Difference between revisions of "Team:TUDelft/Wetlab/Protocols"

This protocol is based on the Pierce BCA protein assay kit by Thermo Scientific protocol.

1. Prepare a set of protein standards using one 2mg/mL Albumin Standard (BSA) ampule according to the table below:
   NOTE: Use the same diluent as the samples. The expected working range = 20-2000µg/mL.
   

Vial	Volume of MilliQ (µL)	Source of BSA	Volume of source BSA (µL)	Final BSA concentration (µg/µL)
A	0	Stock	300	2000
B	125	Stock	375	1500
C	325	Stock	325	1000
D	175	Vial B	175	750
E	325	Vial C	325	500
F	325	Vial E	325	250
G	325	Vial F/td>	325	125
H	400	Vial G	100	25
I	400	n/a/td>	0	0

2. Determine the amount of total volume of working reagent (WR) required by using the the following formula:
   Total volume WR = (# standards + # unknowns) × (# replicates) × (200 µl)
3. Prepare the BCA working reagent by mixing 50 parts of BCA Reagent A with 1 part of BCA Reagent B (50:1, Reagent A:B). NOTE: The WR is stable for several days when stored in a closed container at room temperature (RT).
4. Pipette 25µL of each standard or unknown sample replicate into a microplate well.
5. Add 200µL of the WR to each well and mix plate thoroughly.
6. Cover plate and incubate at 37°C for 30 minutes.
7. Cool plate to room temperature.
8. Measure the absorbance at or near 562nm on a plate reader.

NOTE: All work is performed within a sterile field created by a bunsen burner flame.

1. For one cryostock, take a 1.5mL sample from an overnight liquid cultures.
2. Centrifuge the 2mL tubes at 2000rpm for 10 min.
3. Decant the supernatant without disturbing the pellet.
4. Add fresh sterile LB medium to the pellet, 1/3 volume of the starting volume of the culture.
5. Completely resuspended the pellet by vortexing the tube.
6. Add sterile 80% glycerol solution, the same volume as fresh LB in step 4.
7. Mix by vortexing.
8. Make a 1mL aliquot in cryotubes and label it with the cell type, plasmid type, protein type, operator and date.
9. Store the vials at -80ºC and update the inventory.

This protocol spans over 3 days of execution time, starting from a -80°C mother stock. Throughout the protocol, it is recommended to work under aseptic conditions in order to prevent contamination risks.

Day 1



1. Keep the -80°C strain stock of interest on ice.
2. Streak the strain on solid selective medium and incubate overnight at 37 °C while shaking.



Day 2



3. Prepare a 10mL liquid starter culture with one of the colonies that grew on the selective plate. Let the culture grow overnight at 37°C, shaking at 180rpm.
4. Sterilise solutions of CaCl₂ 100 mM and of CaCl₂ 100 mM + 15 % Glycerol in advance.
   NOTE: Volumes depend on the total culture volume to be prepared in step 5. For example, if 1 L is used in step 7, 300 mL of CaCl₂ 100 mM and 10 mL of CaCl₂ 100 mM + 15 % glycerol will be used.



Day 3



5. Inoculate 1:100 of overnight culture in the desired volume of LB with antibiotic (eg. 10 mL of culture in 1 L of LB).
6. Incubate in a shaker at 37 °C, 180 rpm to OD600nm ~0.4-0.6 (measure OD600nm every 30 minutes).
7. Harvests the cells by centrifugation at 4000 x g for 5 minutes in centrifuge tubes, decant supernatant.
8. Resuspend cells by gently pipetting 1/5 (of the volume of LB from step 5) of ice-cold 100 mM CaCl₂ and incubate on ice for 20 minutes.
9. Pellet the cells by centrifugation at 4000 x g for 5 minutes in centrifuge tubes, decant supernatant.
10. Resuspend cells by gently pipetting 1/10 (of the volume of LB from step 5) of ice-cold 100 mM CaCl₂ and incubate on ice for 60 minutes.
11. Pellet the cells by centrifugation at 4000 x g for 5 minutes in centrifuge tubes, decant supernatant.
12. Resuspend cells by gently pipetting 1/100 (of the volume of LB from step 5) of ice-cold 100 mM CaCl₂ + 15% glycerol and keep on ice.
13. Chemical competent cells can either immediately be used for heat shock transformation , or stored in aliquots of 50 uL in microcentrifuge tubes at -80 °C.

1. Under aseptic conditions, pick a colony, resuspend it in 10 µL of milli-Q water.
   NOTE: A picked colony cannot be used again; it is recommended to restreak on a 'back-up'-plate and incubate it overnight at 37 °C.
2. Incubate the resuspended colony at 90 °C for 10 min. Spin the suspension down and use the supernatant as template DNA for the PCR.
   NOTE: Instead of separate boiling prior to PCR, this step can be incorporated in the PCR program. The initial denaturation step at 98 °C should then be prolonged to 5 minutes.
3. Make sure every PCR reaction is composed as follows:
   

   Component	Volume (µL)	Final concentration
   GoTaq 5x buffer*	10	1 X
   10 mM dNTPs	1	200 µM
   Primer forward (10µM)	1	200 nM
   Primer reverse (10µM)	1	200 nM
   Boiled colony supernatant	5
   Gotaq polymerase (5u/µL)	0.2	20 U/mL
   MilliQ	31.8

   * NOTE: Use GoTaq Buffer Green when it is required to run a verification gel afterwards.
4. Add 5 µL of supernatant of colony mixture to each PCR tube.
5. Close all tubes thoroughly and place them in a thermocycler with the following protocol:
   

   Step	Time (s)	Temperature (°C)
   Initial denaturation	150	98
   Denaturation	60	94
   Annealing	60	Tann*
   Extension	60 sec per kb DNA	72
   Final extension	600	72
   Hold	∞	4

   *NOTE: The annealing temperature (T_ann) is dependent on the melting temperature (T_m) of the primers used. It is recommended to have T_ann = T_m - 5°C.
6. The PCR product(s) can be checked on gel. In order to do so, cast a gel and prepare the samples according to the DNA electrophoresis protocol.

Gel Electrophoresis and DNA staining makes use of mutagenic chemicals like EtBr, SYBR Safe or any other DNA staining. Wear protection (gloves) when carrying out this protocol and work in an assigned area for this work to prevent contamination of the rest of the lab.

1. Weigh agarose powder for a 0.8% (w/v) gel in TAE.
   E.g. weigh 1.6g and add to 200mL TAE buffer for a 0.8% gel.
2. Add weighed agarose in TAE buffer (1X) and warm the solution (in a microwave) until it is completely dissolved. Let the solution cool down to hand warm temperature.
   NOTE: Make sure the lid is not completely closed to avoid possible exploding of the glass bottle.
3. Pour gel in gel tray and mix well with SYBR Safe. For a small gel (~ 40 mL solution) add 1 µL of SYBR Safe; for a large gel (~80 mL) add 2 µL of SYBR Safe. Add a comb to create wells for the samples. Allow the agarose to solidify (approximately 20 minutes).
4. Transfer the gel to the electrophoresis cell, remove the combs and cover the gel in TAE buffer (1X).
   NOTE: Mind the direction of DNA migration when placing the gel in the cell.
5. Prepare the electrophoresis samples by adding Nucleic Acid Loading Buffer conform the manufacturer’s instructions.
6. Load the molecular weight marker (DNA ladder) in the first well according to manufacturer’s instructions (generally 3-5 µL) and load 5-10 µL of the dyed samples in the other wells.
   NOTE: Do not contaminate the loading buffer and ladder with SYBR Safe! Do not touch it while wearing a glove.
7. Connect the cables of the gel tray following the colour code and run at 80-110V for 40-60 min.
   NOTE: NOTE: Time and voltage depend on the density of the gel and the length of the lane in the gel.

This protocol is based on the Wizard® SV Gel and PCR Clean-Up System of Promega Corporation.

1. Place the excised gel slice in a 1.5 mL Eppendorf tube.
2. Add 10 µL Membrane Binding Solution per 10 mg of gel slice. Vortex and incubate at 50-65 °C until gel slice is completely dissolved.
3. Insert the SV Minicolumn into Collection Tube and label both of them according to the labelling of your samples.
4. Transfer the dissolved gel mixture to the Minicolumn assembly. Incubate the Minicolumn at room temperature for 1 minute.
5. Centrifuge the SV Minicolumn at maximum speed for 1 minute.
6. Discard the flow through and reinsert the SV Minicolumn into the Collection Tube.
7. Add 700µL of Membrane Wash Solution.
   NOTE: Upon prior use, dilute the solution with 95% ethanol following the manufacturer's’ instructions.
8. Centrifuge the SV Minicolumn assembly at maximum speed for 1 minute.
9. Discard the flow through and reinsert the SV Minicolumn into the Collection Tube.
10. Repeat the washing step with 500µL of Membrane Wash Solution and centrifuge for 5 minutes at maximum speed.
11. Once the Collection Tube is empty, centrifuge the Minicolumn assembly at maximum speed for 1 minute with the microcentrifuge lid open to allow ethanol full evaporation.
    NOTE: Leaving the column at room temperature ameliorates evaporation of residual ethanol traces.
12. Transfer the SV Minicolumn to a clean labelled 1.5 mL Eppendorf tube.
13. Add 50 µL of pre-warmed MilliQ directly to the centre of the SV Minicolumn, without touching the membrane with the pipette tip.
    NOTE: Use 30 µL when higher final concentrations of DNA are required or when small quantities of DNA are suspected.
14. Incubate the SV Minicolumn at room temperature for 5 minutes.
15. Centrifuge at maximum speed for 1 minute.
16. Discard the SV Minicolumn, cap the tube containing the eluted DNA and keep the DNA at 4 °C (for immediate use) or -20 °C (for storage).

Our DpnI digestions were performed with New England Biolabs DpnI (20.000 units/mL), which is compatible with CutSmart Buffer by the same manufacturer.

1. Prepare a sample in a 0.5 mL microcentrifuge tube as follows:
   

   Component	Volume (µL)
   10x CutSmart buffer (NEB)	4
   Purified PCR product	30
   Restriction Enzyme DpnI	1
   MilliQ	5

   Incubate for 1.5 hours at 37°C.
2. Heat inactivate the enzyme by incubating at 80oC for 20 minutes.

This protocol spans over 3 days of execution time, starting from a -80 °C mother stock. Throughout the protocol, it is recommended to work under aseptic conditions in order to prevent contamination risks.

Day 1



1. Sterilize the required 50 mL MilliQ and 250 µL glycerol in advance.
2. Keep the -80 °C strain stock of interest on ice.
3. Streak the strain on solid selective medium and incubate overnight at 37 °C.



Day 2



4. Prepare a 10mL liquid starter culture with one of the colonies that grew on the selective plate. Let the culture grow overnight at 37 °C, shaking at 180rpm.

Day 3



5. Use 0.5mL of the starter culture to inoculate 35mL selective liquid medium and keep track of the OD660.
6. Grow at 37°C while shaking 250 rpm till an OD660 of ~0.5.
7. Centrifuge for 10 minutes at 4 °C at 3900 rpm.
8. Discard the supernatant and resuspend pellet in 20 mL cold Milli-Q.
9. Centrifuge for 10 minutes at 4 °C at 3900 rpm.
10. Discard the supernatant and resuspend pellet in 20 mL cold Milli-Q.
11. Centrifuge for 10 minutes at 4 °C at 3900 rpm.
12. Discard supernatant and resuspend in 200 µL 50% glycerol.
13. Prepare aliquots of 50 µL.
14. Either transform the cells straight away or store the electrocompetent cells at -80 °C.

NOTE: This protocol allows for a single transformation.

1. Thaw a 50 µL aliquot of electrocompetent cells on ice.
2. Add ~200 ng DNA to the cells and keep on ice for 20 minutes.
   NOTE: in case of purified plasmid DNA, 50 ng DNA is enough.
3. Transfer all the content to an electro-shock cuvette.
4. Electro-shock the cells with the Electro Cell Manipulator at 2.5 kV.
5. Immediately add 0.2-1 mL of recovery medium (eg. SOC-medium). Resuspend and transfer to a 1.5 mL tube.
6. Incubate at 37 °C at 250 rpm for 1 hour.
7. Plate the cells on solid medium with appropriate antibiotics and incubate overnight at 37 °C.

This protocol is based on the instructions for G-Block resuspension given by Integrated DNA Technologies (IDT).

1. Centrifuge the tube containing the gBlock for 3−5 seconds (>3,000 x g) to pellet the material to the bottom of the tube.
2. Add an appropriate volume of sterile Milli-Q to the tube for a desired final concentration. The required volume of Milli-Q can be read from the table below (and the label on the IDT tube that contains your fragment):
   

   Final concentration (ng/µL)	µL MilliQ to add to 250 ng	µL MilliQ to add to 250 ng	µL MilliQ to add to 250 ng
   10	25	50	100
   20	not recommended	25	50
   50	not recommended	10	20

3. Incubate at 50°C for 20 minutes.
4. Briefly vortex and centrifuge.
5. Store the resuspended gBlock at -20 °C.

This Gibson Assembly protocol is based on the protocol provided by New England Biolabs.

1. Thaw 10 µL of 2x Gibson Assembly mastermix (New England Biolabs) on ice.
2. Add backbone and insert; depending on the assembly, assembly pieces can be added in a predetermined ratio (recommended is ratio 1:3 mol vector over mol insert, so not a 1:3 ratio based on weight). Do not exceed the total volume of 10µL.
   NOTE: An online ligation calculator can subsequently be used to calculate the amount of the assembly pieces that is required.
3. If applicable, fill up the reaction volume to 20µL with Milli-Q.
   

   Component	Volume (µL)
   10x Gibson Assembly Master Mix 2X (NEB)	10
   Vector	X (? ng)
   Insert fragment	Y (? ng)
   MilliQ	10-X-Y

4. Incubate the assembly reaction at 50 °C for 60 minutes and place on ice for subsequent transformation. Otherwise, store at -20°C.

1. For each PCR in 50µL reaction volume, make sure the composition is as follows:
   

   Component	Volume (µL)	Final concentration
   5x Phusion HF buffer	10	1 X
   10 mM dNTPs	1	200 µM
   Primer forward (10µM)	2.5	200 nM
   Primer reverse (10µM)	2.5	200 nM
   DNA template	~10 ng to 250 ng*
   Phusion polymerase	0.5	20 U/mL
   MilliQ	up to 50 µL

   
   * NOTE: The 10 ng - 250 ng of template DNA are approximate, choose a volume that works fine for all your samples.
   NOTE: High fidelity PCR with Phusion polymerase can be optimized per case by adding 3% of DMSO.
2. Close all tubes thoroughly and place them in a thermocycler with the following protocol:
   

   Step	Time (s)	Temperature (°C)
   Initial denaturation	30	98
   Denaturation	10	98
   Annealing	15	Tann*
   Extension	15-30 sec per kb DNA	72
   Final extension	300	72
   Hold	∞	4

   *NOTE: The annealing temperature (T_ann) is dependent on the melting temperature (T_m) of the primers used. It is recommended to have T_ann = T_m - 5°C.
3. The PCR product(s) can be checked on gel. In order to do so, cast a gel and prepare the samples according to the DNA electrophoresis protocol.

1. Decide on which enzyme(s) to cut with. Check online what buffer the enzyme(s) work(s) in (NEB). For most of the enzymes, the SmartCut buffer 10X can be used.
2. Prepare a sample a sample as follows:

Component	Volume (µL)
10x CutSmart buffer (NEB)	2
Fragment (~1-2 μg)	X (depending on the concentration)
Restriction Enzyme 1	1
Restriction Enzyme 2 (optional)	1
MilliQ	20 - (3 + X) add up to 20 µL

3. Incubate for 4 hours at 37 °C.
4. Inactivate the restriction enzyme(s) by heating to 65 °C for 10 minutes.
   NOTE: This last step can be skipped if the sample is evaluated on gel electrophoresis immediately. Also, some enzymes are not inactivated by increasing the temperature to 65 °C. It is preferred to perform DNA Clean Up according to the DNA Purification (PCR) protocol for subsequent cloning strategies.