension will be red, as the dried DNA has cresol red dye. We recommend that you do not use TE to resuspend the dried DNA.
Transform 1µL of the resuspended DNA into your desired competent cells, plate your transformation with the appropriate antibiotic* and grow overnight.
Pick a single colony and inoculate broth (again, with the correct antibiotic) and grow for 16 hours.
Use the resulting culture to miniprep the DNA AND make your own glycerol stock.

2. PCR Clean- up

NEB Monarch® PCR & DNA Cleanup Kit (5 μg)

DNA Cleanup and Concentration: for the purification of up to 5 μg of DNA (ssDNA > 200 nt and dsDNA > 50 bp) from PCR and other enzymatic reactions.

Oligonucleotide Cleanup: for the purification of up to 5 μg of DNA fragments ≥ 15 bp (dsDNA) or ≥ 18 nt (ssDNA). Expected recovery is > 70%. When purifying ssDNA of any size, recovery can be increased by using this protocol; however, it is important to note that this protocol shifts the cutoff for smaller fragments to 18 nt (rather than 50 nt for the DNA Cleanup and Concentration Protocol).

General Guidelines:

Input amount of DNA to be purified should not exceed the binding capacity of the column (5 μg). A starting sample volume of 20–100 μl is recommended. For smaller samples, TE can be used to adjust the volume to the recommended volume range. Centrifugation should be carried out at 16,000 x g in a standard laboratory microcentrifuge at room temperature.

Protocol:

Add ethanol to Monarch DNA Wash Buffer prior to use (4 volumes of ≥ 95% ethanol per volume of Monarch DNA Wash Buffer).
Always keep all buffer bottles tightly closed when not in use
All centrifugation steps should be carried out at 16,000 x g. (~13K RPM in a typical microcentrifuge). This ensures all traces of buffer are eluted at each step.
Dilute sample with DNA Cleanup Binding Buffer according to the table below. Mix well by pipetting up and down or flicking the tube. Do not vortex. A starting sample volume of 20–100 μl is recommended. For smaller samples, TE can be used to adjust the volume. For diluted samples larger than 800 μl, load a portion of the sample, proceed with Step 2, and then repeat as necessary.

SAMPLE TYPE	RATIO OF BINDING BUFFER: SAMPLE	EXAMPLE
dsDNA > 2 kb (plasmids, gDNA)	2:1	200μl:100 μl
dsDNA < 2 kb(some amplicons, fragments)	5:1	500 μl:100 μl
ssDNA > 200 nt*	7:1	700 μl:100 μl

Insert column into collection tube and load sample onto column and close the cap. Spin for 1 minute, then discard flow-through.
To save time, spin for 30 seconds, instead of 1 minute.
If using a vacuum manifold* instead of centrifugation, insert the column into the manifold and switch the vacuum on. Allow the solution to pass through the column, then switch the vacuum source off.
Re-insert column into collection tube. Add 200 μl DNA Wash Buffer and spin for 1 minute. Discarding flow-through is optional.
If using a vacuum manifold, add 200 μl of DNA Wash Buffer and switch the vacuum on. Allow the solution to pass through the column, then switch the vacuum source off.
Repeat wash (Step 3).
* Make sure to follow the manifold manufacturer's instructions to set-up the manifold and connect it properly to a vacuum source.
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.
If using a vacuum manifold: Since vacuum set-ups can vary, a 1 minute centrifugation is recommended prior to elution to ensure that no traces of salt or ethanol are carried over to the next step.
Add ≥ 6 μl of DNA Elution Buffer to the center of the matrix. Wait for 1 minute, then spin for 1 minute to elute DNA.
Note: 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. For larger size DNA (≥ 10 kb), heating the elution buffer to 50°C prior to use can improve yield.
Care should be used to ensure the elution buffer is delivered onto the matrix and not the wall of the column to maximize elution efficiency. To save time, spin for 30 seconds, instead of 1 minute.

DNA Clean- up and Concentration System (ReliaPrep)

Materials:

100% isopropanol, RNase-free
95–100% ethanol, RNase-free
1.5ml microcentrifuge tubes • agarose gel (standard or low-melt; only for gel purification)
1X TAE or TBE electrophoresis buffer (only for gel purification)
50–65°C heating block (only for gel purification)
microcentrifuge capable of maintaining at least 10,000 × g (14,000rpm)
Kit

Protocol:

Note: Perform all centrifugation steps at 10,000 × g (14,000rpm).

Pipet 25–400µl of a PCR amplification or reaction pool into a 1.5ml microcentrifuge tube.
Add an equal volume of Membrane Binding Solution and vortex 5 seconds.

PCR Volume (µl)	Membrane Binding Solution (µl)	Total Load (µl)
25	25	50
50	50	100
100	100	200
150	150	300
200	200	400
250	250	500
300	300	600
350	350	700
400	400	800

Load sample onto a ReliaPrep™ Minicolumn seated in a Collection Tube and centrifuge for 30 seconds.
Remove column and discard the contents of the Collection Tube. Reseat the minicolumn into the same Collection Tube.
Add 200µl of Column Wash Solution (CWE) and centrifuge for 15 seconds. Remove column and discard the contents of the Collection Tube. Reseat the minicolumn into the same Collection Tube.
Wash with 300µl of Buffer B (BWB) and centrifuge for 15 seconds. Repeat wash with 300µl of Buffer B (BWB) and centrifuge again.
Remove minicolumn, and discard the contents of the Collection Tube. Reseat the minicolumn into the same Collection Tube and centrifuge for 1 minute to dry the minicolumn; then transfer minicolumn to an Elution Tube.
Pipet 15µl of Nuclease-Free Water or TE buffer (not provided) to the center of the minicolumn, then centrifuge for 30 seconds. Note: Touch the pipette tip to the column bed surface before dispensing Nuclease-Free Water or TE buffer to completely wet the column matrix. The color should change from light to dark tan.
For maximum recovery, repeat elution with an additional 15µl of Nuclease-Free Water or TE buffer.

Wizard® SV Gel and PCR Clean-Up System

Protocol:

Amplify target of choice using standard ampliﬁcation conditions.
Add an equal volume of Membrane Binding Solution to the PCR ampliﬁ cation (Notes 1–4 below).
Notes:

The maximal capacity of a single SV Minicolumn is approximately 1ml of PCR ampliﬁ cation added to 1ml of Membrane Binding Solution (2ml total). For PCR volumes >350µl, continue to pass the sample through the column until all of the sample has been processed.
The maximum binding capacity is approximately 40µg per column, and as little as 10ng has been successfully puriﬁed.
Mineral oil does not interfere with puriﬁcation.
For ampliﬁcation reactions that do not produce a single product or where ampliﬁcation has been ineﬃcient and there is highly visible primer dimer, gel puriﬁcation of the band of interest is recommended. Alternatively, an 80% ethanol wash solution can be substituted for the supplied Membrane Wash Solution to reduce primer-dimer carryover.

Place one SV Minicolumn in a Collection Tube for each dissolved gel slice or PCR ampliﬁ cation.
Transfer the dissolved gel mixture or prepared PCR product to the SV Minicolumn assembly and incubate for 1 minute at room temperature.
Centrifuge the SV Minicolumn assembly in a microcentrifuge at 16,000 × g (14,000rpm) for 1 minute. Remove the SV Minicolumn from the Spin Column assembly, and discard the liquid in the Collection Tube. Return the SV Minicolumn to the Collection Tube. Note: Failure to spin at 16,000 × g (14,000rpm) can result in reduced yield. !
Wash the column by adding 700µl of Membrane Wash Solution, previously diluted with 95% ethanol (Section 4.A), to the SV Minicolumn. Centrifuge the SV Minicolumn assembly for 1 minute at 16,000 × g (14,000rpm). Empty the Collection Tube as before, and place the SV Minicolumn back in the Collection Tube. Repeat the wash with 500µl of Membrane Wash Solution, and centrifuge the SV Minicolumn assembly for 5 minutes at 16,000 × g.
Remove the SV Minicolumn assembly from the centrifuge, being careful not to wet the bottom of the column with the ﬂowthrough. Empty the Collection Tube, and recentrifuge the column assembly for 1 minute with the microcentrifuge lid open (or oﬀ ) to allow evaporation of any residual ethanol.
Carefully transfer the SV Minicolumn to a clean 1.5ml microcentrifuge tube. Apply 50µl of Nuclease-Free Water directly to the center of the column without touching the membrane with the pipette tip. Incubate at room temperature for 1 minute. Centrifuge for 1 minute at 16,000 × g (14,000rpm).
Discard the SV Minicolumn, and store the microcentrifuge tube containing the eluted DNA at 4°C or –20°C. Note: The volume of the eluted DNA will be approximately 42–47µl. If the DNA needs to be further concentrated, perform an ethanol precipitation. Alternatively, the DNA may be eluted in as little as 15µl of Nuclease-Free Water without signiﬁ cant reduction in yield. If using an elution volume of 15µl, verify that the membrane is completely covered with Nuclease-Free Water before centrifugation. Elution volumes less than 15µl are not recommended.

Elution Volume	Percent Recovery compared to 50 L
10 L	35 %
15 L	98%
25 L	98%
50 L	100%
75L	100%
100L	100%

3. PCR - NEB Q5-HF Kit

Materials:

50uL PCR Tubes
Ice Box
Nuclease-Free dH2O
10uM Forward Primer
10uM Reverse Primer

Primers should be at 100um, but check

Template DNA
Q5 High-Fidelity 2X Master Mix
PCR Machine

Protocol:

Get out primers from -20C freezer (4th drawer down, on the left), and keep on ice
Get the Q5 High 2X Master Mix kit from the -20C freezer (4th drawer down, on the left), and keep on ice
Add the following to PCR Tube, adding the Q5 High-Fidelity 2X Master Mix Last:

Q5 High-Fidelity 2X Master Mix	25uL
Template DNA	Variable (~0.2uL for Plasmids)
10uM Forward Primer	2.5uL
10uM Reverse Primer	2.5uL
Nuclease-Free dH2O	Make up to 50uL

Keep the above mixture on ice until you are ready to put it into the machine
Turn on PCR machine at the back and wait for it to boot. Once booted, select the program you intend to run, or create a new program. The program should have:

Initial Denaturation	98C	30s
Denaturation	98C	15-30s
Annealing	45-68C	15-60s
Extension	72C	20s per kb of template DNA
Final Extension	72C	5 minutes
Hold	4-10C	-

The annealing temperature should be ~3C below the lowest melting temperature of the two primers.
If you want to keep the PCR product in the machine overnight, set the time to infinite on the machine and the final hold temperature to 4C. Steps 2-4 in the table above should be repeated for 30 cycles.

4. Agarose Gel

Gel preparation and running

Makes 1L of 50X TAE Buffer for running DNA gels.

Materials:

Tris Base
Disodium EDTA
Glacial Acetic Acid
1L Beaker
1L Shop Bottle (Empty and Clean)
Magnetic Stirrer (Optional)

Protocol:

Weigh out 242g of Tris and add to 800mL of dH2O in the 1L beaker. Stir until all the Tris has dissolved.
Once Tris dissolved, add 18.61g of Disodium EDTA to the mixture. Stir until dissolved.
Once EDTA dissolved, add 57.1mL of Glacial Acetic Acid and stir.
Make the mixture up to 1L with dH2O. Transfer to 1L shop bottle and cap.

Making 50x TAE buffer for gel

Making Agarose gels to run Nucleic acid samples. Makes circa 10 1% Agarose gels.Wear gloves throughout as risk of burns from Agarose, and to avoid skin contact with SYBR Safe.

Materials:

Agarose Powder
1x TAE Buffer (Diluted from 50x Buffer)

500mL Shop Bottle
Gel Cast
Gel Comb
Gel Clamp

Protocol:

Making the Gel Stock:

Weigh out 5g of Agarose Powder, and transfer with x1 TAE Buffer to a clean 500mL Shop Bottle.
Top up the shop bottle to 500mL with x1 TAE Buffer.
Microwave the mixture in the bottle until the Agarose powder is dissolved. This takes circa 6 mins of microwave time, but ensure to swirl and mix the mixture every 2 mins or so. Mix upon boiling.
The gel can now be left to cool before pouring or left to cool on the bench top. Once solidified, the cap is placed on the bottle.
You can now microwave the agarose shop bottle to prepare more gels.

Casting the Gel

Place the Gel cast in the gel clamp and clamp tight.
Once the molten agarose bottle is just cool enough to hold, pour in the agarose to the cast.
If you are unsure about what volume to use, measure out 40-50mL of molten Agarose in a measuring cylinder.
Add 1 uL of SYBR Safe and stir round the molten agarose using the pipette tip.
Place the comb in the cast. Leave the gel to set.

Running the Gel

Remove Gel + Cast from clamp and place in the gel tank, ensuring the gel is covered in x1 TAE buffer.
Remove the comb from the gel gently, once submerged in x1 TAE Buffer
Load samples into the gel
Run the Gel for 1hr at 90V in most conditions, but you can use a lower voltage and longer time if needs be.

5. DNA gel extraction

NEB Monarch kit

The input amount of DNA to be purified should not exceed the binding capacity of the column (5 μg). DNA fragments were excised from an agarose gel and are diluted by the addition of four volumes of Gel Dissolving Buffer. For a typical 100 mg (100 μl) gel slice, 400 μl of Gel Dissolving Buffer is added. Centrifugation should be carried out at 16,000 x g in a standard laboratory microcentrifuge at room temperature.

Buffer Preparation:

Add ethanol to Monarch DNA Wash Buffer prior to use (4 volumes of ≥ 95%ethanol per volume of Monarch DNA Wash Buffer).
For 50-prep kit add 20 ml of ethanol to 5 ml of Monarch DNA Wash Buffer
For 250-prep kit add 100 ml of ethanol to 25 ml of Monarch DNA Wash Buffer. Always keep all buffer bottles tightly closed when not in use.

Protocol:

Excise the DNA fragment to be purified from the agarose gel using a razor blade, scalpel or other clean cutting tool. Use care to trim excess agarose. Transfer it to a 1.5 ml microcentrifuge tube and weigh the gel slice.
Note: Using UV light to visualize the slice is common, but the exposure time should be kept as short as possible to minimize damage to the DNA. Use long-wave UV when possible, as shorter wavelengths induce greater damage. Also, trim off excess agarose from the perimeter of the band to minimize the amount of dissolving buffer needed, and to reduce the time necessary to extract the DNA.
Add 4 volumes of Monarch Gel Dissolving Buffer to the tube with the slice.
Note: If the volume of the dissolved sample exceeds 800 μl, the loading of the sample onto the column should be performed in multiple rounds to not exceed the volume constraints of the spin column.
Incubate the sample between 37–55°C (typically 50°C), vortexing periodically until the gel slice is completely dissolved (generally 5–10 minutes).
Note: For DNA fragments > 8 kb, an additional 1.5 volumes of water should be added after the slice is dissolved to mitigate the tighter binding of larger pieces of DNA (e.g., 100 μl gel slice: 400 μl Gel Dissolving Buffer: 150 μl water). Failure to dissolve all the agarose will decrease the recovery yield due to incomplete extraction of the DNA and potential clogging of the column by particles of agarose.
Insert the column into collection tube and load sample onto the column. Spin for 1 minute, then discard flow-through.
To save time, spin can be reduced to 30 seconds.
If using a vacuum manifold* instead of centrifugation, insert the column into the manifold and switch the vacuum on. Allow the solution to pass through the column, then switch the vacuum source off.
Re-insert column into collection tube. Add 200 μl DNA Wash Buffer and spin for 1 minute. Discarding flow-through is optional.
If using a vacuum manifold, add 200 μl of DNA Wash Buffer and switch the vacuum on. Allow the column solution to pass through the column, then switch the vacuum source off.
Repeat wash (Step 5).
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 in doubt, re-spin for 1 minute before placing into clean microfuge tube.
If using a vacuum manifold: Since vacuum set-ups can vary, a 1 minute centrifugation is recommended prior to elution to ensure that no traces of salt and ethanol are carried over to the next step.
* Make sure to follow the manifold manufacturer's instructions to set-up the manifold and connect it properly to a vacuum source.
Add ≥ 6 μl of DNA Elution Buffer to the center of the matrix. Wait for 1 minute, and spin for 1 minute to elute DNA.
Note: 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. For larger size DNA (≥ 10 kb), heating the elution buffer to 50°C prior to use can improve yield. Care should be used to ensure the elution buffer is delivered onto the matrix and not the wall of the column to maximize elution efficiency. To save time, spin can be reduced to 30 seconds.

6. Ethanol precipitation of plasmid DNA

DNA is polar due to its highly charged phosphate backbone. This polarity, based on the principle of "like dissolves like", makes it soluble in water, which is also highly polar. Ethanol is much less polar than water, its dielectric constant is 24.3 (at 25 °C). This means that adding ethanol to solution disrupts screening of charges by water. If enough ethanol is added electrical attraction between phosphate groups and any positive ions present in solution becomes strong enough to form stable ionic bonds and precipitate DNA. This usually happens when ethanol makes around 64% of the solution. As the mechanism suggests solution has to contain positive ions for precipitation to occur, usually Na+, NH4+ or Li+ play this role.

Optimal incubation time depends on the length and concentration of DNA. Smaller fragments and lower concentrations will require longer times to achieve the same recovery. For very small lengths and low concentrations overnight incubation is recommended. In such cases the use of carriers like tRNA, glycogen or linear polyacrylamide can greatly improve recovery. During incubation DNA and some salts will precipitate from solution, in the next step this precipitate is collected by centrifugation in a microcentrifuge tube at high speeds (~12,000g). Time and speed of centrifugation has the biggest effect on DNA recovery rates. Smaller fragments and higher dilutions require longer and faster centrifugation. Centrifugation can be done either at room temperature or in 4 °C or 0 °C. During centrifugation precipitated DNA has to move through ethanol solution to the bottom of the tube, lower temperatures increase viscosity of the solution and larger volumes make the distance longer, so both those factors lower efficiency of this process requiring longer centrifugation for the same effect. After centrifugation the supernatant solution is removed, leaving a pellet of crude DNA. Whether the pellet is visible depends on the amount of DNA and on its purity (dirtier pellets are easier to see) or the use of co-precipitants. 70% ethanol is then added to the pellet, and the sample gently mixed to break the pellet loose and wash it. This removes some of the salts present in the leftover supernatant and bound to DNA pellet making the final DNA cleaner. This suspension is centrifuged again to once again pellet DNA and the supernatant solution is removed. This step is repeated once. Finally, the pellet is air-dried and the DNA is resuspended in water or other desired buffer. It is important not to over-dry the pellet as it may lead to denaturation of DNA and make it harder to resuspend. Isopropanol can also be used instead of ethanol; the precipitation efficiency of the isopropanol is higher making one volume enough for precipitation. However, isopropanol is less volatile than ethanol and needs more time to air-dry in the final step. The pellet might also adhere less tightly to the tube when using isopropanol.

Materials:

Sodium Acetate (3M, pH 5.2)
Ethanol (>95 %)
Ethanol (70 %)
Water as buffer
Sample

Protocol:

Add 1/10 volume of Sodium Acetate (3 M, pH 5.2).
Add 2.5-3.0 X volume (calculated after addition of sodium acetate) of at least 95% ethanol
Incubate on ice for 15 minutes. In case of small DNA fragments or high dilutions overnight incubation gives best results, incubation below 0 °C does not significantly improve efficiency.
Centrifuge at > 14,000 x g for 30 minutes at room temperature or 4 °C.
Discard supernatant being careful not to throw out DNA pellet which may or may not be visible.
Rinse with 70% Ethanol (see note above for details)
Centrifuge again for 15 minutes.
Discard supernatant and dissolve pellet in desired buffer. Make sure the buffer comes into contact with the whole surface of the tube since a significant portion of DNA may be deposited on the walls instead of in the pellet. o This last step should be done with sterile TE.

Plasmid preparation

1. Miniprep

NEB Monarch Miniprep

Materials:

Monarch Plasmid Resuspension Buffer
Monarch Plasmid Lysis Buffer
Monarch Plasmid Wash Buffer 1
Monarch Plasmid Wash Buffer 2
Monarch DNA Elution Buffer
Monarch Plasmid
Miniprep Columns
Cell Culture
15mL Falcon Tubes
1.5mL Eppendorf Tubes

Protocol:

Culture preparation:

Divide culture evenly between 15mL Falcon Tubes. Try to have no more than 5mL per tube, and no less than 1mL.
Centrifuge cultures in the falcon tubes at max speed for 30s and discard supernatant
Resuspend pellet in 1mL of dH2O and transfer to 1.5mL eppendorf
Centrifuge culture again at max speed for 30s in 1.5mL eppendorf, discard supernatant

Miniprep:

Before beginning the miniprep, ensure that Ethanol has been added to Plasmid Wash Buffer 2, in the correct amount. Correct amount is 4 Ethanol : 1 wash buffer but it should say on the bottle how much to add. If it has already been added, don't add more!

Resuspend the pellet in the eppendorf using 200uL Monarch Plasmid Re-suspension Buffer. Gently pipette up and down to ensure the pellet is completely re-suspended (vortex if necessary)
Add 200uL of Monarch Plasmid Lysis Buffer. Invert tube immediately and gently 5-6 times. Colour should change from light to dark pink. Incubate at room temperature for 1 min. Avoid incubating for longer than 1 min.
Add 400uL of Monarch Plasmid Neutralisation Buffer and gently invert until colour becomes uniformly yellow and precipitate has formed.
Spin the mixture in the eppendorf down at 16000xg for 2-5 mins.
Transfer supernatant to Plasmid Miniprep Spin column and spin at 16000xg for 1 min. Discard flow-through.
Add 200uL of Plasmid Wash Buffer 1 to column and centrifuge at 16000xg for 1 min. You can discard the flow-through at this step but it is only necessary if you think the tip of the column will come into contact with the flow-through - at no point allow this to happen! If it has, spin at 16000xg again for 1 min and proceed as normal.
Add 400uL of Plasmid Wash Buffer 2 to the column and centrifuge at 16000xg for 1 min. Discard flow-through.
Transfer column to clean 1.5mL eppendorf tube.
Add >30uL of DNA Elution Buffer to the mixture. Adding more will increase yield slightly but decrease product concentration. Spin the column at 16000xg for 1 min. Remove column. Do not discard flow-through, this is your DNA product.
Store DNA product labelled at -20C.

2.Eznyme digestion

Restriction enzyme plasmid digestion

Materials:

Plasmid
CutSmart buffer
dH2O
DNA inserts
Restriction enzymes
Water bath

Protocol:

Digestion of plasmid: Combine 50 uL plasmid, 2 uL of restriction enzyme (BamHI-HF), 2 uL of restriction enzyme (NcoI-HF), 8 uL of CutSmart buffer and 20 uL dH2O.Leave at 37°C (water bath) for 1 and a half hours.
Digestion of insert: 8 uL of H2O, 2 uL BamHI, 2 uL NcoI, 2 uL Cutsmart, 8 uL of insert DNA left at 37°C (water bath) for 1 and a half hours.

Double digestion of plasmid/ inserts

Materials:

Restriction enzymes (BamHi-HF, Ncol-HF)
CutSmart buffer
Plasmid
DNA inserts

Protocols:

BamHI and NcoI can be used together in digestion, make sure you use the right buffer (presumably CutSmart for the HF versions of these enzymes). Ensure you have quite a lot of plasmids at hand, e.g. Miniprep plasmid from 2x10ml overnight LB cultures using two separate spin columns, and check concentration using Nanodrop – preferably you would have more than 100ng/μl.
Inserts which you amplified by PCR need to be cleaned up first, e.g. by PCR purification kit.
To ensure each enzyme actually works prepare two separate single digests of the plasmid (you won’t see any difference with the inserts), assuming you have at least 100μl plasmid at hand prepare the following using either NcoI or BamHI:
5μl buffer + 44μl plasmid + 1μl enzyme (added last) = final volume of 50ul
Spin the reactions briefly to make sure everything is at the bottom of the tubes.
From each single digest take 44μl and transfer to a separate tube. Mix the two reactions which will give you now the double digest.
Incubate all three tubes (3 reactions per 1 construct) at 37 degC for a couple of hours; for the gene inserts I sometimes incubate for several hours or even overnight.
Digested plasmid should be gel-extracted; to ensure each enzyme cut properly run a few μl of each single digest and of the double digest on the same gel (using smaller wells) or before you attempt to gel extract. As long as you see just one vector band in each reaction at the appropriate size then it must have been successfully linearised and thus the enzymes work.
After gel extraction it is likely your vector will be at quite a low concentration (e.g. 10ng/μl or less) and the intercalating agents like Sybr Safe or ethidium bromide trailing from gel may interfere as well. If you want to be sure you can run the gel-extracted plasmid on the gel to check it is there.
Digested inserts can be purified by something like “PCR cleanup” – e.g. Qiagen have a kit where you mix the reaction with 5 volumes of buffer PB and you clean that on a spin column and then elute the purified DNA. You just want to remove cleaved bits, buffer and the enzymes, and you can as well gel extract the bands but you always lose more DNA by gel extraction.
Ellute the gel-extracted plasmid and purified linear DNA in the smallest volume possible to concentrate the DNA.

3.DNA ligation reaction

Materials:

Digested insert
Digested plasmid
T4 DNA ligase
Ligase buffer

Protocols:

You want to use an excess of the digested insert over the digested plasmid, usually people use 3-5-fold excess. Remember that your inserts are considerably shorter (say 500bp) than the plasmid backbone (say 5000bp) so if Nanodrop/agarose gel shows similar concentration of plasmid and insert, if fact you have 10x more insert.
As long as you know there is some plasmid from gel extraction and the purified inserts have some reasonable concentration in Nanodrop (above 20ng/μl for short, <500bp constructs, for longer constructs it would be expected even more) Set up a reaction like this, using NEB T4 DNA ligase: 2μl ligase buffer + 13μl digested insert + 4μl digested plasmid + 1μl ligase (added last).
After spinning the tube briefly, incubate the reaction at room temperature for a few hours or even overnight before transforming into DH5 alpha cells.

4. Colony PCR

Colony PCR is a method for determining the presence of insert DNA in a plasmid construct. This procedure was carried out in order to check the presence of a domesticated DNA part or transcriptional unit.

Materials:

250 µL tubes
Ice box
Thermocycler
Pipettes and pipette tips
Nuclease-Free dH2O
Standard Taq buffer (10X) NEB
Primers forward and reverse (10 uM)
dNTPs 2.5 mM
Taq polymerase enzyme - NEB
5 µl diluted colony

Protocol:

Colony PCR protocol from New England Biolabs (NEB). For each PCR reaction, primers annealing temperature was determined using the New England Biolabs online calculator tool (NEM TM calculator).

Colonies are picked and smirred inside the PCR tubes.
On ice, add all components in a 250 µL tube, making up to a 25 µl volume reaction. If checking the presence of a plasmid in many colonies, a Master Mix is recommended to be done by including all reagents except colonies.

Protein purification

Nickel column

Dialysis and reverse nickel purification

Materials:

Buffer A: 50mM Tris-HCl pH 7.5-8, 500mM NaCl
Buffer B: 50mM Tris-HCl pH 7.5-8, 500mM NaCl + 500mM imidazole
Dialysis buffer: 20mM Tris pH 7.5-8, 200mM NaCl

Protocols:

Prepare 5 litres of dialysis buffer (20mM Tris pH 7.5-8, 200mM NaCl)
Thaw out TEV: use about 1-2mg TEV per 50mg protein
Take a sample of the elution for gel and then add the appropriate amount of TEV
Set up a dialysis sausage with the elution+TEV (ensure you use the right cut-off, for proteins 20kDa or smaller use SnakeSkin with cut off 3 kDa!)
Leave the dialysis stirring slowly overnight in the cold room
Next day transfer the dialysate into a falcon tube and spin 15 min at max speed to pellet precipitate, then filter with 0.45um syringe filter (there will probably still be a lot of crap, TEV is notorious…)
Load the cut protein slowly onto the column – the slower the better, you want to remove TEV/HisTag/uncut protein; again you can discard the first 5 ml that go through
When all protein went through, pass another 10ml of dialysis buffer (or just diluted buffer A) through to column to ensure all the cut protein has gone out of the column
To remove TEV/HisTag/uncut protein from the column pass 3-4 column volumes of Buffer B ( 50mM Tris-HCl pH 7.5-8, 500mM NaCl + 500mM imidazole), collect that bulk elution in case cleavage was not efficient – you can dialyse that to remove imidazole, add more TEV and do the reverse nickel again!
To ensure the column is clean, pass some strip buffer (containing 50mM EDTA that chelates nickel ions) until the column is white, then a couple CVs of water, then the green nickel solution to recharge the column, and again some water. [If it looks dirty or you had huge amounts of crashing out protein on the column, you may want to wash it with a couple of CVs of 0.5M NaOH – ideally after stripping and water wash, before recharging with nickel.] Then the column can be stored in 20% EtOH in the fridge. Make sure the peristaltic pump tubing is washed with WATER before the tube is TAKEN OFF THE WHEEL – that makes the tube relax and less prone to cracking.

Large scale cell lysis and Ni purification

Materials:

Buffer A: 50 mM Tris-HCl (pH 7.7), 500 mM NaCl
Buffer B: 50mM Tris-HCl pH 7.5-8, 500mM NaCl + 500mM imidazole

Protocols:

Use 1 L overnight culture
Pellet at 7000 rpm, 4°C for 7 minutes
Take the pellet, add some (10-20ml) Buffer A.
In a small beaker with a stirrer dissolve 1 tablet of complete protease inhibitors (Roche) and add DNase (1 aliquot for pellet up to 2L, 2 aliquots if more)
Add the contents of the beaker to the Falcon tube with the pellet and swirl it around; you can use a spatula to scrape pellet off the sides
Transfer the buffer with chunks of pellet into the beaker, try to take all the pellet out
Keep stirring until pellet is properly resuspended (it can take 20 minutes or so, you can leave the beaker on the stirrer in the cold room for longer)
Switch the cell disruptor and the chiller on (ideally >10 min before you start lysis)
Pass 200ml water through the disruptor at >15 kpsi, then some Buffer A. [Stop the disruptor ideally when the liquid disappears from the cup but no air has yet entered the system]
Pour the resuspended pellet into cup VIA METAL SIEVE; if the solution looks very viscous or clumpy leave it stirring for longer and perhaps add some more DNase to it
Change pressure to 30 kpsi and pass the sample through, collecting it in a beaker
You can do a second round of lysis – stop the disruptor again when the liquid just went down, pour the lysate back in the cup and lyse again
At the end you can pass some (10-20ml) of buffer A to rinse out the remaining lysate
Reduce the pressure back to around 15 kpsi, fill the cup with water and pass it all through
Switch the CHILLER OFF
Pass 200ml 70% EtOH through the disruptor; the foam is tough to get rid of so you can use the EtOH in a squirty bottle to remove the foam from the sides
Remove the cup and the underlying little sieve piece and wash them in the sink to remove any crap, then put them back on
Remove any waste, refill water/EtOH bottles if necessary, log your usage in the book and switch the machine off (button on the side)
Transfer the lysate into cream round-bottom tubes with caps, balance them
Spin the tubes in the JA20 rotor, 20 min at 20 000 rpm, 4 deg
In the meantime equilibrate nickel column on the peristaltic pump: first 1-2 volumes of water, then a few column volumes of buffer A
Pour the supernatant from the cream tubes into a small Duran bottle, supplement it DROPWISE with imidazole, final concentration 20mM (calculate how much of 5M imidazole stock you need to add). If lysate looks dense either dilute it down with buffer or filter with 0.45um filter
If the lysate looks clear, load it onto the nickel column – preferably slowly as this enhances the binding. Collect the flow-through in a beaker
When all lysate is loaded, pass a few column volumes (20-30ml) of Buffer A containing 40-50mM imidazole, collect the wash
Elute in 3-4 column volumes of buffer A containing 250mM imidazole; when you start to elute you can discard the first 5ml that go through as the elution buffer wouldn’t have reached the bottom of the column by then anyway
After elution pass a few column volumes of Buffer B through the column, then at least 5 column volumes of Buffer A – the column can then be ready for the next day
Measure the absorbance of the elution using elution buffer as a blank to estimate the protein content

Protocol for small scale Ni pull-down

Materials:

Sample
1x PBS Buffer
0.1 mm glass beads
Ni- Nta resin
milliQ water
5 mM imidazole

Protocols:

Pellet cells by centrifugation for 20-30 mins at maximum speed.
Discard media.
Resuspend cells in 500uL 1x PBS buffer until homogeneous.
Lyse cells by bead beating, add a scoop or “a bit” of 0.1mm beads and vortex on the vortex genie (chemical hood) for 10-20 mins.
Centrifuge for 20-30 mins at 14000 rpm (4 degrees C) to separate soluble/insoluble proteins.
While centrifugation is happening, take 100uL of Ni-Nta resin slur and pipette it in the column that will be used for pull-down. Centrifuge for 1-3 minutes to remove residual EtOH from storage. Discard Flow through.
Wash the resin slur with at least 500uL milliq H2O. Centrifuge for 1-3 minutes. Discard Flow through
After centrifugation from step 6 is done, take the supernatant containing soluble proteins and add it to the equilibrated resin (KEEP 20 uL of protein before mixing to the Ni resin, this is your LOADED sample for the gel later). Pipette up and down to mix and then incubate then for 30mins-1h at 4 degrees in the spinning wheel to allow protein binding to Nickel resin.
After proteins are bound to Nickel resin, centrifuge for 1-3 minutes or until all liquid has passed through the filter. KEEP the flow through as this are the proteins that did not bind to Nickel. This is the FLOW THROUGH sample.
Add 500uL of PBS+5mM imidazole to the resin to wash residual unbound proteins. KEEP the flow through as this are the proteins that did not bind to Nickel strong enough and came out in the wash. This is the WASH sample.
Add 100uL of PBS+250mM imidazole to the resin to elute proteins bound to Nickel. KEEP the flow through as this potentially has your protein of interest. – ELUTED.
Run a gel: 3-5 uL of the LOADED and FLOW-THROUGH samples, 10-25uL of the WASH sample and 25uL of the ELUTED sample.

Protein analysis

Protein gel preparation and running (Amersham ECL™ gel system)

Electrophoresis

Protocol:

Prepare 1×running buffer by diluting 19ml of Amersham ECL Gel Running Buffer, 10× in 171 ml water.190ml buffer is sufficient for one electrophoresis gel.
Add 90ml of 1×running buffer to each tank of Amersham ECL Gel Box.
Cut open the gel package and gently remove Amersham ECLGel from the package.
Rinse the gel cassette with distilled water.Peel off the tapes from the two legs of the cassette.
Place Amersham ECL Gel in Amersham ECLGel Box so that the well side of the cassette faces toward the cathode(-) and the other cassette leg faces toward the anode(+).
Place the safety lid on top of Amersham ECL Gel Box.
Connect AmershamECL Gel Box to the power supply (EPS301) and pre-run the gel for 12 minutes at 160V.
Do not exceed the maximum operating voltage of 200V .
Note: Expected start current approx.45mA.
Once the pre-run is finished,switch off the power.
Remove the safety lid.
Wigglethecombbackandforth,andbringitstraightupfrom thecassettetomakethewellsavailableforsampleloading.
Note: Small gel pieces can be detached from the well container.This will not affect gel performance. Do not discard the comb.
Add 6ml of 1×running buffer to the well container.
Prepare the samples by adding sample and 2×sample buffer in a 1:1 mixture.
Note: For native conditions: use a sample buffer without SDS and DTT.
Heat the samples at 95°C for 5 minutes.
Spin down the samples quickly in a microcentrifuge and load the samples directly into the wells in the gel. Note: A maximum of 0.5μg /band per well of sample can be loaded. Overloading may cause smearing and distortion.
Note: To ensure uniform mobility ,load an equal volume of 1×SDS sample buffer into any unused well.
Place the safety lid on top of Amersham ECL Gel Box.

Parameter	Value
Voltage	160 V
Expected current	Start:approx.45mA End:23to33mA
Run time	60 minutes

Removing the gel from the cassette

Protocol:

Once the run is completed,shut off the power, disconnect the electrodes, remove the safety lid and finally, remove the gel cassette from Amersham ECL Gel Box.
Open the cassette by inserting the edge of the comb in the slot opposite the sample wells, and twist.
Remove the top plate from the gel cassette and allow the gel to sit on the bottom plate.
Cut the stacking gel with the end of the top plate approximately 2cm downstream of the wells. Repeat the procedure on the other side of the gel to remove the front.
Hold the gel cassette bottom plate over a container with suitable buffer, with the gel facing downwards. Gently push a tweezer between the gel and the cassette bottom plate until the gel is removed from the bottom plate.

Post-staining of the gel

250ml of each solution is used in each step.

Solution	Preparation
Fixing solution	Mix 400ml of ethanol and 100ml of acetic acid with distilled water. Make up to 1.0l with distilled water.
Staining solution	Mix 1 table to PhastGel™ BlueR-350 with destaining solution. Make up to 400ml with destaining solution.Heat to 60°C , stirring constantly, and filter before use.
Washing solution	Distilled water
Destaining solution	Mix 250 ml ethanol and 80 ml acetic acid. Make up to 1.0 l with distilled water.
Preserving solution	Mix 25ml of (87%v/v) glycerol with distilled water. Make up to 250 ml with destaining solution.

Fixation: Immediately after electrophoresis, immerse the gel in fixing solution for 30 minutes.This solution precipitates the proteins and allows the SDS to diffuse out of the gel.
Staining: Discard the fixing solution.Stain the gel for 10 minutes in staining solution. Cover the staining dish.
Washing: Rinse the gel once in distilled water.
Destaining: Destain the gel by changing the destaining solution several times until the stained protein bands are clearly visible against the clear background.
Preserving: Soak the destained gel in preserving solution for 30 minutes.

Western blotting

Protocol:

Run an SDS-Page gel with samples - any gel running system can be used.
Stack the gel between sponges, filter paper and a nitrocellulose membrane using the following order:
Negative electrode - sponge - filter paper - protein gel - nitrocellulose - filter paper - sponge - positive electrode
Run the transfer to the membrane at 90 V for 90 minutes using transfer buffer (Tris-glycin with 10% methanol)
Remove the membrane and block with milk powder dissolved in 50 mL PBS.A tween 20 for 1 hour
Pour off used blocking solution
Prepare 25 mL of fresh milk powder in PBS.A tween 20 for 1 hour and add 5 uL of anti-His antibody
Leave for shaking for 1 hour.
Wash the membrane with excess PBS A Tween 20 then at least 3x50 mL of PBS.A Tween 20 and leave for 5 minutes each wash
Spread 1 mL of ECL (the substrate solution for the horse-radish peroxidase) in order to visualize the blot
Use the ChemiDoc stain free blot setting to visualize the blot

Laboratory Notes

Our laboratory work was documented using Benchling and the laboratory notes are presented here.

1. Basic laboratory preparations: Made competent cells, agar plates and plasmids that can be digested and used for cloning.

2. Preparing cells containing the correct plasmids for antibody expression: Cloned in the antibody fragment DNA of all our wild-types and mutant antibody CH3 domains into pEHISTEV expression vector. Transfomed SoluBL21 and Origami cells with expression plasmids.

3. Expressing and purifying antibody fragments: Expressed antibody fragments in SoluBL21 and Origami cells, purified these proteins via nickel affinity chromatography and reverse nickel chromatography. Analysed expressed proteins via mass-spectrometry

4. Expressing a commercially available antibody fragment as an antibody expression test in E. coli.:

5. Expressing the Bal-3 isopeptide domain, as a positive control for bacterial isopeptide domain expression:

6. Expressing wild-type and mutant Thermotoga maritima enzyme mutants:

7. Characterizing the mOrange-SpyTag conjugate.:

8. Neomycin resistance characterization in DH5Alpha cells:

9. Characterizing T7 promoter using in vitro transcription: