Team:MIT/Recombinase Group Notebook Week 7

• BACKGROUND
• EXPERIMENTS
  Synthetic Promoters microRNA Recombinase Proof of Concept Collaborations
• FUTURE WORK
  Circuit Design Clinical Applications
• HUMAN PRACTICES
  Parts Toolbox Accessible Protocols Integrated Human Practices Public Outreach Interlab Study
• NOTEBOOK
  Lab Notebook Parts Mammalian Parts Collection Attributions Safety
• AWARDS
  HP Silver HP Gold Integrated HP Basic Part
• TEAM

TN 07/18/16 Circuit design meeting

Made with Benchling

Project: iGem 2016 _ Recombinase subgroup

Authors: Trinh Nguyen

Date: 2016-07-18

Monday, 7/18

sensitivity in micro level range of the progesterone degradation tag

Problems:

Contact George Church's lab for more sensitive progesterone degradation tag

Contact other labs that are expert in this field

Hybrid promoter: alternative between estrogen and progesterone response elements

Problems:

Loose sensitivity of both inputts

Recepter group:

Do in parallel both just estrogen response promoter and progesterone promoter

Then, construct the hybrid

Clinical Story:

If the circuit works, how do we do ex-vivo diagnostic: (biopsy)

Eutopic endometrium samples

Transfect the samples with the circuit

Induce with estrogen & progesterone

Detect dz traits and look for biomarkers/fluorescent

TN 7/18/2016

Made with Benchling

Project: iGem 2016 _ Recombinase subgroup

Authors: Trinh Nguyen

Date: 2016-07-18

Monday, 7/18

Resuspending oligos:

Q4 attP Qx: 4nmol => 40uL

Q4 attP Qx RC: 4nmol => 40uL

Q1 attB Q3: 4nmol => 40uL

Q1 attB Q3 RC: 4nmol => 40uL

Resuspending gBlocks:

Q1 TP901 Qx: 1068 fmol => 21.4 ul (final concentration 50fmol/uL)

Miniprep Concentrations:

	A	B
1	1-1	635.4
2	1-2	695.9
3	1-3	585.3
4	1-4	575
5	2-1	536.3
6	2-2	584.4
7	2-3	655.5
8	2-4	486
9	3-1	628.1
10	3-2	259.4
11	3-3	676.5
12	3-4	595.9
13	4-1	696.1
14	4-2	663.2
15	4-3	731.7
16	4-4	616.2

Table1

Restriction digest

1-1, 1-2, 1-3, 1-4: AflII

2-1, 2-2, 2-3, 2-4: AflII

3-1, 3-2, 3-3, 3-4: NcoI

3-1 DD, 3-2 DD, 3-3 DD, 3-4 DD: NcoI + BglII

4-1, 4-2, 4-3, 4-4: NcoI

4-1 DD, 4-2 DD, 4-3 DD, 4-4 DD: NcoI + XbaI

BB 1: AflII + pDEST

BB 2: NcoI + BglII + pDEST

BB 3: NcoI + XbaI + pDEST

BB 4: XbaI + pDEST

PCR

Introduction

PCR (the polymerase chain reaction) uses a polymerase and DNA oligonucleotides to amplify a length of double-stranded DNA. For general purpose PCR, I like to use the Q5 polymerase from NEB: the product documentation is here.

Materials

• 200 µl thin-wall PCR strip tubes, one per reaction
• Forward and reverse primers, 10 µM working stock
• Template DNA, 1 ng/ul
  • Make SURE that you have diluted out the template DNA appropriately! Miniprep-concentration template will screw up your PCR!
• Q5 master mix, thawed on ice
• Ice bucket with ice
• Empty 10 µl or 200 µl tip box
• Gel for gel electrophoresis
• Parafilm
• 6X NEB Purple Loading Dye

Procedure

• Set up your PCR program

1. Use tmcalculator.neb.com to compute the annealing temperature Ta of your reaction based on your primers' melting temperatures.

• Make sure to select Q5 2X Master Mix under "Polymerase/Kit"

2. Compute the extension time Tex based on your predicted product length: 30 seconds per kilobase. So if my amplicon is 1500 bp, my extension time is 1'30"

3. Program the thermocycler with the following (there should be a "Q5" program you can easily edit):

	A
1	Heated lid --> 105°C
2	HOLD at 98°C
3	98° for 30"
4	Cycle 30 times:
5	-- 98° for 5 seconds
6	-- Ta for 15 seconds
7	-- 72° for time Tex
8	Close cycle
9	72° for 2'
10	Store at 8°C

Table1

4. Start the program on the thermocycler. It will heat the lid, heat the block to 98° then hold at that temperature.

• Set up the PCR reactions

5. Use the following table to fill in your template and primers for each reaction you're running. For a 20 µl reaction, use 1 ul of forward primer (10 uM), 1 ul of reverse primer (10 uM), 1 ul of template (1 ng/ul), 7 ul nuclease-free water, and 10 ul 2X Q5 master mix.

	A	B	C	D
1		Q1_PhlF_QX
2	Fwd Primer	1 ul BT28 PhlF-F
3	Rev Primer	1 ul BT29 PhlF-R
4	Template	1 ul pENTR L1_PhlF_L2
5	Water	7 ul
6	2X Q5 MM	10 ul

Table2

6. Take the grey platform off of the tip box. Fill the top box with ice, then replace the grey platform. Fill the box with water until the level is just below the grey platform.

7. Label the PCR tubes.

8. Set up the PCR reactions. Start with the water; end with the Q5 master mix.

9. Working quickly, re-cap the PCR reactions. Flick several times to mix, then pulse spin in the tube microfuge. Return the tubes to ice.

10. Carry the tubes to the pre-heated thermocycler. Put the tubes in the block, close the lid and then press the button to continue through the hold and run the rest of the program.

• Analyze the PCR reaction with gel electrophoresis

11. Prepare or secure a gel.

12. Slice a small piece of parafilm off of the roll.

13. Spot 4 ul of water in a droplet on the parafilm. Add 1 ul of the PCR reaction and 1 ul of loading dye.

14. Pipette up and down with a 5 ul pipettor, then load in a well on the gel.

15. Run the gel.

16. If there is a single clean band, proceed to clean the reaction up with a QIAgen PCR cleanup column.

17. If there are multiple bands but you see the one you want, proceed to a gel extraction of the band you want.

Golden Gate Assembly

Introduction

Golden Gate is an assembly method using Type II-S restriction enzymes and predefined overlap sequences to assemble multiple pieces of DNA in one reaction. Golden Gate design is beyond the scope of this protocol; this protocol is the reaction setup only.

Materials

• 200 µl thin-wall PCR strip tubes, one per reaction
• Backbone plasmid
  • Usually pDONR-GG L1_lacZ_L2 or pDONR-GG L4_lacZ_R1
• Parts being assembled, either in plasmids or gBlocks
• Nuclease-free water
• T4 ligase buffer, 1.5 µl per reaction
  • Found in single-use aliquots of 10 (??) µl in the -20° freezer. T4 ligase buffer does NOT like to be thawed and frozen again; once you use an aliquot, discard what's left!
• 10X BSA, 1.5 µl per reaction
  • Found in the -20° freezer. ...and that's Bovine Serum Albumin, NOT the BsaI enzyme!
• BsaI (or AarI) restriction enzyme, 1 µl per reaction
  • Leave the BsaI and T4 ligase in the freezer until the very end, and add after the rest of the reaction has been set up.
• T4 ligase, 1 µl per reaction
  • Many protocols (both online and in the lab) call for high-concentration T4 ligase. This is only necessary if you're doing VERY LARGE Golden Gate reactions (5-8 parts). You do NOT need the HC ligase for most of the GGs you'll be doing.
  • Leave the BsaI and the T4 ligase in the freezer until the very end, and add after the rest of the reaction has been set up.
• An empty 10µl or 200µl tip box
• Ice in an ice bucket

Procedure

• Record your reaction setup.

1. A Golden Gate works best when there are equimolar amounts of each part in the reaction. By convention, we use 50 fmol of each part. For each part (plasmid and gBlock), compute the concentration (fmol/ul) and the volume required for 50 fmol in the table below. If you resuspended your gBlocks at 50 fmol/ul, you can omit them in this table.

	A	B	C	D	E
1	Part	Size (kb)	Concentration, (ng/ul)	Concentration, (fmol/ul)	Volume for 50 fmol
2	pDONR-GG	3.4	100	44.6	1.1
3
4
5

Table1

2. Record the reaction setup for each reaction below. For each reaction, include:

• -- 50 fmol of each part

• -- 1.5 ul T4 ligase buffer

• -- 1.5 ul 10X BSA

• -- 1 ul BsaI

• -- 1 ul T4 ligase

• -- Water to a total volume of 15 ul.

• -- NOTE: If you're using AarI, you need to add the AarI oligo as well!

	A	B	C	D
1	A19 L4_PhlF_R1
2	pDONR-GG L1_L2	1.1 ul
3	BT-01 gBlock	1 ul
4	BT-02 gBlock	1 ul
5	Water	6.9 ul
6	T4 ligase buffer	1.5 ul
7	10X BSA	1.5 ul
8	BsaI or AarI	1 ul
9	T4 ligase	1 ul

Table2

• Set up your thermocycler

3. Program a thermocycler with the following program:

• NOTE: If you are using AarI, use 50 cycles instead of 15.

	A	B
1	Heat lid	70°
2	Start cycle	15X
3	--	37° for 1' 30"
4	--	16° for 3' 0"
5	Close Cycle
6	50° for 5'
7	80° for 10'
8	Store at 8°

Table3

• Set up your reactions

4. Fill an empty tip box with ice, put the grey platform back on, then fill with water until just under the rack.

5. Label one thin-walled PCR tube per reaction.

6. Add the reaction components. Add the water first; add the restriction enzyme and buffer last.

7. Cap the tubes. Flick a few times to mix, then pulse spin.

8. Load the tubes in the thermocycler and start the program.

• Transform

9. Transform as usual. Plate only 10 ul of the outgrowth in a 200 ul puddle of water, or you will get a lawn of colonies.

Annealing Oligos

Introduction

Anealing Oligos

Materials

• Nuclease-free TE
  • We resuspend in TE to help supress nuclease activity that would degrade the primers.
• Annealing buffer: TE + 50 mM NaCl
• Dehydrated oligos from IDT
• One extra epi tube per oligo
• One thin-walled 200 ul PCR tube

Procedure

• Preparing stock solution

1. Briefly spin each tube BEFORE opening

2. Add sterile 1X TE (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) in a volume of uL equal to 10 times the nmol amount listed

• Concentration of stock solution is 100 uM

3. Store at -20 ^oC

• Procedure

4. Set a thermocycler to a block temperature of 95°C and a lid temperature of 105°C.

5. Dilute oligo stock solutions to 5 uM working stock (95 uL TE + 5 uL of 100 uM stock)

6. Combine 98 uL of annealing buffer, and 1 uL of 5 uM forward primer, and 1 uL of 5 uM reverse primer in a thin-walled PCR tube. Mix well.

• The concentration for the anealed oligos is 50 nM (50 fmol/uL)

7. Place tube in a standard heatblock at 90–95 °C for 3–5 minutes.

8. Remove the heat block from the apparatus and allow to cool to room temperature (or at least below 30 °C) on the workbench. Slow cooling to room temperature should take 45–60 minutes.

9. Store on ice or at 4 °C until ready to use.

Miniprep

Introduction

The miniprep uses silica gel to isolate plasmid DNA from an E. coli culture

Materials

• Buffer P1 (resuspension buffer)
  • Retrieve from refrigerator. If you are opening a new miniprep kit, add the RNAse and LyseBlue reagent and check the box on the cap.
• Buffer P2 (lysis buffer)
  • Open the cap and look at the lysis buffer. Swirl it around. If it appears cloudy, the SDS has fallen out of solution; warm it for a few minutes in the 55°C water bath.
• Buffer N3 (neutralization buffer)
• Buffer PB (binding buffer)
• Buffer PE (rinse buffer)
  • Make sure the "Ethanol added?" box has been checked. If you are opening a new miniprep kit, add absolute ethanol as per the kit instructions and check the box on the cap.
• Buffer EB (elution buffer)
• Miniprep waste container
  • Miniprep buffers contain salts that can't go down the sink.
• Per miniprep: two microcentrifuge tubes and one blue spin column, with collection tube.

Procedure

• Harvest and resuspension

1. For each culture, label two microcentrifuge tubes on the cap and one blue spin column on the side.

• The spin columns should be in their (cap-less) collection vials.

2. Pipette 1.6 mL of each culture into the corresponding microcentrifuge tubes.

• (This is for a 1.7 ml microcentrifuge tube; set your pipettor to 800 ul and do two transfers. If we ever go back to 2-ml tubes, do 1.9 ml (2x950 ul))

3. Centrifuge at maximum speed (10,000 or 13,000xg) for three minutes.

5. Aspirate the supernatant, or pour it off into the bleach bucket.

6. Pipette ANOTHER 1.8 ml of each culture into the corresponding microcentrifuge tubes.

7. Centrifuge at maximum speed for three minutes.

8. While the centrifuge is running, move the remaining cultures to 4degC.

9. Aspirate the supernatant off with the bench aspirator. Be careful not to disturb the pelleted E. coli.

• We use an aspirator here because the less extra salt and protein we put in the miniprep, the better the yield is.

10. Add 250 µl Buffer P1 to each tube.

11. Resuspend the E. coli pellet. The preferred way is with the roto-mixer at the other end of the lab.

• Alternately, if you have just a few tubes, you can resuspend on a vortex.

• Make sure to resuspend fully and thoroughly. The resulting suspension should be smooth and cloudy; if there is particulate matter floating around, vortex some more.

• Lysis

12. Add 250 µl Buffer P2 to each tube.

• Work quickly; the lysis step should take less than 5 minutes.

13. Snap the tubes closed and invert them 4-6 times, until the tube is thoroughly mixed and the entire solution turns blue.

• If you have many many tubes, you can stack a second tube rack on top of them and invert the entire thing.

14. Add 350 ul Buffer N3 to each tube.

15. Snap the tubes closed and invert 4-6 times, until the solution is thouroughly mixed and no longer blue.

• The solution will become cloudy or flocculent.

16. Centrifuge on high speed for 10 minutes.

• Separation

17. Remove the tubes from the microcentrifuge, being careful not to disturb the white pellet.

18. Using P-1000 micropipettor set to 850 ul, carefully transfer the supernatant from each centrifuge tube to the corresponding blue spin column.

19. Centrifuge the spin columns for 30 seconds at maximum speed.

• Don't forget to put the lid on the rotor! Some of the salts get aerosolized because the spin columns don't have caps.

20. Pour the flow-through from each column into the miniprep waste container.

21. Pipette 500 ul of Buffer PB onto each spin column.

22. Centrifuge the spin columns for 30 seconds at maximum speed.

23. Pour the flow-through from each column into the miniprep waste container.

24. Pipette 750 ul of Buffer PE onto each spin column

25. Wait 1-3 minutes.

• This allows some of the salt that's still bound to the silica matrix to resuspend in the buffer.

26. Centrifuge the spin columns for 30 seconds at maximum speed.

27. Pour the flow-through from each column into the miniprep waste container.

28. Return each spin column to its collection tube and centrifuge an additional 1 minute at high speed.

• This removes every last trace of buffer PE; the ethanol can screw up downstream steps.

29. Transfer each spin column to a clean labelled microcentrifuge tube.

30. Pipette 50 ul of Buffer EB onto the center of each column.

• The volume of EB is comparable to the volume of silica gel matrix; if you pipette down the side, you might not get the entire transfer to the matrix.

31. Wait 1-3 minutes.

• This gives the DNA a chance to dissociate from the silica matrix.

32. Centrifuge the spin columns, in their collection tubes, for one minute at maximum speed.

33. Proceed directly to analyze the samples on the Nanodrop.

Resuspending gBlocks

Introduction

gBlocks are long double-stranded linear pieces of DNA that arrive from IDT dehydrated; to use them, you need to resuspend them in TE to a known concentration. They are synthesized on a much smaller scale than primers and other oligonucleotides; we generally resuspend to a concentration of 50 fmol/µl.

Materials

• Nuclease-free TE
  • We resuspend in TE to help supress nuclease activity that would degrade the gBlocks
• Dehydrated gBlocks from IDT

Procedure

• Resuspend the dried gBlock

1. Label the top of the gBlock tube. I recommend the group initials and a number.

2. In the little microfuge, spin the (dry) gBlocks briefly.

• Sometimes the freeze-dried gBlocks flake off the bottom of the tube.

3. Determine how many fmol of gBlock are in the tube.

• The number of fmol is on the bottom line of the label. For example, one of my gBlocks says

• "500 ng = 1879 fmol"

4. Resuspend in the appropriate amount of TE.

• For my gBlock, if there are 1879 fmol in the tube, and I want 50 fmol/ul, then I need to resuspend in 37.5 ul of TE.

5. Vortex briefly. Pulse spin.

Resuspending Primers

Introduction

Primers arrive from IDT dehydrated; to use them, you need to resuspend them in TE to a known concentration. By convention, the stock concentration (in the blue-capped IDT tube) is 100 µM; the working concentration depends on the application: - For PCR, the working concentration is usually 10 µM - For sequencing, the working concentration is usually 5 µM

Materials

• Nuclease-free TE
  • We resuspend in TE to help supress nuclease activity that would degrade the primers.
• Dehydrated primers from IDT
• One extra epi tube per primer
  • ...for the working stock.

Procedure

• Resuspend the dried primer

1. Label the top of the oligo tubes. I recommend the group initials and a number.

2. In the little microfuge, spin the (dry) oligos briefly.

• Sometimes the freeze-dried primers flake off the bottom of the tube.

3. Determine how many nanomoles of primer are in the tube.

• This is the bottom line on the tube label. As an example, the BT-01 primer says

• 7.3OD = 33.4 nmol = 0.23 mg. <-- the number of nanomoles is "33.4".

4. Add 10 µl of nuclease-free TE for each nanomole of primer.

• For the BT-01 primer, this volume is 334 µl.

5. Vortex briefly.

6. Check the bottom of the tube to see if the primer is fully resuspended. If not, vortex again.

7. Pulse spin the primers.

• Make a working stock

8. Label the top of the epi tube the same as the oligo.

9. Determine the working stock concentration that you want. Label the side of the epi tube with the working concentration.

10. Make 100 µl of working stock.

• For example, if you're making a 10 µM working stock, dilute 10 µl of the concentrated stock into 90 µl of TE.

11. Vortex briefly.

12. Freeze both the concentrated and working stocks in the primers box.

Transformation of E. coli

Introduction

Transformation is the process of inducing chemically competent E. coli to take up DNA.

Materials

• Dry bath, set to 42°C
  • Fill the wells in the dry bath block 1/2 full with DI water.
• Ice bucket, with ice
  • For thawing competent cells.
• DNA to transform
  • Could be an assembly reaction (LR, Golden Gate, etc) or a miniprepped plasmid.
  • If you removed it from the freezer, make sure it's entirely thawed out.
• pUC19 Transformation Control, 1 pg/µl
  • The pUC19 control will tell you how efficient your transformations were.
• SOC growth media, at room temperature
  • Check to make sure it's clear and NOT CLOUDY.
• Antibiotic plates, one per transformation, plus 1 Amp plate for the pUC19 control
  • Make sure the plates you use match the resistance cassette of the plasmid!
• Competent E. coli, one tube per transformation + one for the pUC19 control
  • These live in the -80 in 235.
  • Thaw on ice 3-4 minutes.
• A timer, set for 30 seconds.

Procedure

• Setup

1. Make sure the dry bath is set to 42°C and the wells in the block are 1/2 full of DI water

2. Remove selection plates from the refrigerator. Double-check that they match the selection marker on your plasmid, then place them in the 37° incubator.

3. Retrieve the DNA to transform.

• If frozen: thaw, completely, flick a few times to mix, then pulse down in the microfuge.

4. Fill an ice bucket with ice. Retrieve one tube of competent E. coli per transformation from the -80 and thaw on ice, 3-4 minutes.

00:03:00

5. While the transformation tubes are thawing, label their tops with something descriptive. Record the labels here:

	A	B	C	D	E	F	G	H
1	puc-19L
2	puc-19TC
3	Pdest_mCherry
4	pDest

Table1

• Transformation

6. Add 2 µl DNA from each reaction to a tube of competent cells.

• Immediately after adding the DNA to each tube, stir the cells a few times with the pipette tip.

7. Add 1 µl of the pUC19 transformation control to the positive control tube.

8. Incubate on ice for 30 minutes.

00:30:00

9. Heat shock the cells for exactly 30 seconds in the 42° heat block. (Yes, set a timer.)

00:00:30

10. Place back on ice for 2 minutes.

00:02:00

11. Add 250 µl SOC to each tube.

12. Tape the tubes to the platform of a shaker at 37°C and shake at 270 RPM for 60 minutes.

01:00:00

• Plating

13. Label the selection plates using the labels you recorded above.

14. Shake ~10 plating beads onto each plate.

15. Pipette 100 µl of each transformation onto the corresponding plates. NOTE: if you are using DNA from a golden gate reaction, see the golden gate protocol for instructions to dilute your sample.

16. -----Golden Gate: Plate only 10 ul of the outgrowth in a 200 ul puddle of water, or you will get a lawn of colonies.

17. Cover the plates and shake the beads around to spread the cells out.

18. Dispose of the beads by tapping them into the waste container.

19. Incubate the plates upside down overnight in the 37° incubator.

• Don't incubate for more than 18-24 hours.

• Compute transformation efficiency

20. Count the colonies on your positive transformation plate.

• If there are many many colonies, then hooray! You had a great transformation. Just estimate.

21. Divide the number of colonies by the fraction of the transformation you plated.

• So, if you resuspended your transformation in a total volume of 300 ul, then plated 100 ul, multiply the number of colonies by 3.

22. Transformation efficiency is expressed in colonies per microgram pUC19. Multiply the number of colonies by the appropriate conversion factor.

• So if you transformed 1 picogram of pUC19 DNA, multiply by 10⁶.

23. Record your transformation efficiency in your (daily) lab notebook.

Pouring LB Agar Plates

Introduction

The media must be autoclaved before you pour plates. This is not something for iGEMers to do; ask an advisor.

Materials

• One sleeve of empty plates per 500 ml of media
  • Make sure you are using the Weiss lab's bacteriological plates, not tissue culture plates.
• LB-Agar media, either solid or already melted
• 1000X stock solution of the antibiotic
  • (Amp)icillin, (Kan)amycin, (Chlor)amphenicol

Procedure

• Making media

1. Make sure you don't already have a bunch of the plates already in the fridge

2. Fill a 1-liter bottle with 500 ml of water.

3. Weigh out XXXX of powdered LB-Agar (check the bottle for the weight.)

• IF YOU MAKE A MESS OF THE WEIGH STATION, CLEAN UP AFTER YOURSELF.

4. Ask an instructor to autoclave your media.

• Melting media

5. Solid media must be melted in the microwave. Be careful -- it gets very hot.

6. Loosen the cap to prevent the bottle from exploding. 1 full turn of the cap is sufficient.

7. Microwave 2-3 minutes, watching carefully, until the media begins to melt and bubble.

8. Continue microwaving in short bursts until the media is completely melted.

• IF YOU MAKE A MESS OF THE MICROWAVE, CLEAN UP AFTER YOURSELF.

9. Cool the media to <= 60°C before adding the antibiotics.

• You can either swirl it under a cold tap until you can handle it without burning yourself, or you can leave it for a few hours in the 55°C water bath.

• To check the media temperature, swirl the bottle, wait ~30 seconds and measure with an IR thermometer. (Pew pew.)

• Pouring plates

10. Add antibiotic.

• Unless otherwise noted, it is 1000X -- so for 500 ml of media, add 500 µl of antibiotic.

• Note: If you're pouring Golden Gate plates, XGal is 500X! (for 500 ml of media, add 1000 µl of XGal)

11. Pour all of the media you add these too. Refreezing and thawing has unpredictable results on the amount of antibiotic destroyed by the heat, so to avoid over/under dosing don't refreeze. 500 ml makes a sleeve of plates.

12. Tighten the lid to avoid spills and swirl the bottle to mix

13. Use a seriological pipette to add 20ml to each plate.

• Use sterile practices

• If you get bubbles, use the pipette to try and suck the air out of the bubbles, this removes them

• 20ml is a rough estimate, what is important is that you get a solid layer with no holes.

14. Mark the plates with a stripe of marker up the side, based on which antibiotic(s) you added:

• Blue: Amp

• Red: Kan

• Green: Cm

• Black: Non-antibiotic additive (like X-gal)

15. Dry on the bench overnight.

16. Re-stack the plates upside-down and label the sleeve with the antibiotics, your initials and the date.

Diagnostic Restriction Digestion

Introduction

A diagnostic restriction digest helps identify correctly assembled clones from incorrect clones.

Materials

• 200 ul PCR strip tubes, one per reaction
• Restriction enzyme (chosen below), 1 ul per reaction.
• 10X restriction enzyme buffer (chosen below), 1 ul per reaction.

Procedure

• Choose A Good Restriction Enzyme

1. Using Benchling, choose a restriction enzyme that meets the following criteria:

• - Cuts at least once in the insert (for a pEXPR, either the promoter or the gene.)

• - Cuts at least once in the backbone.

• - Gives bands that aren't too large (> 8 kb) or too small (< 200 bp).

• - Gives a band pattern that is significantly different from the expected error mode.

• * For LR reactions, the most common error is a pDEST that slipped through the selection.

• - Begin in the "Brian's Favorites" list, then expand to the main Weiss lab list.

• - If you can't find a single enzyme that gives an acceptable band pattern, choose two enzymes that give a acceptable band pattern when used together.

• * This double-digest is subject to buffer compatibility, outlined below.

2. Record your enzyme choice on the plasmid's Description page.

3. Benchling will tell you the enzyme's buffer compatibility and active temperature. Record the buffer in which the enzyme is most active.

• If there are multiple bufers in which the enzyme is equally active, choose in this order: Buffer 3.1, Buffer CS, Buffer 2.1, Buffer 1.1.

• Set up the restriction digest

4. Retrieve the minipreps and the appropriate 10X buffer concentrate from the freezer. Thaw on the benchtop or in your fingers.

5. Label the PCR tubes with your initials and an incrementing number.

• ie: BT-1, -2, -3, -4

6. Vortex the minipreps and the 10X buffer concentrate briefly, then pulse down in the microfuge.

7. For each miniprep, set up a PCR tube containing the following in order:

• - 5 ul enzyme-quality H2O

• - 1 ul enzyme buffer

• - 3 ul miniprep DNA

• - 1 ul enzyme

• Remove the enzyme from the freezer for as little time as possible.

• I have specified an "arbitrary" 3 µl volume of miniprep DNA; this should be fine as long as your miniprep concentration is >= 100 ng/ul.

8. Flick the strip tubes a few times to mix the reaction, then pulse down in the strip tube microfuge.

9. Incubate at the appropriate temperature for at least 1 hour and more more than 16 hours.

• If the enzyme's active temperature is 37°C, use the 37°C plate incubator.

10. Stop the reaction by adding 2 ul of 6X NEB purple gel loading dye to each reacti on.

11. Flick the strip tubes a few times to mix the reaction, then pulse down in the strip tube microfuge.

• PAUSE POINT: The reaction can be stored almost indefinitely at room temperature once it's been stopped.

12. Proceed to gel electrophoresis.

JG 7/19/16

Made with Benchling

Project: iGem 2016 _ Recombinase subgroup

Authors: Trinh Nguyen

Date: 2016-07-19

Tuesday, 7/19

Silico digest

clipboard_2016-07-19_11:01:46.png

Top gel image:

hEF1a:mKate: Single digest AflII

hEF1a:EYFP: Single digest AflII

EGSH:SCRE: Single digest NcoI

EGSH:SCRE: Double digest NcoI+BglII

Botton gel image:

EGSH:FLPO: Single digest NcoI

EGSH:FLPO: Double digest NcoI+XbaI

RecombinaseLRTOP.jpg

To Sequnce: 1-1 (hef1a-eyfp), 1-2,

2-1, 2-2, (hef1a-mKate)

3-1, 3-2, (EGSH-SCre)

4-1, 4-2 (EGSH-FLPO)

50*30/concentration

RecombinaseLRBOTTOM.jpg

Transfection:

Cell line: HEK293

Total DNA: 500ng

Total volume: 50uL

Plate map:

	A	B	C	D
1	Untransfect control	Single color: 250ng hEF1a:mKate 250ng pDONR	Single color: 250ng hEF1a:BFP 250ng pDONR	Two color: 250ng hEF1a:mKate 250ng hEF1a:BFP
2	300ng EGSH:mKate 100ng hEF1a:VgEcr 100ng hEF1a:BFP 0uM	300ng EGSH:mKate 100ng hEF1a:VgEcr 100ng hEF1a:BFP 0.5uM	300ng EGSH:mKate 100ng hEF1a:VgEcr 100ng hEF1a:BFP 1uM	300ng EGSH:mKate 100ng hEF1a:VgEcr 100ng hEF1a:BFP 5uM
3	300ng EGSH:mKate 100ng hEF1a:VgEcr 100ng hEF1a:BFP 10uM

Table1

Transfection mixture:

	A	B	C	D
1	uL	Concentration (ng/uL)	1 well	6 wells
2	EGSH:mKate	100	3	18
3	hEF1a:VgEcr	100	1	6
4	hEF1a:BFP	100	1	6
5	Viafect		1	6
6	Opt Mem		44	264
7	Total		50	300

Table2

	A	B	C	D	E
1	uL	Concentration (ng/uL)	Single color (mKate)	Single color (BFP)	Two color
2	hEF1a:mKate	100	2.5	0	2.5
3	hEF1a:BFP	100	0	2.5	2.5
4	pDONR	200	1.5	1.5	0
5	viafect		1	1	1
6	Opt Mem		45	45	44
7	Total		50	50	50

Table3

Gel Extraction:

Weight of Gel Band: 0.062g

Pouring LB Agar Plates

Introduction

The media must be autoclaved before you pour plates. This is not something for iGEMers to do; ask an advisor.

Materials

• One sleeve of empty plates per 500 ml of media
  • Make sure you are using the Weiss lab's bacteriological plates, not tissue culture plates.
• LB-Agar media, either solid or already melted
• 1000X stock solution of the antibiotic
  • (Amp)icillin, (Kan)amycin, (Chlor)amphenicol

Procedure

• Making media

1. Make sure you don't already have a bunch of the plates already in the fridge

2. Fill a 1-liter bottle with 500 ml of water.

3. Weigh out XXXX of powdered LB-Agar (check the bottle for the weight.)

• IF YOU MAKE A MESS OF THE WEIGH STATION, CLEAN UP AFTER YOURSELF.

4. Ask an instructor to autoclave your media.

• Melting media

5. Solid media must be melted in the microwave. Be careful -- it gets very hot.

6. Loosen the cap to prevent the bottle from exploding. 1 full turn of the cap is sufficient.

7. Microwave 2-3 minutes, watching carefully, until the media begins to melt and bubble.

8. Continue microwaving in short bursts until the media is completely melted.

• IF YOU MAKE A MESS OF THE MICROWAVE, CLEAN UP AFTER YOURSELF.

9. Cool the media to <= 60°C before adding the antibiotics.

• You can either swirl it under a cold tap until you can handle it without burning yourself, or you can leave it for a few hours in the 55°C water bath.

• To check the media temperature, swirl the bottle, wait ~30 seconds and measure with an IR thermometer. (Pew pew.)

• Pouring plates

10. Add antibiotic.

• Unless otherwise noted, it is 1000X -- so for 500 ml of media, add 500 µl of antibiotic.

• Note: If you're pouring Golden Gate plates, XGal is 500X! (for 500 ml of media, add 1000 µl of XGal)

11. Pour all of the media you add these too. Refreezing and thawing has unpredictable results on the amount of antibiotic destroyed by the heat, so to avoid over/under dosing don't refreeze. 500 ml makes a sleeve of plates.

12. Tighten the lid to avoid spills and swirl the bottle to mix

13. Use a seriological pipette to add 20ml to each plate.

• Use sterile practices

• If you get bubbles, use the pipette to try and suck the air out of the bubbles, this removes them

• 20ml is a rough estimate, what is important is that you get a solid layer with no holes.

14. Mark the plates with a stripe of marker up the side, based on which antibiotic(s) you added:

• Blue: Amp

• Red: Kan

• Green: Cm

• Black: Non-antibiotic additive (like X-gal)

15. Dry on the bench overnight.

16. Re-stack the plates upside-down and label the sleeve with the antibiotics, your initials and the date.

Gel electrophoresis

Introduction

Gel electrophoresis separates pieces of DNA by length.

Materials

• Agarose gel, 1 lane per sample, plus at least 1 lane for the ladder.
  • If you just poured the gel, make sure it has had at least 30 minutes to set.
  • If you have a large gel but only need a few lanes, cut out just a piece of it with a razor blade. Cut straight!
• Samples to run
• 6X NEB Purple Loading Dye
• Molecular weight standard (aka "ladder")
  • Commonly available ladders are Hyperladder I and NEB 2-Log.
• Gel box, lid, leads
• Electrophoresis power supply
• 1X TAE buffer, enough to fill the gel box.

Procedure

• Prepare your samples

1. If your samples are frozen, thaw them completely, flick or vortex to mix, then pulse down in the microfuge.

2. If your samples are not already in loading dye, mix them 1:6 with 6X NEB Purple Loading Dye

• Add 1 µl of loading dye for every 5 µl of sample.

• Pro tip: If you don't want to mix loading gel with your entire sample (PCR products, for example), cut off a strip of Parafilm; pipette 2-3 ul of sample onto the parafilm; add 1 ul loading dye; mix by pipetting and load directly.

• Set up the gel box

3. Program the voltage on the power supply. For the small gel box, use 100V; for the large gel box, use 150V.

4. Program the time on the power supply. For a small analytical gel, set the timer for 30 minutes.

• For larger gels, start at 45 minutes, and then check regularly!

5. Attach the leads to the gel box cover. Make sure that the red (positive) lead is attached to the side of the box farthest away from the wells in the gel.

• Remember, the DNA is negatively charged and will move toward the positive terminal. A useful neumonic for remembering this is "Run to Red."

6. If the gel box has not been used previously that day, empty it and rinse it out with DI water.

• The TAE buffer can be re-used, but if it's been sitting out for too long it evaporates and the salt concentration (and conductivity) changes. TAE is cheap; when in doubt, replace it.

7. Place the gel in the gel box.

8. Pour TAE into the gel box until it just barely covers the gel.

• The TAE's purpose is to conduct electricity; over-filling the gel box results in a larger conductive path, more current flow, and more heating (which can screw up your run.)

9. Check the wells to see that they are free of bubbles. If there are bubbles, blow them out by pipetting 100 µl of TAE from the gel box into the well.

• Load your samples

10. Load your samples. For large combs, load 10 µl; for small combs, load 5 µl.

• Work quickly. The samples begin to diffuse in the buffer, leading to smeared bands.

11. Load the ladder in the last lane. Load 1/2 the volume of your samples: for large combs, load 5 µl, for small combs, load 2.5 µl.

12. Place the lid on the gel box. Make sure it is seated on the brass contacts.

• Run the gel

13. Press the Start or Run button on the power supply.

14. Make sure the power supply doesn't complain about an open circuit. If it does, re-seat the gel box lid and press Run again.

15. Double-check that there are bubbles forming on the platinum wires at either end of the gel box.

16. Double-check that the red (positive) lead is on the side farthest from the wells. Remember, Run to Red.

17. Run until the pink band is 2/3 to 3/4 of the way down the gel. For small analytical gels, this should take 30 minutes. For larger gels, start at 45 minutes and check regularly!

• Image the gel

18. If the power supply is still running, press the Stop button.

19. Lift the lid off of the gel box. Lift the lid straight up. If you try to "hinge" it up, the lid will break.

20. Transfer the gel to the GelDoc.

21. If it's not running, start QuantityOne from the toolbar.

• If necessary, click the top button on the toolbox to select the scanner.

• If necessary, reset the camera (as per instructions on the GelDoc computer.)

22. Press the Epifluorescent Illumination button on the GelDoc. Check the gel's position, zoom and focus.

• You want to be zoomed in so that the gel fills the field of view, and focussed so that the well edges are sharp.

23. Close the GelDoc door. Click Auto Expose.

• The Auto Expose functionality generally over-exposes my gels a bit. If your gel is over-exposed, remove 1/3 of the exposure time, type it into the Exposure box, then click Manually Expose.

24. In the File menu, select, Export as JPEG.... Save your gel to the iGEM folder on the desktop.

25. Open Benchling. Log in, and copy the gel to the Description page for the plasmid you're building.

• If there are multiple plasmids on the gel, save it to each plasmid's Description.

26. Discard the gel in the biowaste box. Wipe down the gel doc with a little water and a paper towel or Kimwipe.

• Annotate the gel

27. Immediately, before you forget what's where, annotate the gel.

• List what is in each lane.

• Describe whether the pattern is what you expected or not. (You should have an in silico digestion to compare it to!)

Overnight liquid cultures (picking colonies)

Introduction

Overnight cultures are used to prepare miniprep DNA.

Materials

• The plate from which you are picking colonies
• 15 ml round-bottom polystyrene tubes, one per culture
  • The ones with the snap caps, NOT conical tubes with screw caps
• 5 mL LB per culture
• A container that can hold 5ml x the number of cultures
  • For a modest number of minipreps, a 50 ml conical tube works well.
  • For larger minipreps, use a sterile bottle (100 ml is frequently useful.)
• Antibiotic stock, 1000X

Procedure

• Materials Setup

1. Warm up the LB to at least room temperature (if it came from the fridge), but not warmer than 37°C

2. Label one round-bottom culture tube for each miniprep. Use "NAME-1, NAME-2, ..."etc for the naming convention, where NAME is a shortened name of the plasmid (eg, "hEF1a:mKate").

• Your impulse is to just use number, or initials and number, but trust me -- you will want to be able to identify this tube in three weeks when you've forgotten what you were doing.

3. Using a sterile pipette, transfer 5 ml of LB to the mixing container for each culture PLUS 5 ML.

4. Add antibiotic stock to a final concentration of 1X (1 µl stock for each 1 ml in the mixing container.)

5. Cap tightly and mix well.

• Culture Setup

6. Using a sterile pipette, transfer 5 ml of LB+antibiotic to each round-bottom culture tube.

• If you are making cultures with different antibiotics, take care that the right media goes in each tube.

7. Squirt ethanol on a pair of foreceps and wipe dry with a Kimwipe.

8. Use the foreceps to pick up a sterile 200µl pipette tip, scrape a colony off of the plate, and drop the pipette tip in the corresponding tube.

• Repeat for each tube.

9. Transfer to an incubating shaker at 37°C and incubate 14-16 hours.

• Don't over-grow too badly, or your yield will suffer.

• If you need to grow longer, you can grow at 30°C instead for 20 hours.

Transformation of E. coli

Introduction

Transformation is the process of inducing chemically competent E. coli to take up DNA.

Materials

• Dry bath, set to 42°C
  • Fill the wells in the dry bath block 1/2 full with DI water.
• Ice bucket, with ice
  • For thawing competent cells.
• DNA to transform
  • Could be an assembly reaction (LR, Golden Gate, etc) or a miniprepped plasmid.
  • If you removed it from the freezer, make sure it's entirely thawed out.
• pUC19 Transformation Control, 1 pg/µl
  • The pUC19 control will tell you how efficient your transformations were.
• SOC growth media, at room temperature
  • Check to make sure it's clear and NOT CLOUDY.
• Antibiotic plates, one per transformation, plus 1 Amp plate for the pUC19 control
  • Make sure the plates you use match the resistance cassette of the plasmid!
• Competent E. coli, one tube per transformation + one for the pUC19 control
  • These live in the -80 in 235.
  • Thaw on ice 3-4 minutes.
• A timer, set for 30 seconds.

Procedure

• Setup

1. Make sure the dry bath is set to 42°C and the wells in the block are 1/2 full of DI water

2. Remove selection plates from the refrigerator. Double-check that they match the selection marker on your plasmid, then place them in the 37° incubator.

3. Retrieve the DNA to transform.

• If frozen: thaw, completely, flick a few times to mix, then pulse down in the microfuge.

4. Fill an ice bucket with ice. Retrieve one tube of competent E. coli per transformation from the -80 and thaw on ice, 3-4 minutes.

00:03:00

5. While the transformation tubes are thawing, label their tops with something descriptive. Record the labels here:

	A	B	C	D	E	F	G	H
1	puc-19L
2	puc-19TC
3	Pdest_mCherry
4	pDest

Table1

• Transformation

6. Add 2 µl DNA from each reaction to a tube of competent cells.

• Immediately after adding the DNA to each tube, stir the cells a few times with the pipette tip.

7. Add 1 µl of the pUC19 transformation control to the positive control tube.

8. Incubate on ice for 30 minutes.

00:30:00

9. Heat shock the cells for exactly 30 seconds in the 42° heat block. (Yes, set a timer.)

00:00:30

10. Place back on ice for 2 minutes.

00:02:00

11. Add 250 µl SOC to each tube.

12. Tape the tubes to the platform of a shaker at 37°C and shake at 270 RPM for 60 minutes.

01:00:00

• Plating

13. Label the selection plates using the labels you recorded above.

14. Shake ~10 plating beads onto each plate.

15. Pipette 100 µl of each transformation onto the corresponding plates. NOTE: if you are using DNA from a golden gate reaction, see the golden gate protocol for instructions to dilute your sample.

16. -----Golden Gate: Plate only 10 ul of the outgrowth in a 200 ul puddle of water, or you will get a lawn of colonies.

17. Cover the plates and shake the beads around to spread the cells out.

18. Dispose of the beads by tapping them into the waste container.

19. Incubate the plates upside down overnight in the 37° incubator.

• Don't incubate for more than 18-24 hours.

• Compute transformation efficiency

20. Count the colonies on your positive transformation plate.

• If there are many many colonies, then hooray! You had a great transformation. Just estimate.

21. Divide the number of colonies by the fraction of the transformation you plated.

• So, if you resuspended your transformation in a total volume of 300 ul, then plated 100 ul, multiply the number of colonies by 3.

22. Transformation efficiency is expressed in colonies per microgram pUC19. Multiply the number of colonies by the appropriate conversion factor.

• So if you transformed 1 picogram of pUC19 DNA, multiply by 10⁶.

23. Record your transformation efficiency in your (daily) lab notebook.

Gel Extraction

Introduction

Get started by giving your protocol a name and editing this introduction.

Materials

• QIAgen gel extraction kit
  • Purple columns
  • Buffer QG
  • Buffer PE
  • Buffer EB
• Isopropanol
• 3M Sodium Acetate, pH=5.0
• Microcentrifuge tubes

Procedure

• Setup

1. Set a dry bath to 50°C and fill the holes in the metal block with water. Transfer some Buffer EB (30 ul for each purification plus an extra 30 ul) to a microcentrifuge tube and place it in the dry back to warm.

• Excise gel

2. On the blue-light transilluminator, cut out the gel band and put it in an eppendorf tube.

3. Weigh the gel band.

• TARE the balance with an empty eppendorf tube.

• Weigh the tube with the gel band in it.

• Gel extraction

4. Add 6 volumes of Buffer QG to 1 volume of gel. (Compute volume assuming 100 mg of gel ~ 100 ul.)

5. Incubate at 50°C until the gel is disolved, about 10 minutes; vortex briefly every 2-3 minutes.

6. Add 10 µl 3M sodium acetate pH 5.0. Vortex briefly.

7. Add 1 gel volume of isopropanol.

8. Apply a maximum of 750 µl of the disolved gel to the QIAquick column and centrifuge for 30 seconds at maximum speed.

9. Pipette the flow-through back into the column and centrifuge again. Discard the flow-through.

10. If needed, apply the rest of the disolved gel to the column. Centrifuge; then re-apply and centrifuge again. Discard flow-through.

11. Add 500 µl of buffer QG and centrifuge 30 seconds. Discard flow-through.

12. Add 750 µl of Buffer PE to the column. Incubate at room temperature for 3-5 minutes, then centrifuge for 30 seconds.

13. Discard flow-through and centrifuge again for 1 minute.

14. Move column to a clean microcentrifuge tube.

15. Add 30 µl of the warm EB buffer to the center of the column.

16. Incubate in the 50-degree dry bath for 3 minutes.

17. Centrifuge for 1 minute.

18. Measure the concentration using the nanodrop (make sure to vortex your samples before).

DNA Transfection into HEK Cells

Introduction

Get started by giving your protocol a name and editing this introduction.

Materials

• OptiMEM (Serum-free media)
• Viafect
• DNA from midipreps

Procedure

• Phase I: Calculations

1. Select DNA to be transfected and determine the concentration through Nanodrop

2. Determine the volume of DNA required to have 0.5 ug in reaction mixture.

• Ex. for a 521 ng/ul stock of DNA you would need approximately 1 uL. If the calculated volume you need is less than 0.5 uL (i.e. hard to pipette) you may want to dilute your DNA solution so that you can get an appropriate volume.

3. Determine the amount of Viafect to use from the following table:

	A	B	C	D	E	F	G	H
1	Volume of cells in media (per well)	Total volume transfection complex (per well)	Amount of DNA (per well)	Amt of Viafect Reagent for 1.5:1 Reagent:DNA	2:1	3:1	4:1	6:1
2	500 uL	50 uL	0.5 ug	0.75 uL	1 uL	1.5 uL	2 uL	3 uL

Table1

4. Calculate the amount of serum free DMEM media required to get a reaction volume of 50 uL

• Ex. if you are adding 1 uL of DNA and 3 uL of Viafect, you would want to add 46 uL of DMEM

• Phase II: Reaction Mixture

5. This phase consists of mixing the reactants required to create the transfection complex. Do so for all ratios of reagent to DNA above - add media first, then reagent, then DNA.

6. Let the reaction mixture sit for 10-20 minutes

• Closer to 20 min is better

• Phase III: Transfection

7. Retrieve cells from incubator. Make sure to leave this step for last because you don’t want to keep the cells at room temperature for too long.

8. Add entire 50 uL transfection complex to each well in the well plate and store your cells back in the incubator.

• Phase IV: Checking for Fluorescence

9. Close off lights in room and turn on the microscope’s light.

10. Apply the appropriate fluorescence filter.

11. Check for fluorescence.

JG 7/20/16

Made with Benchling

Project: iGem 2016 _ Recombinase subgroup

Authors: Maya Kaul

Date: 2016-07-20

Wednesday, 7/20

Miniprep Concentrations - 1 is SloxP-SV40 GG, 2 is FRT-SV40 GG

	A	B
1	1-1	226.2 ng/ul
2	1-2	239.6 ng/ul
3	1-3	205.7 ng/ul
4	1-4	226.2 ng/ul
5	2-1	271.8 ng/ul
6	2-2	241.9 ng/ul
7	2-3	304.0 ng/ul
8	2-4	360.1 ng/ul

Table2

Gel Data:

RecombinaseGG.jpg

E, L, 1-1, 1-2, 1-3, 1-4, L, 2-1, 2-2, 2-3, 2-4, E

clipboard_2016-07-15_14:55:25.png

Gel Extraction

Introduction

Get started by giving your protocol a name and editing this introduction.

Materials

• QIAgen gel extraction kit
  • Purple columns
  • Buffer QG
  • Buffer PE
  • Buffer EB
• Isopropanol
• 3M Sodium Acetate, pH=5.0
• Microcentrifuge tubes

Procedure

• Setup

1. Set a dry bath to 50°C and fill the holes in the metal block with water. Transfer some Buffer EB (30 ul for each purification plus an extra 30 ul) to a microcentrifuge tube and place it in the dry back to warm.

• Excise gel

2. On the blue-light transilluminator, cut out the gel band and put it in an eppendorf tube.

3. Weigh the gel band.

• TARE the balance with an empty eppendorf tube.

• Weigh the tube with the gel band in it.

• Gel extraction

4. Add 6 volumes of Buffer QG to 1 volume of gel. (Compute volume assuming 100 mg of gel ~ 100 ul.)

5. Incubate at 50°C until the gel is disolved, about 10 minutes; vortex briefly every 2-3 minutes.

6. Add 10 µl 3M sodium acetate pH 5.0. Vortex briefly.

7. Add 1 gel volume of isopropanol.

8. Apply a maximum of 750 µl of the disolved gel to the QIAquick column and centrifuge for 30 seconds at maximum speed.

9. Pipette the flow-through back into the column and centrifuge again. Discard the flow-through.

10. If needed, apply the rest of the disolved gel to the column. Centrifuge; then re-apply and centrifuge again. Discard flow-through.

11. Add 500 µl of buffer QG and centrifuge 30 seconds. Discard flow-through.

12. Add 750 µl of Buffer PE to the column. Incubate at room temperature for 3-5 minutes, then centrifuge for 30 seconds.

13. Discard flow-through and centrifuge again for 1 minute.

14. Move column to a clean microcentrifuge tube.

15. Add 30 µl of the warm EB buffer to the center of the column.

16. Incubate in the 50-degree dry bath for 3 minutes.

17. Centrifuge for 1 minute.

18. Measure the concentration using the nanodrop (make sure to vortex your samples before).

Miniprep

Introduction

The miniprep uses silica gel to isolate plasmid DNA from an E. coli culture

Materials

• Buffer P1 (resuspension buffer)
  • Retrieve from refrigerator. If you are opening a new miniprep kit, add the RNAse and LyseBlue reagent and check the box on the cap.
• Buffer P2 (lysis buffer)
  • Open the cap and look at the lysis buffer. Swirl it around. If it appears cloudy, the SDS has fallen out of solution; warm it for a few minutes in the 55°C water bath.
• Buffer N3 (neutralization buffer)
• Buffer PB (binding buffer)
• Buffer PE (rinse buffer)
  • Make sure the "Ethanol added?" box has been checked. If you are opening a new miniprep kit, add absolute ethanol as per the kit instructions and check the box on the cap.
• Buffer EB (elution buffer)
• Miniprep waste container
  • Miniprep buffers contain salts that can't go down the sink.
• Per miniprep: two microcentrifuge tubes and one blue spin column, with collection tube.

Procedure

• Harvest and resuspension

1. For each culture, label two microcentrifuge tubes on the cap and one blue spin column on the side.

• The spin columns should be in their (cap-less) collection vials.

2. Pipette 1.6 mL of each culture into the corresponding microcentrifuge tubes.

• (This is for a 1.7 ml microcentrifuge tube; set your pipettor to 800 ul and do two transfers. If we ever go back to 2-ml tubes, do 1.9 ml (2x950 ul))

3. Centrifuge at maximum speed (10,000 or 13,000xg) for three minutes.

5. Aspirate the supernatant, or pour it off into the bleach bucket.

6. Pipette ANOTHER 1.8 ml of each culture into the corresponding microcentrifuge tubes.

7. Centrifuge at maximum speed for three minutes.

8. While the centrifuge is running, move the remaining cultures to 4degC.

9. Aspirate the supernatant off with the bench aspirator. Be careful not to disturb the pelleted E. coli.

• We use an aspirator here because the less extra salt and protein we put in the miniprep, the better the yield is.

10. Add 250 µl Buffer P1 to each tube.

11. Resuspend the E. coli pellet. The preferred way is with the roto-mixer at the other end of the lab.

• Alternately, if you have just a few tubes, you can resuspend on a vortex.

• Make sure to resuspend fully and thoroughly. The resulting suspension should be smooth and cloudy; if there is particulate matter floating around, vortex some more.

• Lysis

12. Add 250 µl Buffer P2 to each tube.

• Work quickly; the lysis step should take less than 5 minutes.

13. Snap the tubes closed and invert them 4-6 times, until the tube is thoroughly mixed and the entire solution turns blue.

• If you have many many tubes, you can stack a second tube rack on top of them and invert the entire thing.

14. Add 350 ul Buffer N3 to each tube.

15. Snap the tubes closed and invert 4-6 times, until the solution is thouroughly mixed and no longer blue.

• The solution will become cloudy or flocculent.

16. Centrifuge on high speed for 10 minutes.

• Separation

17. Remove the tubes from the microcentrifuge, being careful not to disturb the white pellet.

18. Using P-1000 micropipettor set to 850 ul, carefully transfer the supernatant from each centrifuge tube to the corresponding blue spin column.

19. Centrifuge the spin columns for 30 seconds at maximum speed.

• Don't forget to put the lid on the rotor! Some of the salts get aerosolized because the spin columns don't have caps.

20. Pour the flow-through from each column into the miniprep waste container.

21. Pipette 500 ul of Buffer PB onto each spin column.

22. Centrifuge the spin columns for 30 seconds at maximum speed.

23. Pour the flow-through from each column into the miniprep waste container.

24. Pipette 750 ul of Buffer PE onto each spin column

25. Wait 1-3 minutes.

• This allows some of the salt that's still bound to the silica matrix to resuspend in the buffer.

26. Centrifuge the spin columns for 30 seconds at maximum speed.

27. Pour the flow-through from each column into the miniprep waste container.

28. Return each spin column to its collection tube and centrifuge an additional 1 minute at high speed.

• This removes every last trace of buffer PE; the ethanol can screw up downstream steps.

29. Transfer each spin column to a clean labelled microcentrifuge tube.

30. Pipette 50 ul of Buffer EB onto the center of each column.

• The volume of EB is comparable to the volume of silica gel matrix; if you pipette down the side, you might not get the entire transfer to the matrix.

31. Wait 1-3 minutes.

• This gives the DNA a chance to dissociate from the silica matrix.

32. Centrifuge the spin columns, in their collection tubes, for one minute at maximum speed.

33. Proceed directly to analyze the samples on the Nanodrop.

Pouring an agarose gel

Introduction

Agarose gels are used to separate DNA fragments. Their most common use is

Materials

• TAE buffer, 50-100 ml
  • There is generally 1X TAE buffer at the iGEM bench in a 1L or 2L bottle.
  • If not, mix some more up from the carboy 10X stock by the main lab gel station. (Use the 1L graduated cylinder at the gel station.)
• UltraPure agarose
• Casting tray, casting stand, combs

Procedure

• Mix and melt the agarose

1. Check to see if there is a gel waiting in the fridge.

2. Determine how much molten agarose you'll need.

• A small gel cast takes 50 ml; a large gel cast is 100 ml.

3. Measure out the appropriate amount of TAE into a glass bottle or flask.

4. Add 5 µl SYBRSafe (0.5X ; stock solution is 10000X) per 100 ml of the solution and mix well.

5. Add UltraPure agarose to a final concentration of 1% (mass / volume)

• So, if you're making 50 ml gel: 0.5 g.

6. Swirl the bottle or flask to distribute the agarose.

7. Heat the solution in the microwave with frequent stirring to dissolve the agarose homogenously. ~30 seconds/100ml solution

8. Let sit until cool enough to handle.

• The agarose MUST cool some -- if it's too hot, it can warp the casting trays.

• Don't allow it to get too cold!

• PAUSE POINT - You can put the flask in the 55°C water bath almost indefinitely at this point.

• Pour the gel

9. Set up the gel tray in the casting stand. Make sure the rubber gaskets are flat up against the edges of the casting tray.

10. Set up the gel co mbs to form the wells.

• Rinse the combs with water and wipe dry.

• Note for combs: 15-well combs hold about 6 ul liquid per well, 12-well combs hold about 15 ul per well, 8-well combs hold about 20 ul per well

• Taping two 8-well comb wells together results in a well that holds up to 100 ul

• Taping three 8-well comb wells together result in a well that holds up to 200 ul

11. Pour the molten agarose into the casting tray.

• If bubbles form around the combs, remove and re-insert.

12. Wait 30 mintues for the gel to solidify.

13. Use immediately, or place in a plastic zip-lock baggie with a little 1X TAE and store at 4°C.

Golden Gate Assembly

Introduction

Golden Gate is an assembly method using Type II-S restriction enzymes and predefined overlap sequences to assemble multiple pieces of DNA in one reaction. Golden Gate design is beyond the scope of this protocol; this protocol is the reaction setup only.

Materials

• 200 µl thin-wall PCR strip tubes, one per reaction
• Backbone plasmid
  • Usually pDONR-GG L1_lacZ_L2 or pDONR-GG L4_lacZ_R1
• Parts being assembled, either in plasmids or gBlocks
• Nuclease-free water
• T4 ligase buffer, 1.5 µl per reaction
  • Found in single-use aliquots of 10 (??) µl in the -20° freezer. T4 ligase buffer does NOT like to be thawed and frozen again; once you use an aliquot, discard what's left!
• 10X BSA, 1.5 µl per reaction
  • Found in the -20° freezer. ...and that's Bovine Serum Albumin, NOT the BsaI enzyme!
• BsaI (or AarI) restriction enzyme, 1 µl per reaction
  • Leave the BsaI and T4 ligase in the freezer until the very end, and add after the rest of the reaction has been set up.
• T4 ligase, 1 µl per reaction
  • Many protocols (both online and in the lab) call for high-concentration T4 ligase. This is only necessary if you're doing VERY LARGE Golden Gate reactions (5-8 parts). You do NOT need the HC ligase for most of the GGs you'll be doing.
  • Leave the BsaI and the T4 ligase in the freezer until the very end, and add after the rest of the reaction has been set up.
• An empty 10µl or 200µl tip box
• Ice in an ice bucket

Procedure

• Record your reaction setup.

1. A Golden Gate works best when there are equimolar amounts of each part in the reaction. By convention, we use 50 fmol of each part. For each part (plasmid and gBlock), compute the concentration (fmol/ul) and the volume required for 50 fmol in the table below. If you resuspended your gBlocks at 50 fmol/ul, you can omit them in this table.

	A	B	C	D	E
1	Part	Size (kb)	Concentration, (ng/ul)	Concentration, (fmol/ul)	Volume for 50 fmol
2	pDONR-GG	3.4	100	44.6	1.1
3
4
5

Table1

2. Record the reaction setup for each reaction below. For each reaction, include:

• -- 50 fmol of each part

• -- 1.5 ul T4 ligase buffer

• -- 1.5 ul 10X BSA

• -- 1 ul BsaI

• -- 1 ul T4 ligase

• -- Water to a total volume of 15 ul.

• -- NOTE: If you're using AarI, you need to add the AarI oligo as well!

	A	B	C	D
1	A19 L4_PhlF_R1
2	pDONR-GG L1_L2	1.1 ul
3	BT-01 gBlock	1 ul
4	BT-02 gBlock	1 ul
5	Water	6.9 ul
6	T4 ligase buffer	1.5 ul
7	10X BSA	1.5 ul
8	BsaI or AarI	1 ul
9	T4 ligase	1 ul

Table2

• Set up your thermocycler

3. Program a thermocycler with the following program:

• NOTE: If you are using AarI, use 50 cycles instead of 15.

	A	B
1	Heat lid	70°
2	Start cycle	15X
3	--	37° for 1' 30"
4	--	16° for 3' 0"
5	Close Cycle
6	50° for 5'
7	80° for 10'
8	Store at 8°

Table3

• Set up your reactions

4. Fill an empty tip box with ice, put the grey platform back on, then fill with water until just under the rack.

5. Label one thin-walled PCR tube per reaction.

6. Add the reaction components. Add the water first; add the restriction enzyme and buffer last.

7. Cap the tubes. Flick a few times to mix, then pulse spin.

8. Load the tubes in the thermocycler and start the program.

• Transform

9. Transform as usual. Plate only 10 ul of the outgrowth in a 200 ul puddle of water, or you will get a lawn of colonies.

Resuspending gBlocks

Introduction

gBlocks are long double-stranded linear pieces of DNA that arrive from IDT dehydrated; to use them, you need to resuspend them in TE to a known concentration. They are synthesized on a much smaller scale than primers and other oligonucleotides; we generally resuspend to a concentration of 50 fmol/µl.

Materials

• Nuclease-free TE
  • We resuspend in TE to help supress nuclease activity that would degrade the gBlocks
• Dehydrated gBlocks from IDT

Procedure

• Resuspend the dried gBlock

1. Label the top of the gBlock tube. I recommend the group initials and a number.

2. In the little microfuge, spin the (dry) gBlocks briefly.

• Sometimes the freeze-dried gBlocks flake off the bottom of the tube.

3. Determine how many fmol of gBlock are in the tube.

• The number of fmol is on the bottom line of the label. For example, one of my gBlocks says

• "500 ng = 1879 fmol"

4. Resuspend in the appropriate amount of TE.

• For my gBlock, if there are 1879 fmol in the tube, and I want 50 fmol/ul, then I need to resuspend in 37.5 ul of TE.

5. Vortex briefly. Pulse spin.

Annealing Oligos

Introduction

Anealing Oligos

Materials

• Nuclease-free TE
  • We resuspend in TE to help supress nuclease activity that would degrade the primers.
• Annealing buffer: TE + 50 mM NaCl
• Dehydrated oligos from IDT
• One extra epi tube per oligo
• One thin-walled 200 ul PCR tube

Procedure

• Preparing stock solution

1. Briefly spin each tube BEFORE opening

2. Add sterile 1X TE (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) in a volume of uL equal to 10 times the nmol amount listed

• Concentration of stock solution is 100 uM

3. Store at -20 ^oC

• Procedure

4. Set a thermocycler to a block temperature of 95°C and a lid temperature of 105°C.

5. Dilute oligo stock solutions to 5 uM working stock (95 uL TE + 5 uL of 100 uM stock)

6. Combine 98 uL of annealing buffer, and 1 uL of 5 uM forward primer, and 1 uL of 5 uM reverse primer in a thin-walled PCR tube. Mix well.

• The concentration for the anealed oligos is 50 nM (50 fmol/uL)

7. Place tube in a standard heatblock at 90–95 °C for 3–5 minutes.

8. Remove the heat block from the apparatus and allow to cool to room temperature (or at least below 30 °C) on the workbench. Slow cooling to room temperature should take 45–60 minutes.

9. Store on ice or at 4 °C until ready to use.

Diagnostic Restriction Digestion

Introduction

A diagnostic restriction digest helps identify correctly assembled clones from incorrect clones.

Materials

• 200 ul PCR strip tubes, one per reaction
• Restriction enzyme (chosen below), 1 ul per reaction.
• 10X restriction enzyme buffer (chosen below), 1 ul per reaction.

Procedure

• Choose A Good Restriction Enzyme

1. Using Benchling, choose a restriction enzyme that meets the following criteria:

• - Cuts at least once in the insert (for a pEXPR, either the promoter or the gene.)

• - Cuts at least once in the backbone.

• - Gives bands that aren't too large (> 8 kb) or too small (< 200 bp).

• - Gives a band pattern that is significantly different from the expected error mode.

• * For LR reactions, the most common error is a pDEST that slipped through the selection.

• - Begin in the "Brian's Favorites" list, then expand to the main Weiss lab list.

• - If you can't find a single enzyme that gives an acceptable band pattern, choose two enzymes that give a acceptable band pattern when used together.

• * This double-digest is subject to buffer compatibility, outlined below.

2. Record your enzyme choice on the plasmid's Description page.

3. Benchling will tell you the enzyme's buffer compatibility and active temperature. Record the buffer in which the enzyme is most active.

• If there are multiple bufers in which the enzyme is equally active, choose in this order: Buffer 3.1, Buffer CS, Buffer 2.1, Buffer 1.1.

• Set up the restriction digest

4. Retrieve the minipreps and the appropriate 10X buffer concentrate from the freezer. Thaw on the benchtop or in your fingers.

5. Label the PCR tubes with your initials and an incrementing number.

• ie: BT-1, -2, -3, -4

6. Vortex the minipreps and the 10X buffer concentrate briefly, then pulse down in the microfuge.

7. For each miniprep, set up a PCR tube containing the following in order:

• - 5 ul enzyme-quality H2O

• - 1 ul enzyme buffer

• - 3 ul miniprep DNA

• - 1 ul enzyme

• Remove the enzyme from the freezer for as little time as possible.

• I have specified an "arbitrary" 3 µl volume of miniprep DNA; this should be fine as long as your miniprep concentration is >= 100 ng/ul.

8. Flick the strip tubes a few times to mix the reaction, then pulse down in the strip tube microfuge.

9. Incubate at the appropriate temperature for at least 1 hour and more more than 16 hours.

• If the enzyme's active temperature is 37°C, use the 37°C plate incubator.

10. Stop the reaction by adding 2 ul of 6X NEB purple gel loading dye to each reacti on.

11. Flick the strip tubes a few times to mix the reaction, then pulse down in the strip tube microfuge.

• PAUSE POINT: The reaction can be stored almost indefinitely at room temperature once it's been stopped.

12. Proceed to gel electrophoresis.

Gel electrophoresis

Introduction

Gel electrophoresis separates pieces of DNA by length.

Materials

• Agarose gel, 1 lane per sample, plus at least 1 lane for the ladder.
  • If you just poured the gel, make sure it has had at least 30 minutes to set.
  • If you have a large gel but only need a few lanes, cut out just a piece of it with a razor blade. Cut straight!
• Samples to run
• 6X NEB Purple Loading Dye
• Molecular weight standard (aka "ladder")
  • Commonly available ladders are Hyperladder I and NEB 2-Log.
• Gel box, lid, leads
• Electrophoresis power supply
• 1X TAE buffer, enough to fill the gel box.

Procedure

• Prepare your samples

1. If your samples are frozen, thaw them completely, flick or vortex to mix, then pulse down in the microfuge.

2. If your samples are not already in loading dye, mix them 1:6 with 6X NEB Purple Loading Dye

• Add 1 µl of loading dye for every 5 µl of sample.

• Pro tip: If you don't want to mix loading gel with your entire sample (PCR products, for example), cut off a strip of Parafilm; pipette 2-3 ul of sample onto the parafilm; add 1 ul loading dye; mix by pipetting and load directly.

• Set up the gel box

3. Program the voltage on the power supply. For the small gel box, use 100V; for the large gel box, use 150V.

4. Program the time on the power supply. For a small analytical gel, set the timer for 30 minutes.

• For larger gels, start at 45 minutes, and then check regularly!

5. Attach the leads to the gel box cover. Make sure that the red (positive) lead is attached to the side of the box farthest away from the wells in the gel.

• Remember, the DNA is negatively charged and will move toward the positive terminal. A useful neumonic for remembering this is "Run to Red."

6. If the gel box has not been used previously that day, empty it and rinse it out with DI water.

• The TAE buffer can be re-used, but if it's been sitting out for too long it evaporates and the salt concentration (and conductivity) changes. TAE is cheap; when in doubt, replace it.

7. Place the gel in the gel box.

8. Pour TAE into the gel box until it just barely covers the gel.

• The TAE's purpose is to conduct electricity; over-filling the gel box results in a larger conductive path, more current flow, and more heating (which can screw up your run.)

9. Check the wells to see that they are free of bubbles. If there are bubbles, blow them out by pipetting 100 µl of TAE from the gel box into the well.

• Load your samples

10. Load your samples. For large combs, load 10 µl; for small combs, load 5 µl.

• Work quickly. The samples begin to diffuse in the buffer, leading to smeared bands.

11. Load the ladder in the last lane. Load 1/2 the volume of your samples: for large combs, load 5 µl, for small combs, load 2.5 µl.

12. Place the lid on the gel box. Make sure it is seated on the brass contacts.

• Run the gel

13. Press the Start or Run button on the power supply.

14. Make sure the power supply doesn't complain about an open circuit. If it does, re-seat the gel box lid and press Run again.

15. Double-check that there are bubbles forming on the platinum wires at either end of the gel box.

16. Double-check that the red (positive) lead is on the side farthest from the wells. Remember, Run to Red.

17. Run until the pink band is 2/3 to 3/4 of the way down the gel. For small analytical gels, this should take 30 minutes. For larger gels, start at 45 minutes and check regularly!

• Image the gel

18. If the power supply is still running, press the Stop button.

19. Lift the lid off of the gel box. Lift the lid straight up. If you try to "hinge" it up, the lid will break.

20. Transfer the gel to the GelDoc.

21. If it's not running, start QuantityOne from the toolbar.

• If necessary, click the top button on the toolbox to select the scanner.

• If necessary, reset the camera (as per instructions on the GelDoc computer.)

22. Press the Epifluorescent Illumination button on the GelDoc. Check the gel's position, zoom and focus.

• You want to be zoomed in so that the gel fills the field of view, and focussed so that the well edges are sharp.

23. Close the GelDoc door. Click Auto Expose.

• The Auto Expose functionality generally over-exposes my gels a bit. If your gel is over-exposed, remove 1/3 of the exposure time, type it into the Exposure box, then click Manually Expose.

24. In the File menu, select, Export as JPEG.... Save your gel to the iGEM folder on the desktop.

25. Open Benchling. Log in, and copy the gel to the Description page for the plasmid you're building.

• If there are multiple plasmids on the gel, save it to each plasmid's Description.

26. Discard the gel in the biowaste box. Wipe down the gel doc with a little water and a paper towel or Kimwipe.

• Annotate the gel

27. Immediately, before you forget what's where, annotate the gel.

• List what is in each lane.

• Describe whether the pattern is what you expected or not. (You should have an in silico digestion to compare it to!)

Transformation of E. coli

Introduction

Transformation is the process of inducing chemically competent E. coli to take up DNA.

Materials

• Dry bath, set to 42°C
  • Fill the wells in the dry bath block 1/2 full with DI water.
• Ice bucket, with ice
  • For thawing competent cells.
• DNA to transform
  • Could be an assembly reaction (LR, Golden Gate, etc) or a miniprepped plasmid.
  • If you removed it from the freezer, make sure it's entirely thawed out.
• pUC19 Transformation Control, 1 pg/µl
  • The pUC19 control will tell you how efficient your transformations were.
• SOC growth media, at room temperature
  • Check to make sure it's clear and NOT CLOUDY.
• Antibiotic plates, one per transformation, plus 1 Amp plate for the pUC19 control
  • Make sure the plates you use match the resistance cassette of the plasmid!
• Competent E. coli, one tube per transformation + one for the pUC19 control
  • These live in the -80 in 235.
  • Thaw on ice 3-4 minutes.
• A timer, set for 30 seconds.

Procedure

• Setup

1. Make sure the dry bath is set to 42°C and the wells in the block are 1/2 full of DI water

2. Remove selection plates from the refrigerator. Double-check that they match the selection marker on your plasmid, then place them in the 37° incubator.

3. Retrieve the DNA to transform.

• If frozen: thaw, completely, flick a few times to mix, then pulse down in the microfuge.

4. Fill an ice bucket with ice. Retrieve one tube of competent E. coli per transformation from the -80 and thaw on ice, 3-4 minutes.

00:03:00

5. While the transformation tubes are thawing, label their tops with something descriptive. Record the labels here:

	A	B	C	D	E	F	G	H
1	puc-19L
2	puc-19TC
3	Pdest_mCherry
4	pDest

Table1

• Transformation

6. Add 2 µl DNA from each reaction to a tube of competent cells.

• Immediately after adding the DNA to each tube, stir the cells a few times with the pipette tip.

7. Add 1 µl of the pUC19 transformation control to the positive control tube.

8. Incubate on ice for 30 minutes.

00:30:00

9. Heat shock the cells for exactly 30 seconds in the 42° heat block. (Yes, set a timer.)

00:00:30

10. Place back on ice for 2 minutes.

00:02:00

11. Add 250 µl SOC to each tube.

12. Tape the tubes to the platform of a shaker at 37°C and shake at 270 RPM for 60 minutes.

01:00:00

• Plating

13. Label the selection plates using the labels you recorded above.

14. Shake ~10 plating beads onto each plate.

15. Pipette 100 µl of each transformation onto the corresponding plates. NOTE: if you are using DNA from a golden gate reaction, see the golden gate protocol for instructions to dilute your sample.

16. -----Golden Gate: Plate only 10 ul of the outgrowth in a 200 ul puddle of water, or you will get a lawn of colonies.

17. Cover the plates and shake the beads around to spread the cells out.

18. Dispose of the beads by tapping them into the waste container.

19. Incubate the plates upside down overnight in the 37° incubator.

• Don't incubate for more than 18-24 hours.

• Compute transformation efficiency

20. Count the colonies on your positive transformation plate.

• If there are many many colonies, then hooray! You had a great transformation. Just estimate.

21. Divide the number of colonies by the fraction of the transformation you plated.

• So, if you resuspended your transformation in a total volume of 300 ul, then plated 100 ul, multiply the number of colonies by 3.

22. Transformation efficiency is expressed in colonies per microgram pUC19. Multiply the number of colonies by the appropriate conversion factor.

• So if you transformed 1 picogram of pUC19 DNA, multiply by 10⁶.

23. Record your transformation efficiency in your (daily) lab notebook.

Glycerol Stock

Introduction

While minipreping, remaining miniprep culture should be have been kept in the 4 degree C fridge. After sequencing and confirmation, the miniprep culure of the cells you've chosen to midiprep should be made into a glycerol stock and put into the -80 iGEM glycerol box.

Materials

• Cryo Tubes (orange caps, in bottom drawer)
• 50% Glycerol solution (on shelves above bench) 500uL (for each miniprep culture)
• Miniprep 500uL
• Label Printer and other components
• P1000 and tips

Procedure

• Initial Set Up

1. First make sure that the tape printer located on top of the computer tower has enough tape. If out of tape, contact Brian for more.

2. Take our your miniprep cultures and aspirate the ones that you will not be using for midiprep

3. Check that you have enough glycerol. If out, contact Brian.

4. Take out the number of cryo tubes equal to the amount of minipreps cultures you'll be using for midiprep. If out, check the Weiss Lab cabinets located in hall.

• Labels

5. Labels should be made first before making the glycerol stock so you don't confuse tubes.

6. The label making program, P-touch Editor 5.0, can be found on the desktop of the computer

7. If the iGEM label template is not open, click File--->Open---->iGEM 2016 Template Label

8. Follow template of label. Fill in date, your initiails, plasmid name, cell type, and add 'iGEM 2016'

9. Once finished, print the labels using the printer named Brother PT-2430PC

10. The labels are peel stickers. Stick labels onto cryo tubes

• Making Glycerol Stock

11. Pipette 500uL glycerol into each cryo tube.

12. Pipette 500ul of each miniprep culture into their respective cryo tubes. Remember to switch pipette tips each time

13. Cap tubes are store in -80 degree freezer in iGEM glycerol stock box. Should be located in the bottom of the freezer.

14. Store remaining miniprep cultures in 4 degree fridge until used to grow midiprep culture

TN 07/21/16

Made with Benchling

Project: iGem 2016 _ Recombinase subgroup

Authors: Elizabeth Strand

Date: 2016-07-21

Thursday, 7/21

Miniprep Concentrations - hef1a-SV40-eYFP:

	A	B
1	1-1	226.3
2	1-2	216.8
3	1-3	230.3
4	1-4	231.3

Table1

clipboard_2016-07-21_10:38:32.png

RecombinaseGel 7-21-16.jpg

Sample Preparation for Flow Cytometry

Introduction

Materials

• Versene
  • Clear liquid on Brian's side of drawer
• Trypsin
  • Remember to put Trypsin and cell media into dry bath to warm them up before starting
• HEK cell media
• Flow cytometry tubes
  • Tubes with Blue filters on top
  • One for each well
• Ice Bucket
• Glass pipette tips
• P200 and P1000 pipette tips

Procedure

• Preparing Samples

1. Retrieve cells from incubator

2. Look at the wells under the EVOS microscope by the Lu Bench to check the cells and look for fluorescence

3. For each well:

• - Aspirate the media (make sure the tip does not touch the bottom of the well)

• - Pipette 200 uL of versene slowly down the side of the well

4. Again, for each well:

• - Aspirate the versene

• - Pipette 200 uL of trypsin and wait three minutes

5. Add 500 uL of media to each well

6. Label flow cytometry tubes and put in an ice bucket

7. Pipette up and down in circles, and then add entire mixture of media and trypsin to a flow cytometry tube through the filter

8. Centrifuge at 2300 rpm for 4 min

9. Aspirate out the supernatant for each tube without disturbing the cell pellet

• - Pellet is very small and hard to see

• - Don't put tip lower than the line around the bottom of the tube so that you don't disturb the pellet

10. Add 500 uL of versene to each tube (making sure to avoid bubbles, as it can clog the cytometer)

11. Go find Brian, and he will load your samples into the cytometer

Sequencing protocol

Introduction

Preparing sample to send to Genewiz for sequencing Sequencing Pick-up Times: 9:30am, 3:00 pm, 5:30 pm

Materials

• DNA stock
• Nuclease-free water
• Primer (5nM stock)
• One microcentrifuge tube for each DNA plasmid
• 2 PCR tubes per plasmid (must be in strips of eight, round up if needed!)

Procedure

• Diluting the DNA

1. For orders with <48 samples, use 8-strip PCR tubes. Label your tubes on the side vertically with "MI" and sample number (01, 02, 03, etc.). Use image below as guideline. Each plasmid you want to sequence will require 2 tubes, one for the forward primer and one for the reverse primer. Use "MI" instead of the "GW" in the picture.

2. Dilute your sequencing primer to 5 µM (pmol/µl) using water. You will need 5 µl for each sequencing reaction. Note, the primers in the tubes from IDT have not been diluted! Use already diluted stock (should be in regular microcentrifuge tubes).

3. Using the recorded concentration, dilute the DNA to the template concentration in 10 µl as detailed below. For example, a 5 kb plasmid would need a template concentration of ~50 ng/µl. Final volume for your working stock should 30 uL. Please make dilutions in water or Tris.

4. Add 10 µl of diluted template DNA to each tube.

5. Add 5 µl of diluted primer to each tube. Be careful about putting the forward and reverse primers in the correct tubes! Remember, for each DNA template, there should be two tubes - one with reverse primer and the other with forward primer

6. Vortex briefly

7. Log into genewiz. Username: igem-sequence@mit.edu Password: igem2014citrus

8. Click "create sequencing order"

9. Sevice Priority: click on "standard"

10. Create Order by: click on "online form"

11. Sample Type: click on "Pre-mixed"

12. Enter in how many samples you are sending in and click "Create new form"

13. Enter in descriptive names for your DNA (this is to help you remember when you get the results back), the DNA type, the DNA length (get this from your construct on benchling), the primer name (again, this is more for you). When you are done hit save and next.

14. It will give you an list of what you have entered and a price for each reaction. Hit "next step"

15. The payment information field should be autofilled. There has been an issue recently with the PO box number not being entered. If this is the case, click "credit card" (next to "payment info") and then click "PO" to bring you back to the original form. Now you should be able to enter in a PO number. Open up a previous submitted order (there should be some saved to the desktop called "sequencing" or something like that) and copy over the PO number. PO Number: 5510061907

16. Hit "next step". Once it gives you a submitted order form, print 2 copies and pick them up from the printer (exit lab, through glass doors, on your left)

17. Put your PCR tubes in a large falcon tube.

18. Put the falcon tube in a ziploc bag (located in Qiagen column drawer) and put in one order form. Label the bag with something descriptive ("Weiss Lab MIT iGEM Recombinase 6/30/16" for example)

19. Put the bag in the Genewiz pickup box (located by the elevators, make sure you put in the genewiz box)

20. Open up the weiss lab orders spreadsheet (bookmarked on the computer). You need to request access to the spreadsheet if it is your first time ordering. Once you request access, email Brian so he knows that you have sent in a request.

21. Click the tab at the bottom called "genewiz".

22. Fill out the form. Make sure you indicate that the order is from iGEM so the right account can be charged.

23. You did it! YAY! Genewiz should get back to you pretty quickly, generally between 12-24 hrs.

Midiprep

Introduction

Purifying large amount of DNA to tranfect mammalian cells

Materials

• Qiagen MidiPrep

Procedure

Miniprep

Introduction

The miniprep uses silica gel to isolate plasmid DNA from an E. coli culture

Materials

• Buffer P1 (resuspension buffer)
  • Retrieve from refrigerator. If you are opening a new miniprep kit, add the RNAse and LyseBlue reagent and check the box on the cap.
• Buffer P2 (lysis buffer)
  • Open the cap and look at the lysis buffer. Swirl it around. If it appears cloudy, the SDS has fallen out of solution; warm it for a few minutes in the 55°C water bath.
• Buffer N3 (neutralization buffer)
• Buffer PB (binding buffer)
• Buffer PE (rinse buffer)
  • Make sure the "Ethanol added?" box has been checked. If you are opening a new miniprep kit, add absolute ethanol as per the kit instructions and check the box on the cap.
• Buffer EB (elution buffer)
• Miniprep waste container
  • Miniprep buffers contain salts that can't go down the sink.
• Per miniprep: two microcentrifuge tubes and one blue spin column, with collection tube.

Procedure

• Harvest and resuspension

1. For each culture, label two microcentrifuge tubes on the cap and one blue spin column on the side.

• The spin columns should be in their (cap-less) collection vials.

2. Pipette 1.6 mL of each culture into the corresponding microcentrifuge tubes.

• (This is for a 1.7 ml microcentrifuge tube; set your pipettor to 800 ul and do two transfers. If we ever go back to 2-ml tubes, do 1.9 ml (2x950 ul))

3. Centrifuge at maximum speed (10,000 or 13,000xg) for three minutes.

5. Aspirate the supernatant, or pour it off into the bleach bucket.

6. Pipette ANOTHER 1.8 ml of each culture into the corresponding microcentrifuge tubes.

7. Centrifuge at maximum speed for three minutes.

8. While the centrifuge is running, move the remaining cultures to 4degC.

9. Aspirate the supernatant off with the bench aspirator. Be careful not to disturb the pelleted E. coli.

• We use an aspirator here because the less extra salt and protein we put in the miniprep, the better the yield is.

10. Add 250 µl Buffer P1 to each tube.

11. Resuspend the E. coli pellet. The preferred way is with the roto-mixer at the other end of the lab.

• Alternately, if you have just a few tubes, you can resuspend on a vortex.

• Make sure to resuspend fully and thoroughly. The resulting suspension should be smooth and cloudy; if there is particulate matter floating around, vortex some more.

• Lysis

12. Add 250 µl Buffer P2 to each tube.

• Work quickly; the lysis step should take less than 5 minutes.

13. Snap the tubes closed and invert them 4-6 times, until the tube is thoroughly mixed and the entire solution turns blue.

• If you have many many tubes, you can stack a second tube rack on top of them and invert the entire thing.

14. Add 350 ul Buffer N3 to each tube.

15. Snap the tubes closed and invert 4-6 times, until the solution is thouroughly mixed and no longer blue.

• The solution will become cloudy or flocculent.

16. Centrifuge on high speed for 10 minutes.

• Separation

17. Remove the tubes from the microcentrifuge, being careful not to disturb the white pellet.

18. Using P-1000 micropipettor set to 850 ul, carefully transfer the supernatant from each centrifuge tube to the corresponding blue spin column.

19. Centrifuge the spin columns for 30 seconds at maximum speed.

• Don't forget to put the lid on the rotor! Some of the salts get aerosolized because the spin columns don't have caps.

20. Pour the flow-through from each column into the miniprep waste container.

21. Pipette 500 ul of Buffer PB onto each spin column.

22. Centrifuge the spin columns for 30 seconds at maximum speed.

23. Pour the flow-through from each column into the miniprep waste container.

24. Pipette 750 ul of Buffer PE onto each spin column

25. Wait 1-3 minutes.

• This allows some of the salt that's still bound to the silica matrix to resuspend in the buffer.

26. Centrifuge the spin columns for 30 seconds at maximum speed.

27. Pour the flow-through from each column into the miniprep waste container.

28. Return each spin column to its collection tube and centrifuge an additional 1 minute at high speed.

• This removes every last trace of buffer PE; the ethanol can screw up downstream steps.

29. Transfer each spin column to a clean labelled microcentrifuge tube.

30. Pipette 50 ul of Buffer EB onto the center of each column.

• The volume of EB is comparable to the volume of silica gel matrix; if you pipette down the side, you might not get the entire transfer to the matrix.

31. Wait 1-3 minutes.

• This gives the DNA a chance to dissociate from the silica matrix.

32. Centrifuge the spin columns, in their collection tubes, for one minute at maximum speed.

33. Proceed directly to analyze the samples on the Nanodrop.

Diagnostic Restriction Digestion

Introduction

A diagnostic restriction digest helps identify correctly assembled clones from incorrect clones.

Materials

• 200 ul PCR strip tubes, one per reaction
• Restriction enzyme (chosen below), 1 ul per reaction.
• 10X restriction enzyme buffer (chosen below), 1 ul per reaction.

Procedure

• Choose A Good Restriction Enzyme

1. Using Benchling, choose a restriction enzyme that meets the following criteria:

• - Cuts at least once in the insert (for a pEXPR, either the promoter or the gene.)

• - Cuts at least once in the backbone.

• - Gives bands that aren't too large (> 8 kb) or too small (< 200 bp).

• - Gives a band pattern that is significantly different from the expected error mode.

• * For LR reactions, the most common error is a pDEST that slipped through the selection.

• - Begin in the "Brian's Favorites" list, then expand to the main Weiss lab list.

• - If you can't find a single enzyme that gives an acceptable band pattern, choose two enzymes that give a acceptable band pattern when used together.

• * This double-digest is subject to buffer compatibility, outlined below.

2. Record your enzyme choice on the plasmid's Description page.

3. Benchling will tell you the enzyme's buffer compatibility and active temperature. Record the buffer in which the enzyme is most active.

• If there are multiple bufers in which the enzyme is equally active, choose in this order: Buffer 3.1, Buffer CS, Buffer 2.1, Buffer 1.1.

• Set up the restriction digest

4. Retrieve the minipreps and the appropriate 10X buffer concentrate from the freezer. Thaw on the benchtop or in your fingers.

5. Label the PCR tubes with your initials and an incrementing number.

• ie: BT-1, -2, -3, -4

6. Vortex the minipreps and the 10X buffer concentrate briefly, then pulse down in the microfuge.

7. For each miniprep, set up a PCR tube containing the following in order:

• - 5 ul enzyme-quality H2O

• - 1 ul enzyme buffer

• - 3 ul miniprep DNA

• - 1 ul enzyme

• Remove the enzyme from the freezer for as little time as possible.

• I have specified an "arbitrary" 3 µl volume of miniprep DNA; this should be fine as long as your miniprep concentration is >= 100 ng/ul.

8. Flick the strip tubes a few times to mix the reaction, then pulse down in the strip tube microfuge.

9. Incubate at the appropriate temperature for at least 1 hour and more more than 16 hours.

• If the enzyme's active temperature is 37°C, use the 37°C plate incubator.

10. Stop the reaction by adding 2 ul of 6X NEB purple gel loading dye to each reacti on.

11. Flick the strip tubes a few times to mix the reaction, then pulse down in the strip tube microfuge.

• PAUSE POINT: The reaction can be stored almost indefinitely at room temperature once it's been stopped.

12. Proceed to gel electrophoresis.

Transformation of E. coli

Introduction

Transformation is the process of inducing chemically competent E. coli to take up DNA.

Materials

• Dry bath, set to 42°C
  • Fill the wells in the dry bath block 1/2 full with DI water.
• Ice bucket, with ice
  • For thawing competent cells.
• DNA to transform
  • Could be an assembly reaction (LR, Golden Gate, etc) or a miniprepped plasmid.
  • If you removed it from the freezer, make sure it's entirely thawed out.
• pUC19 Transformation Control, 1 pg/µl
  • The pUC19 control will tell you how efficient your transformations were.
• SOC growth media, at room temperature
  • Check to make sure it's clear and NOT CLOUDY.
• Antibiotic plates, one per transformation, plus 1 Amp plate for the pUC19 control
  • Make sure the plates you use match the resistance cassette of the plasmid!
• Competent E. coli, one tube per transformation + one for the pUC19 control
  • These live in the -80 in 235.
  • Thaw on ice 3-4 minutes.
• A timer, set for 30 seconds.

Procedure

• Setup

1. Make sure the dry bath is set to 42°C and the wells in the block are 1/2 full of DI water

2. Remove selection plates from the refrigerator. Double-check that they match the selection marker on your plasmid, then place them in the 37° incubator.

3. Retrieve the DNA to transform.

• If frozen: thaw, completely, flick a few times to mix, then pulse down in the microfuge.

4. Fill an ice bucket with ice. Retrieve one tube of competent E. coli per transformation from the -80 and thaw on ice, 3-4 minutes.

00:03:00

5. While the transformation tubes are thawing, label their tops with something descriptive. Record the labels here:

	A	B	C	D	E	F	G	H
1	puc-19L
2	puc-19TC
3	Pdest_mCherry
4	pDest

Table1

• Transformation

6. Add 2 µl DNA from each reaction to a tube of competent cells.

• Immediately after adding the DNA to each tube, stir the cells a few times with the pipette tip.

7. Add 1 µl of the pUC19 transformation control to the positive control tube.

8. Incubate on ice for 30 minutes.

00:30:00

9. Heat shock the cells for exactly 30 seconds in the 42° heat block. (Yes, set a timer.)

00:00:30

10. Place back on ice for 2 minutes.

00:02:00

11. Add 250 µl SOC to each tube.

12. Tape the tubes to the platform of a shaker at 37°C and shake at 270 RPM for 60 minutes.

01:00:00

• Plating

13. Label the selection plates using the labels you recorded above.

14. Shake ~10 plating beads onto each plate.

15. Pipette 100 µl of each transformation onto the corresponding plates. NOTE: if you are using DNA from a golden gate reaction, see the golden gate protocol for instructions to dilute your sample.

16. -----Golden Gate: Plate only 10 ul of the outgrowth in a 200 ul puddle of water, or you will get a lawn of colonies.

17. Cover the plates and shake the beads around to spread the cells out.

18. Dispose of the beads by tapping them into the waste container.

19. Incubate the plates upside down overnight in the 37° incubator.

• Don't incubate for more than 18-24 hours.

• Compute transformation efficiency

20. Count the colonies on your positive transformation plate.

• If there are many many colonies, then hooray! You had a great transformation. Just estimate.

21. Divide the number of colonies by the fraction of the transformation you plated.

• So, if you resuspended your transformation in a total volume of 300 ul, then plated 100 ul, multiply the number of colonies by 3.

22. Transformation efficiency is expressed in colonies per microgram pUC19. Multiply the number of colonies by the appropriate conversion factor.

• So if you transformed 1 picogram of pUC19 DNA, multiply by 10⁶.

23. Record your transformation efficiency in your (daily) lab notebook.

Overnight liquid cultures (picking colonies)

Introduction

Overnight cultures are used to prepare miniprep DNA.

Materials

• The plate from which you are picking colonies
• 15 ml round-bottom polystyrene tubes, one per culture
  • The ones with the snap caps, NOT conical tubes with screw caps
• 5 mL LB per culture
• A container that can hold 5ml x the number of cultures
  • For a modest number of minipreps, a 50 ml conical tube works well.
  • For larger minipreps, use a sterile bottle (100 ml is frequently useful.)
• Antibiotic stock, 1000X

Procedure

• Materials Setup

1. Warm up the LB to at least room temperature (if it came from the fridge), but not warmer than 37°C

2. Label one round-bottom culture tube for each miniprep. Use "NAME-1, NAME-2, ..."etc for the naming convention, where NAME is a shortened name of the plasmid (eg, "hEF1a:mKate").

• Your impulse is to just use number, or initials and number, but trust me -- you will want to be able to identify this tube in three weeks when you've forgotten what you were doing.

3. Using a sterile pipette, transfer 5 ml of LB to the mixing container for each culture PLUS 5 ML.

4. Add antibiotic stock to a final concentration of 1X (1 µl stock for each 1 ml in the mixing container.)

5. Cap tightly and mix well.

• Culture Setup

6. Using a sterile pipette, transfer 5 ml of LB+antibiotic to each round-bottom culture tube.

• If you are making cultures with different antibiotics, take care that the right media goes in each tube.

7. Squirt ethanol on a pair of foreceps and wipe dry with a Kimwipe.

8. Use the foreceps to pick up a sterile 200µl pipette tip, scrape a colony off of the plate, and drop the pipette tip in the corresponding tube.

• Repeat for each tube.

9. Transfer to an incubating shaker at 37°C and incubate 14-16 hours.

• Don't over-grow too badly, or your yield will suffer.

• If you need to grow longer, you can grow at 30°C instead for 20 hours.

JG 7/22/16

Made with Benchling

Project: iGem 2016 _ Recombinase subgroup

Authors: Elizabeth Strand

Date: 2016-07-22

Friday, 7/22

clipboard_2016-07-22_15:20:46.png

RecombinaseTOP.jpg

RecombinaseGelBOTTOM.jpg

Miniprep Concentrations:

1 = hef1a-SloxP-SV40

3= attb-SV40-attp

4 = TP901

	A	B
1	1-1	415.5
2	1-2	386.5
3	1-3	426.3
4	1-4	304.5
5	3-1	328.8
6	3-2	244.5
7	3-3	346.4
8	3-4	267.5
9	4-1	212.2
10	4-2	279.5
11	4-3	285.6
12	4-4	334.2

Table1

Diagnostic Restriction Digestion

Introduction

A diagnostic restriction digest helps identify correctly assembled clones from incorrect clones.

Materials

• 200 ul PCR strip tubes, one per reaction
• Restriction enzyme (chosen below), 1 ul per reaction.
• 10X restriction enzyme buffer (chosen below), 1 ul per reaction.

Procedure

• Choose A Good Restriction Enzyme

1. Using Benchling, choose a restriction enzyme that meets the following criteria:

• - Cuts at least once in the insert (for a pEXPR, either the promoter or the gene.)

• - Cuts at least once in the backbone.

• - Gives bands that aren't too large (> 8 kb) or too small (< 200 bp).

• - Gives a band pattern that is significantly different from the expected error mode.

• * For LR reactions, the most common error is a pDEST that slipped through the selection.

• - Begin in the "Brian's Favorites" list, then expand to the main Weiss lab list.

• - If you can't find a single enzyme that gives an acceptable band pattern, choose two enzymes that give a acceptable band pattern when used together.

• * This double-digest is subject to buffer compatibility, outlined below.

2. Record your enzyme choice on the plasmid's Description page.

3. Benchling will tell you the enzyme's buffer compatibility and active temperature. Record the buffer in which the enzyme is most active.

• If there are multiple bufers in which the enzyme is equally active, choose in this order: Buffer 3.1, Buffer CS, Buffer 2.1, Buffer 1.1.

• Set up the restriction digest

4. Retrieve the minipreps and the appropriate 10X buffer concentrate from the freezer. Thaw on the benchtop or in your fingers.

5. Label the PCR tubes with your initials and an incrementing number.

• ie: BT-1, -2, -3, -4

6. Vortex the minipreps and the 10X buffer concentrate briefly, then pulse down in the microfuge.

7. For each miniprep, set up a PCR tube containing the following in order:

• - 5 ul enzyme-quality H2O

• - 1 ul enzyme buffer

• - 3 ul miniprep DNA

• - 1 ul enzyme

• Remove the enzyme from the freezer for as little time as possible.

• I have specified an "arbitrary" 3 µl volume of miniprep DNA; this should be fine as long as your miniprep concentration is >= 100 ng/ul.

8. Flick the strip tubes a few times to mix the reaction, then pulse down in the strip tube microfuge.

9. Incubate at the appropriate temperature for at least 1 hour and more more than 16 hours.

• If the enzyme's active temperature is 37°C, use the 37°C plate incubator.

10. Stop the reaction by adding 2 ul of 6X NEB purple gel loading dye to each reacti on.

11. Flick the strip tubes a few times to mix the reaction, then pulse down in the strip tube microfuge.

• PAUSE POINT: The reaction can be stored almost indefinitely at room temperature once it's been stopped.

12. Proceed to gel electrophoresis.

Gel electrophoresis

Introduction

Gel electrophoresis separates pieces of DNA by length.

Materials

• Agarose gel, 1 lane per sample, plus at least 1 lane for the ladder.
  • If you just poured the gel, make sure it has had at least 30 minutes to set.
  • If you have a large gel but only need a few lanes, cut out just a piece of it with a razor blade. Cut straight!
• Samples to run
• 6X NEB Purple Loading Dye
• Molecular weight standard (aka "ladder")
  • Commonly available ladders are Hyperladder I and NEB 2-Log.
• Gel box, lid, leads
• Electrophoresis power supply
• 1X TAE buffer, enough to fill the gel box.

Procedure

• Prepare your samples

1. If your samples are frozen, thaw them completely, flick or vortex to mix, then pulse down in the microfuge.

2. If your samples are not already in loading dye, mix them 1:6 with 6X NEB Purple Loading Dye

• Add 1 µl of loading dye for every 5 µl of sample.

• Pro tip: If you don't want to mix loading gel with your entire sample (PCR products, for example), cut off a strip of Parafilm; pipette 2-3 ul of sample onto the parafilm; add 1 ul loading dye; mix by pipetting and load directly.

• Set up the gel box

3. Program the voltage on the power supply. For the small gel box, use 100V; for the large gel box, use 150V.

4. Program the time on the power supply. For a small analytical gel, set the timer for 30 minutes.

• For larger gels, start at 45 minutes, and then check regularly!

5. Attach the leads to the gel box cover. Make sure that the red (positive) lead is attached to the side of the box farthest away from the wells in the gel.

• Remember, the DNA is negatively charged and will move toward the positive terminal. A useful neumonic for remembering this is "Run to Red."

6. If the gel box has not been used previously that day, empty it and rinse it out with DI water.

• The TAE buffer can be re-used, but if it's been sitting out for too long it evaporates and the salt concentration (and conductivity) changes. TAE is cheap; when in doubt, replace it.

7. Place the gel in the gel box.

8. Pour TAE into the gel box until it just barely covers the gel.

• The TAE's purpose is to conduct electricity; over-filling the gel box results in a larger conductive path, more current flow, and more heating (which can screw up your run.)

9. Check the wells to see that they are free of bubbles. If there are bubbles, blow them out by pipetting 100 µl of TAE from the gel box into the well.

• Load your samples

10. Load your samples. For large combs, load 10 µl; for small combs, load 5 µl.

• Work quickly. The samples begin to diffuse in the buffer, leading to smeared bands.

11. Load the ladder in the last lane. Load 1/2 the volume of your samples: for large combs, load 5 µl, for small combs, load 2.5 µl.

12. Place the lid on the gel box. Make sure it is seated on the brass contacts.

• Run the gel

13. Press the Start or Run button on the power supply.

14. Make sure the power supply doesn't complain about an open circuit. If it does, re-seat the gel box lid and press Run again.

15. Double-check that there are bubbles forming on the platinum wires at either end of the gel box.

16. Double-check that the red (positive) lead is on the side farthest from the wells. Remember, Run to Red.

17. Run until the pink band is 2/3 to 3/4 of the way down the gel. For small analytical gels, this should take 30 minutes. For larger gels, start at 45 minutes and check regularly!

• Image the gel

18. If the power supply is still running, press the Stop button.

19. Lift the lid off of the gel box. Lift the lid straight up. If you try to "hinge" it up, the lid will break.

20. Transfer the gel to the GelDoc.

21. If it's not running, start QuantityOne from the toolbar.

• If necessary, click the top button on the toolbox to select the scanner.

• If necessary, reset the camera (as per instructions on the GelDoc computer.)

22. Press the Epifluorescent Illumination button on the GelDoc. Check the gel's position, zoom and focus.

• You want to be zoomed in so that the gel fills the field of view, and focussed so that the well edges are sharp.

23. Close the GelDoc door. Click Auto Expose.

• The Auto Expose functionality generally over-exposes my gels a bit. If your gel is over-exposed, remove 1/3 of the exposure time, type it into the Exposure box, then click Manually Expose.

24. In the File menu, select, Export as JPEG.... Save your gel to the iGEM folder on the desktop.

25. Open Benchling. Log in, and copy the gel to the Description page for the plasmid you're building.

• If there are multiple plasmids on the gel, save it to each plasmid's Description.

26. Discard the gel in the biowaste box. Wipe down the gel doc with a little water and a paper towel or Kimwipe.

• Annotate the gel

27. Immediately, before you forget what's where, annotate the gel.

• List what is in each lane.

• Describe whether the pattern is what you expected or not. (You should have an in silico digestion to compare it to!)

Midiprep

Introduction

Purifying large amount of DNA to tranfect mammalian cells

Materials

• Qiagen MidiPrep

Procedure

Miniprep

Introduction

The miniprep uses silica gel to isolate plasmid DNA from an E. coli culture

Materials

• Buffer P1 (resuspension buffer)
  • Retrieve from refrigerator. If you are opening a new miniprep kit, add the RNAse and LyseBlue reagent and check the box on the cap.
• Buffer P2 (lysis buffer)
  • Open the cap and look at the lysis buffer. Swirl it around. If it appears cloudy, the SDS has fallen out of solution; warm it for a few minutes in the 55°C water bath.
• Buffer N3 (neutralization buffer)
• Buffer PB (binding buffer)
• Buffer PE (rinse buffer)
  • Make sure the "Ethanol added?" box has been checked. If you are opening a new miniprep kit, add absolute ethanol as per the kit instructions and check the box on the cap.
• Buffer EB (elution buffer)
• Miniprep waste container
  • Miniprep buffers contain salts that can't go down the sink.
• Per miniprep: two microcentrifuge tubes and one blue spin column, with collection tube.

Procedure

• Harvest and resuspension

1. For each culture, label two microcentrifuge tubes on the cap and one blue spin column on the side.

• The spin columns should be in their (cap-less) collection vials.

2. Pipette 1.6 mL of each culture into the corresponding microcentrifuge tubes.

• (This is for a 1.7 ml microcentrifuge tube; set your pipettor to 800 ul and do two transfers. If we ever go back to 2-ml tubes, do 1.9 ml (2x950 ul))

3. Centrifuge at maximum speed (10,000 or 13,000xg) for three minutes.

5. Aspirate the supernatant, or pour it off into the bleach bucket.

6. Pipette ANOTHER 1.8 ml of each culture into the corresponding microcentrifuge tubes.

7. Centrifuge at maximum speed for three minutes.

8. While the centrifuge is running, move the remaining cultures to 4degC.

9. Aspirate the supernatant off with the bench aspirator. Be careful not to disturb the pelleted E. coli.

• We use an aspirator here because the less extra salt and protein we put in the miniprep, the better the yield is.

10. Add 250 µl Buffer P1 to each tube.

11. Resuspend the E. coli pellet. The preferred way is with the roto-mixer at the other end of the lab.

• Alternately, if you have just a few tubes, you can resuspend on a vortex.

• Make sure to resuspend fully and thoroughly. The resulting suspension should be smooth and cloudy; if there is particulate matter floating around, vortex some more.

• Lysis

12. Add 250 µl Buffer P2 to each tube.

• Work quickly; the lysis step should take less than 5 minutes.

13. Snap the tubes closed and invert them 4-6 times, until the tube is thoroughly mixed and the entire solution turns blue.

• If you have many many tubes, you can stack a second tube rack on top of them and invert the entire thing.

14. Add 350 ul Buffer N3 to each tube.

15. Snap the tubes closed and invert 4-6 times, until the solution is thouroughly mixed and no longer blue.

• The solution will become cloudy or flocculent.

16. Centrifuge on high speed for 10 minutes.

• Separation

17. Remove the tubes from the microcentrifuge, being careful not to disturb the white pellet.

18. Using P-1000 micropipettor set to 850 ul, carefully transfer the supernatant from each centrifuge tube to the corresponding blue spin column.

19. Centrifuge the spin columns for 30 seconds at maximum speed.

• Don't forget to put the lid on the rotor! Some of the salts get aerosolized because the spin columns don't have caps.

20. Pour the flow-through from each column into the miniprep waste container.

21. Pipette 500 ul of Buffer PB onto each spin column.

22. Centrifuge the spin columns for 30 seconds at maximum speed.

23. Pour the flow-through from each column into the miniprep waste container.

24. Pipette 750 ul of Buffer PE onto each spin column

25. Wait 1-3 minutes.

• This allows some of the salt that's still bound to the silica matrix to resuspend in the buffer.

26. Centrifuge the spin columns for 30 seconds at maximum speed.

27. Pour the flow-through from each column into the miniprep waste container.

28. Return each spin column to its collection tube and centrifuge an additional 1 minute at high speed.

• This removes every last trace of buffer PE; the ethanol can screw up downstream steps.

29. Transfer each spin column to a clean labelled microcentrifuge tube.

30. Pipette 50 ul of Buffer EB onto the center of each column.

• The volume of EB is comparable to the volume of silica gel matrix; if you pipette down the side, you might not get the entire transfer to the matrix.

31. Wait 1-3 minutes.

• This gives the DNA a chance to dissociate from the silica matrix.

32. Centrifuge the spin columns, in their collection tubes, for one minute at maximum speed.

33. Proceed directly to analyze the samples on the Nanodrop.

Pouring an agarose gel

Introduction

Agarose gels are used to separate DNA fragments. Their most common use is

Materials

• TAE buffer, 50-100 ml
  • There is generally 1X TAE buffer at the iGEM bench in a 1L or 2L bottle.
  • If not, mix some more up from the carboy 10X stock by the main lab gel station. (Use the 1L graduated cylinder at the gel station.)
• UltraPure agarose
• Casting tray, casting stand, combs

Procedure

• Mix and melt the agarose

1. Check to see if there is a gel waiting in the fridge.

2. Determine how much molten agarose you'll need.

• A small gel cast takes 50 ml; a large gel cast is 100 ml.

3. Measure out the appropriate amount of TAE into a glass bottle or flask.

4. Add 5 µl SYBRSafe (0.5X ; stock solution is 10000X) per 100 ml of the solution and mix well.

5. Add UltraPure agarose to a final concentration of 1% (mass / volume)

• So, if you're making 50 ml gel: 0.5 g.

6. Swirl the bottle or flask to distribute the agarose.

7. Heat the solution in the microwave with frequent stirring to dissolve the agarose homogenously. ~30 seconds/100ml solution

8. Let sit until cool enough to handle.

• The agarose MUST cool some -- if it's too hot, it can warp the casting trays.

• Don't allow it to get too cold!

• PAUSE POINT - You can put the flask in the 55°C water bath almost indefinitely at this point.

• Pour the gel

9. Set up the gel tray in the casting stand. Make sure the rubber gaskets are flat up against the edges of the casting tray.

10. Set up the gel co mbs to form the wells.

• Rinse the combs with water and wipe dry.

• Note for combs: 15-well combs hold about 6 ul liquid per well, 12-well combs hold about 15 ul per well, 8-well combs hold about 20 ul per well

• Taping two 8-well comb wells together results in a well that holds up to 100 ul

• Taping three 8-well comb wells together result in a well that holds up to 200 ul

11. Pour the molten agarose into the casting tray.

• If bubbles form around the combs, remove and re-insert.

12. Wait 30 mintues for the gel to solidify.

13. Use immediately, or place in a plastic zip-lock baggie with a little 1X TAE and store at 4°C.

Sequencing protocol

Introduction

Preparing sample to send to Genewiz for sequencing Sequencing Pick-up Times: 9:30am, 3:00 pm, 5:30 pm

Materials

• DNA stock
• Nuclease-free water
• Primer (5nM stock)
• One microcentrifuge tube for each DNA plasmid
• 2 PCR tubes per plasmid (must be in strips of eight, round up if needed!)

Procedure

• Diluting the DNA

1. For orders with <48 samples, use 8-strip PCR tubes. Label your tubes on the side vertically with "MI" and sample number (01, 02, 03, etc.). Use image below as guideline. Each plasmid you want to sequence will require 2 tubes, one for the forward primer and one for the reverse primer. Use "MI" instead of the "GW" in the picture.

2. Dilute your sequencing primer to 5 µM (pmol/µl) using water. You will need 5 µl for each sequencing reaction. Note, the primers in the tubes from IDT have not been diluted! Use already diluted stock (should be in regular microcentrifuge tubes).

3. Using the recorded concentration, dilute the DNA to the template concentration in 10 µl as detailed below. For example, a 5 kb plasmid would need a template concentration of ~50 ng/µl. Final volume for your working stock should 30 uL. Please make dilutions in water or Tris.

4. Add 10 µl of diluted template DNA to each tube.

5. Add 5 µl of diluted primer to each tube. Be careful about putting the forward and reverse primers in the correct tubes! Remember, for each DNA template, there should be two tubes - one with reverse primer and the other with forward primer

6. Vortex briefly

7. Log into genewiz. Username: igem-sequence@mit.edu Password: igem2014citrus

8. Click "create sequencing order"

9. Sevice Priority: click on "standard"

10. Create Order by: click on "online form"

11. Sample Type: click on "Pre-mixed"

12. Enter in how many samples you are sending in and click "Create new form"

13. Enter in descriptive names for your DNA (this is to help you remember when you get the results back), the DNA type, the DNA length (get this from your construct on benchling), the primer name (again, this is more for you). When you are done hit save and next.

14. It will give you an list of what you have entered and a price for each reaction. Hit "next step"

15. The payment information field should be autofilled. There has been an issue recently with the PO box number not being entered. If this is the case, click "credit card" (next to "payment info") and then click "PO" to bring you back to the original form. Now you should be able to enter in a PO number. Open up a previous submitted order (there should be some saved to the desktop called "sequencing" or something like that) and copy over the PO number. PO Number: 5510061907

16. Hit "next step". Once it gives you a submitted order form, print 2 copies and pick them up from the printer (exit lab, through glass doors, on your left)

17. Put your PCR tubes in a large falcon tube.

18. Put the falcon tube in a ziploc bag (located in Qiagen column drawer) and put in one order form. Label the bag with something descriptive ("Weiss Lab MIT iGEM Recombinase 6/30/16" for example)

19. Put the bag in the Genewiz pickup box (located by the elevators, make sure you put in the genewiz box)

20. Open up the weiss lab orders spreadsheet (bookmarked on the computer). You need to request access to the spreadsheet if it is your first time ordering. Once you request access, email Brian so he knows that you have sent in a request.

21. Click the tab at the bottom called "genewiz".

22. Fill out the form. Make sure you indicate that the order is from iGEM so the right account can be charged.

23. You did it! YAY! Genewiz should get back to you pretty quickly, generally between 12-24 hrs.

DB 7/23/16

Made with Benchling

Project: iGem 2016 _ Recombinase subgroup

Authors: Elizabeth Strand

Date: 2016-07-23

Saturday, 7/23

LR Reactions

1 - hEF1a: L7Ae

2 - TRE: L7Ae

3 - EGSH: TP901

4 - hEF1a: attB-flipped EYFP-attP