Difference between revisions of "Team:UCL/Protocols"

Restriction digestion

1. Prepare restriction digestion mixture:
   IDT gBlocks
   - 10 ul of DNA (10 ng/ul)
   - 2 ul of 10 x 2.1 buffer
   - 0.3 ul of EcoR1
   - 0.3 ul of Pst1
   - 7.4 ul of milliQ H2O
   
   
   Other digestions
   - Required amount of DNA
   - 1 ul of 10 x 2.1 buffer
   - 0.3 ul of EcoR1
   - 0.3 ul of Pst1
   - add milliQ H2O up to 10 ul
   (if using total volume greater than 10ul, increase the amount of buffer accordingly)
   
   
2. Incubate at 37C for an hour
3. Heat inactivate by incubating at 80C for 20 minutes
4. Run a sample of digested DNA on a gel in order to confirm digestion:
   - 2 ul of DNA
   - 1 ul of 6 x Gel Loading Dye
   - 3 ul of milliQ H2O
5. If digestion is confirmed, proceed to ligation

Ligation

1. Calculate the amount of insert DNA required to maintain 1:3 backbone:insert molar ratio using formula below. For standard ligations use 50 ng of vector DNA, increase the amounts of DNA if unsuccessful.
   
$$Insert\:Mass\:in\:ng = 3\times \bigg[\frac{Insert\:Length\:in\:bp}{Vector\:Length\:in\:bp}\bigg] \times Vector\:Mass\:in\:ng$$
2. Prepare the ligation mixture:
   - required amount of insert DNA
   - required amount of vector DNA
   - 1 ul of T4 ligase
   - 2 ul of 10 x ligase buffer
   - add milliQ H2O up to 20 ul
3. Incubate at 16C for 30 minutes
4. Heat inactivate by incubating at 80C for 20 minutes
5. Keep on ice until ready to proceed with transformation protocol

Transformation

1. (If using part from the distribution: resuspend the DNA in 10 ul of MiliQ water, making sure that it turns red. Wait 10 minutes before adding the DNA to cells)
2. Put a tube of NEB DH 5 alpha E. coli cells on ice and wait until they thaw completely. Divide the cells into 50 ul aliquotes.
3. Add 1 ul of plasmid DNA to 50 ul of cells.
4. Mix by carefully flicking the tube. Do not vortex or pipette in and out!
5. Place the mixture on ice for 30 minutes.
6. Heat shock the cells at 42 °C for 30 seconds and immediately put on back on ice.
7. Keep cells on ice for next 5 minutes. Do not mix.
8. Pipette 950 ul of SOC media kept at room temperature into the mixture. If SOC is not available, use LB.
   
9. Incubate the mixture at 37 °C for 60 minutes
10. Prepare plates with appropriate antibiotics. Bring plates to room temperature before plating. Use 2 plates per transformation reaction.
11. Plate 200 ul of cells on one plate.
12. Pellet the remaining cells and resuspend in 200ul of LB.
13. Plate the remaining cells on second plate.
14. Incubate plates overnight at 37 °C.

Agarose gel electrophoresis

1. Measure 0.50 g of agarose
2. Measure 50 ml of 1x TAE buffer using measuring cylinder
3. Add agarose and TAE buffer to conical flask and gently mix
4. Microwave the flask for 1 min
5. Wait for the mixture to cool down slightly before proceeding
6. Add 10 ul of 10mg/ml ethidium bromide solution and mix
7. Assemble the casting tray and pour the gel into it
8. Wait around 30 minutes until gel gets solidified
9. Put the gel into the gel chamber and pour 1x TAE buffer until it is fully covered
10. Load 6 ul of DNA ladder to the first well.
11. Prepare the samples by adding appropriate volume of 6x gel loading dye and load them
12. Assemble the gel chamber and run the gel for 40 minutes at 120V
13. Visualise the gel using the gel visualizer

Assembly of 2 parts using gel extraction

1. Digest at least 500 ng of each part according to the restriction digestion
2. Run the digested DNA on the gel according to gel electrophoresis protocol
3. Identify the parts that you want to ligate on a gel and cut the bands out using razor blade
4. Purify the excised bands using the commercial kit according to the manufacturer's instructions
5. Quantify the DNA yield using DNA nanodrop
6. Proceed to ligation

Polymerase Chain Reaction

1. Prepare the PCR mix:
   - 12.5 ul of 2 x Q5 PCR master mix
   - 1.25 ul of 10 uM forward primer
   - 1.25 ul of 10 uM reverse primer
   - 2 ng of DNA to be PCRed
   - add milliQ H2O up to 25
2. Set up the PCR cycles according to the following rules:
   Initial denaturation
   - 98C for 30 seconds
   35 cycles
   - 98C for 10 seconds
   - 30 seconds at primer melting temperature
   - 72C for 30sec/kb of PCRed fragment
   Final extension
   - 72C for 2 minutes
   - Hold at 4C
3. Confirm the PCR by running 2 ul of the product on the gel according to the gel electrophoresis protocol

Gibson Assembly

1. When designing the gBlock fragments for Gibson Assembly, make sure that the fragments have ~20 bp overlap and that first and last insert fragment have ~20 bp overlap with respective ends of PSB1C3
2. Convert the concentration of vector and inserts from ng/ul to pmol/ul using the following formula:
   
3. Prepare the Gibson Assembly mixture:
   - 0.08 pmol of each insert
   - 0.04 pmol of vector
   - 10 ul of Gibson Assembly mix
   - add milliQ H2O up to 20 ul
4. Incubate the reaction at 50C for 15 minutes. Following incubation, put samples on ice.
5. Proceed to transformation protocol. Use 2 ul of the Gibson Assembly reaction mixture for transformation

Agar plate preparation

IPTG-induced protein expression

1. On the afternoon before the induction, start seed culture with appropriate antibiotic from glycerol stock and leave to incubate overnight at 37C shaking
2. The next morning, use 2 µl of seed culture to inoculate 100 ml of fresh media with appropriate antibiotic in shaker flask and grow until an OD600 nm of 0.4-0.6 is reached
3. If necessary, prepare these in the meantime :
   - 50 ml stock of lysis buffer (25 mM TRIS-Cl, 2 mM EDTA, pH 7.6):
   • weight 0.151g of Tris base
   • add 45 ml of water
   • titrate with HCL to pH 7.6
   • fill up to 50 ml with water
   • add 29.2 mg of EDTA
   - IPTG 100 mM stocks: (23.8 mg IPTG per 1ml ddH2O )
4. When culture reaches OD of 0.4-0.6, add IPTG to a final concentration of 1 mM (=100ul of 100 mM stock)
5. Incubate induced culture at 30 °C for 4 hours
6. Split the culture into 4 falcon tubes (~25 ml each) and harvest the pellets by centrifugation for 20mins at max speed
7. Resuspend each pellet in 2 ml of lysis buffer, lyse by sonication (10 cycles of 10 sec with 10 sec breaks)
8. Centrifuge 20 min at max speed.
9. Transfer all supernatants to separate tube
10. Measure the concentration of protein in supernatant using Bradford Assay
11. Store in the freezer

Spectrophotometric assays

- Assay for tryptophan hydroxylase (BBa_K1598002) activity: download protocol

Golden Gate Assembly

DNA Assembly

Components	Amount (μl)
Sap1(15 U)	0,75
T4 Ligase (400 U)	1
DTT (10 mM)	1
ATP (10 mM)	1
G-Buffer (10x, Fermentas)	1
parts	40 fmoles each
ddH2O	Fill up to 10
Total Volume	10

Thermocycler programme:

1. 37°C, 5 min
2. 20°C, 5 min
3. repeat (1. and 2.) 50 times
4. 50°C, 10 min
5. 80°C, 10 min

Gut on Chip

Fabrication of the upper and lower microchannel

1. Pour 3g of 15:1 (wt:wt) PDMS mixture onto the bottom of the petri dish and spread across evenly.
2. Place the master on the PDMS with SU-8 facing up and wait 10mins on an even surface.
3. Pour 15:1 (wt/wt) PDMS mixture onto the silicon master and degas it in a vacuum desiccator for 30min. use 15g and 3g of PDMS for the fabrication of the upper and lower microchannel slabs.
4. Fully cure PDMS in an oven maintained at 60c for at least 4 hours
5. Peel the cured PDMS off the master, and cut it into a 2-cm wide x 3-cm long rectangular block using a scalpel.
6. Punch holes through the upper channel layer by using a biopsy punch with a diameter of 2mm.

Fabrication of porous membranes

1. Pour degassed 15:1 (wt/wt) PDMS mixture into an empty petri dish to gener4ate a 1-cm-thick flat PDMS slab.
3. Cure PDMS in a levelled dry oven at 60c for 4 h
4. Use a scalpel to cut the fully cured PDMS along the edge of the petri dish, and remove it from the dish by using tweezers.
5. Rinse the removed PDMS slab with 100% ethanol, and dry it completely using compressed nitrogen or air
6. Treat the PDMS slab with oxygen plasma for 1.5 min, and salinize it in a vacuum desiccator overnight
7. Place a silicon master containing an array of micro fabricated circular pillars (10µm in diameter and 30µm In height) with a centre to centre facing of 25µm at the centre of the bottom surface of a clean petri dish
8. Pour 3g of degassed 15:1 (wt/wt) PDMS mixture into the wafer, spread it evenly. Avoid the formation of bubbles.
9. Gently put a salinized PDMS slab from step 5 on the surface of the silicon master covered with uncured PDMS. Release the slab very slowly to prevent the formation of air bubbles
10. Place a frosted glass slide on the salinized PDMS block, and place 3 kg weight on the glass slide.
11. Wait for 30 min to allow for intimate contact between the PDMS slab and micro fabricated master surface. Perform this step on a level surface.
12. Move to the entire assembly to a dry oven at 60c and incubate it overnight.
13. Remove the sample from the oven, take off the weight, and cool the assembly to room temperature over 30 min
14. Use a scalpel to lift up a corner of the slab, and slowly peel it from the wafer. Apply 100% ethanol to the gap between the PDMS surface and the silicon wafer during this step to facilitate the detachment of the PDMS layer.

Alignment and Assembly of the micro device

1. Clean the upper PDMS layer and the porous membrane with packaging tape. Do not apply excessive pressure when you are cleaning the membrane surface, in order to avoid unwanted damage of the device and delamination.
2. Treat the membrane surface and the channel of the upper PDMS layer with the plasma by using a corona generator for 3s and 1min, respectively.
3. Use sweeping motions to achieve uniform treatment, and keep the tip of the electrode of the corona generator 5 mm away from the sample surface for best results.
4. Overlay the upper microchannel layer on the PDMS membrane, and bring them in contact. Press the PDMD slabs to permit intimate contact between layers and remove trapped air.
5. Incubate the assembled layers in the dry oven at 80 co. for at least 12h
6. Remove the sample from the oven, and cool it down at room temperature for 1h
7. Cut along the edges of the upper PDMS channel layer bonded to the membrane by using a scalpel, and gently peel the salinized flat PDMS from the assembly.
8. Put a few drops of 100 % ethanol between the layers for easier detachment.
9. Tear off the portions of a porous membrane located over the lateral vacuum chambers using fine tip tweezers under a stereoscope.
10. Expose the membrane surface and the channel side of the lower PDMS layer to the corona under a stereoscope.
11. Align and bond the device. Attempt to pull apart the upper and lower PDMS slabs to qualitatively determine the success of device bonding. Incomplete or unsuccessful bonding results in peeling off and separation of the PDMS slabs
12. Bend the tips of six 18 – gauge blunt needles by 90 co. by using pliers, and break off the needle at a point near the syringe entry port.
13. Insert the needles into the access ports of the central cell culture channels and side vacuum chambers.
14. Cut six pieces of 4 – cm long silicone tubing, and connect them to the free ends of the needles