Difference between revisions of "Team:Potsdam/Protocols"

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Our research work

We are describing our research work. Below you can find the protocols we used.

Protocols

1. Aim

2. Preparation of starting plasmids

1. Preparation restriction of components

1. Restriction using biobrick assembly enzymes.
2. This preparation step is needed to create sticky ends on the cassettes.
3. This step is only performed once.
4. Restriction with EcoRI and PstI (see restriction protocol) for all components.

2. Ligation of cassettes into plasmids

1. Ligation using T4-ligase (see ligation protocol)

plasmid	cassettes
psB1C3	1	2	3
psB1A3	1	2	3
psB1K3	1	2	3

3. Final preparation steps

1. Transformation of 9 different combinations into competent cells (see transformation protocol).
2. Selection with corresponding antibiotics.

4. Next day

1. Colony pcr and gel run to check for sizes of cassettes.
2. Miniprep and check concentration via nanodrop.
3. Many aliquots needed (small volume because thawing time) for future reactions!

3. 3A-assembly

1. Good to know

1. The 3-A-assembly will be used to add more and more cassettes in a row.
2. It is important to only combine cassettes with the same number (1, 2 and 3 have varying spacer length).
3. We will add cassettes and test frequently for viability to determine the maximum target-sequence length.
4. We want to combine about five cassettes.

2. Assembly scheme

1. In each assembly cycle, there will be two cassettes (same number/length) added to one linearized plasmid.
2. In the first step, we can either just use the prepared plasmid with cassettes already inserted or use an empty one, because the part in between will be cut out anyway.
3. The be able to select for plasmid with higher cassette content the resistances will cycle.
4. The resistance cycle (for plasmids) is K C A.
5. For the inserts, the resistance signals from which plasmid they will be cut.
6. The plasmids resistance determines the selection antibiotics for that step!

3. Legend

CX – Chloramphenicol (Insert/Plasmid) from step X
AX – Ampicillin (Insert/Plasmid) from step X
KX – Kanamycin (Insert/Plasmid) from step X
E – EcoRI
S – SpeI
X – XbaI
P – PstI

Insert 1 E+S	Insert 2 X+P	Plasmid E+P
C1	A1	K1
A2	K1	C2
K3	C2	A3
C4	A3	K4

4. Starting a cycle

1. Insert 1 is cut out of plasmid C1/Insert 2 is cut out of plasmid A1 and plamsid K1 is linearized .
2. Insert 1 and 2 are ligated into plasmid K1.
3. Insert 1 is cut out of plasmid A2/Insert 2 (here the fusion from Insert 1+2 from the first cycle) is cut out of K1 and plasmid C2 is linearized.
4. Insert 1 and 2 are ligated into plasmid C2.
5. With insert 1 coming from K3 and insert 2 from C2 (fusion) the steps will repeated until maximum number of inserts is reached.
6. The assembly needs to be done for all 3 cassettes simultaneously after each step.
7. Transformation of ligation into competent cells (see transformation protocol).
8. Selection with corresponding antibiotics.

5. Next day

1. Colony pcr and gel run to check for sizes of cassettes.
2. Miniprep and check concentration via nanodrop.

^[1] http://parts.igem.org/Help:Protocols/3A_Assembly

1. Aim

1. Is the insert DNA in the plasmid present or absent?

2. Much easier than to isolate and purify the vector.

2. Good to know before the start

1. Take typical measures to prevent PCR cross over contamination, keep your bench clean, wear gloves, use sterile tubes and filter pipet tips.
2. Include a no-template control and positive control in parallel.
3. Thaw and keep reagents on ice.
4. Mix well before use.
5. The longer the amplicon, the longer the extension time: Use 15 sec/kb extension.
6. Use 90 sec extension for multiplexing.
7. Run an annealing temperature gradient from 55 °C to 65 °C to choose the best specificity conditions. Do not use fast cycling for multiplexing.
8. ALLin™ Red Taq Mastermix, 2X is premixed with red dye and density reagents for direct loading on the gels after the PCR. In a 2 % agarose TAE gel the dye migrates with ~350 bp DNA, in 1 % agarose TAE gel with ~600 bp DNA fragments.

3. Are you working with A. E.coli or B. yeast?

A.step by step for E.coli:

1. Resuspend colonies.

2. Prepare masterplate.

3. Prepare a PCR master mix (always prepare at least 10 % more).

4. Mix gently, avoid bubbles.

5. Aliquote the 22.0 μl of PCR master mix into each PCR tube.

6. Pick colony and put the toothpick or tip one into the Masterplate and then in the PCR tube/mix.

7. Do not forget the negative control!

8. Close tube.

9. Perform the PCR using Thermocycler as follow:

Steps	Cycles	Temperature	Time
Initial denaturation	1 cycle	95°C	60s
Denaturation	30-40 cycles	95°C	15s
Annealing	30-40 cycles	55-65°C	15s
Extension	30-40 cycles	72°C	15-90s (15 sec per 1 kb)
Final extension	1 cycle	72°C	5 min

10. Store probes for short time on ice, for long at -20 °C.

11. Load probes on the agarose gel e.g. 10 μl (so in case you have enough left for another round).

B. Step by step for yeast:

1. If resuspended colonies are to be used: pipette 50 μl of a 0.02 M NaOH solution into each of a set of appropriately labelled PCR tubes or wells of a PCR plate. Using sterile pipette tips or toothpicks, transfer transformants to individual tubes/wells. The amount of cells resuspended must just be visible. Resuspend cells by pipetting or vortexing and incubate for ≥ 5 min at 37 °C.
2. If overnight cultures are to be used: pipette 40 μl of a 0.01 M NaOH solution into each of a set of appropriately labelled PCR tubes or wells of a PCR plate. Transfer 10 μl of each overnight culture to be tested to the appropriate tube/well and mix by pipetting up and down. Incubate for ≥ 5 min at 37 °C.
3. Prepare a PCR master mix (always prepare at least 10 % more).
4. Aliquot 22.5 μl of PCR master mix into each PCR tube.
5. Add 2.5 μl of the resuspended colony or overnight culture mixed with NaOH to the appropriate PCR tube.
6. Close the tubes
7. Perform the PCR using the following cycling profile:

Steps	Cycles	Temperature	Time
Initial denaturation	1 cycle	95°C	60s
Denaturation	30-40 cycles	95°C	15s
Annealing	30-40 cycles	55-65°C	15s
Extension	30-40 cycles	72°C	15-90s
Final extension	1 cycle	72°C	5 min

8. Load probes on the agarose gel.

9. Store probes for short time on ice, for long at -20°C.

^{[1]https://www.highqu.com/media/wysiwyg/ressources/manuals/PCM02_ALLin_Red_Taq_Mastermix_PI.pdf}

1. What is it ?

1. Standard lab procedure for separating DNA by size (e.g., length in base pairs) for visualization and purification.
2. Uses an electrical field to move the negatively charged DNA through an agarose gel matrix toward a positive electrode.
3. Shorter DNA fragments migrate through the gel more quickly than longer ones.

2. Why are we doing it ?

1. to determine the approximate length of a DNA fragment by running it on an agarose gel alongside a DNA ladder (a collection of DNA fragments of known lengths).

3. Protocol

1. Pouring a Standard 1% Agarose Gel:

1. Measure 1g agarose and and mix it with 100 ml of TBE in a microwaveable flask.
Note: Agarose gels are commonly used in concentrations of 0.7 % to 2 % depending on the size of bands needed to be separated - Simply adjust the mass of agarose in a given volume to make gels of other agarose concentrations (e.g., 2 g of agarose in 100 mL of TAE will make a 2 % gel).
2. Microwave for 1-3 min until the agarose is completely dissolved (but do not overboil the solution, as some of the buffer will evaporate and thus alter the final percentage of agarose in the gel. Many people prefer to microwave in pulses, swirling the flask occasionally as the solution heats up.).
Note: gloves and glasses ! Caution HOT! Be careful stirring, eruptive boiling can occur.
It is a good idea to microwave for 30-45 sec, stop and swirl, and then continue towards a boil. Keep an eye on it as the initial boil has a tendency to boil over. Placing saran wrap over the top of the flask can help with this, but is not necessary if you pay close attention.

4. Pouring of the gel

1. Let agarose solution cool down to about 50 °C (about when you can comfortably keep your hand on the flask), about 5 mins.
Note: Or cool down in water bath about 30 min.
2. Add ethidium bromide (EtBr) to a final concentration of approximately 0.2-0.5 μg/mL (usually about 2-3 μl of lab stock solution per 100 mL gel). EtBr binds to the DNA and allows you to visualize the DNA under ultraviolet (UV) light.
Note: Caution EtBr is a known mutagen. Wear a lab coat, eye protection and gloves when working with this chemical. If you add EtBr to your gel, you will also want to add it to the running buffer when you run the gel.
3. Pour the agarose into a gel tray with the well comb in place.
Note: Think about witch gel tray size you need (a small one or a big one).
Pour slowly to avoid bubbles which will disrupt the gel. Any bubbles can be pushed away from the well comb or towards the sides/edges of the gel with a pipette trip.
4. Let the newly poured gel sit at room temperature for 20-30 mins, until it has completely solidified.
if you are in a hurry the gel can also be set more quickly if you place the gel tray at 4 °C earlier so that it is already cold when the gel is poured into it.

5. Loading Samples and Running an Agarose Gel

1. Add loading buffer to each of your digest samples.
Note: Loading buffer serves two purposes: 1) it provides a visible dye that helps with gel loading and will also allows you to gauge how far the gel has run while you are running your gel; and 2) it contains a high percentage of glycerol, so it increases the density of your DNA sample causing it settle to the bottom of the gel well, instead of diffusing in the buffer.
2. Once solidified, place the agarose gel into the gel box (electrophoresis unit).
3. Fill gel box with 1xTAE (or TBE) until the gel is covered.
4. Carefully load a molecular weight ladder into the first lane of the gel.
Note: When loading the sample in the well, maintain positive pressure on the sample to prevent bubbles or buffer from entering the tip. Place the very top of the tip of the pipette into the buffer just above the well. Very slowly and steadily, push the sample out and watch as the sample fills the well. After all of the sample is unloaded, push the pipettor to the second stop and carefully raising the pipette straight out of the buffer.
5. Carefully load your samples into the additional wells of the gel.
6. Run the gel at 80-150 V until the dye line is approximately 75-80 % of the way down the gel.
Note: Black is negative, red is positive. (The DNA is negatively charged and will run towards the positive electrode.) Always Run to Red.
Note: A typical run time is about 1-1.5 hours, depending on the gel concentration and voltage.
7. Turn OFF power, disconnect the electrodes from the power source, and then carefully remove the gel from the gel box.
8. Using any device that has UV light, visualize your DNA fragments.
Note: When using UV light, protect your skin by wearing safety goggles or a face shield, gloves and a lab coat.
Note: If you will be purifying the DNA for later use, use long-wavelength UV and expose for as little time as possible to minimize damage to the DNA.
Note: The fragments of DNA are usually referred to as ‘bands’ due to their appearance on the gel.

6. Analyzing Your Gel

Using the DNA ladder in the first lane as a guide (the manufacturer's instruction will tell you the size of each band), you can interpret the bands that you get in your sample lanes to determine if the resulting DNA bands that you see are as expected or not. For more details on doing diagnostic digests and how to interpret them please see the Diagnostic Digest page.

7. Purifying DNA from Your Gel

If you are conducting certain procedures, such as molecular cloning, you will need to purify the DNA away from the agarose gel. For instructions on how to do this, visit the Gel Purification page.

1. Aim

A restriction digest is the division of DNA ina specific area by the help of restriction enzymes. The aim afterwards could be to analyze and characterize (restriction maps) the DNA, to compare it to others or to clone it into e.g. a vector. Previous to the restriction, the DNA has to be isolated (see protocol miniprep).

2. Test digest - What for?

In a test digest the cleavage products are analyzed to verify that the used DNA has e.g. taken in a specific fragment. Only few of the DNA is digested, because the uncut DNA will be used in further steps. The whole DNA strand is often too long to analyze, therefore:

targeted cuts between specific base sequences
cleavage fragments small enough to run on gel
segregation and analysis by gel electrophoresis

→ always with a control: uncut plasmid (see protocol gel electrophoresis) In a preparative digest normally 1 U enzyme digests 1 µg DNA in one hour.

3. Preparative digest - What for?

In a preparative digest the entire available DNA is digested, because the cleavage products are used in further steps. The cut DNA can be extracted from a gel. It is important that as much DNA as possible is digested. Therefore 0,2 - 0,4 µl enzyme per µg DNA are applied and the reaction should run for ~2h.

4. Procedure

exemplary pipetting scheme:

component	volume
DNA (typically 200 - 500 ng)	1 µl
10x buffer	1 µl
H₂O	7,6 µl
enzyme 1	0,1 µl
enzyme 2	0,1 µl
final volume	10 µl

If more than one reaction is done, it is convenient to prepare a master mix containing everything except for the DNA (!). This saves time and tips and keeps you from having to pipet very small volumes (e.g. 0,1 µl).
It is also possible to use more than two or just one restriction enzyme per reaction. They just need to have the same buffer preferences. Volumes have to be adjusted to the number of enzymes.

^[1] http://www.log2embl.de/sites/default/files/Labor-Protokoll-Restriktionsverdau.pdf

^[2] https://www.uni-hohenheim.de/fileadmin/einrichtungen/pflanzenphysiologie/Protokolle/V.Klonierung/restriktionsverdau_de.pdf

^[3] http://www.biochemie.uni-jena.de/files/Praktikum/plasmid%20dna%20+%20restrictionsverdau.pdf

1. What is it?

Transmission of genetic information into competent cells (or plants, algas, mushrooms) (the target organismn)

2. What you need

1.5 mL tube, pipette (50µL), pipette(1000µL), plate
Ice, SOC Medium, competent cells

3. Steps

1. Prechil 1.5 ml tube on ice
2. Thaw a tube competent E. coli cells on ice for 10 minutes.

1. Mix gently
2. Pipette 50 µl of the cells into the 1.5ml tube (Temperatures over 0°C decrease the efficiency of the transformation!)

3. Add 1-5 µl (containing 1 pg-100 ng of plasmid) DNA to the cell mixture.
(as soon as as the last bit of ice in the tube is disappeared!)
4. Flick the tube 4-5 times to mix cells and DNA. (No vortexing!)
5. Place the mixture on ice for 30 minutes. (without mixing!)
(2-fold loss in transformation efficiency for every 10 minutes this step is shortened!)
6. Heat shock at exactly 42°C for exactly 30 seconds. (without mixing!)
(temperature and timing specific to transformation volume and vessel)
7. Place on ice for 5 minutes.
(without mixing!)
8. Pipette 950 µl of room temperature SOC into the mixture.
9. Place at 37°C for 60 minutes and shake vigorously (800 rpm in thermo mix block)
(2-fold loss in transformation efficiency for every 15 minutes this step is shortened)
(SOC gives 2-fold higher transformation efficiency than LB medium)
(incubation with shaking or rotating the tube gives 2-fold higher transformation efficiency than without)
10. Warm selection plates to 37°C
(plates can be used warm or cold, wet or dry…efficiency is nearly the same… warm plates are easier to spread and allow most rapid colony formation.)
11. Mix the cells thoroughly by flicking the tube and inverting.
12. Spread 200 µl onto a selection plate and incubate overnight at 37°C.
13.For low efficiency cloning reactions: spin down the whole transformation mixture and remove the nearly complete supernatant (approx. 900 µl). Resuspend cells in remaining liquid and plate completely.

^[1]https://www.neb.com/protocols/1/01/01/high-efficiency-transformation-protocol-c2987

Promega “Wizard Plus SV Miniprep Purification System“

1.Production of cleared lysate

1. Isolation of the bacteria

1. harvest 1–5 ml (high-copy-number plasmid) or 10 ml (low-copy-number plasmid) of bacterial culture
2. centrifugation for 5 minutes at 10,000 xg in a tabletop centrifuge
3. pour off the supernatant
4. reinsert again bacterial culture to the pellet and repeat step 2 and 3
5. blot the inverted tube on a paper towel to remove excess media

2. Resuspension of the cells

1. add 250 μl of Cell Resuspension Solution
2. completely resuspend the cell pellet by vortexing or pipetting
3. it is essential to thoroughly resuspend the cells

3. Lysing

1. add 250 μl of Cell Lysis Solution
2. mix by inverting the tube 4 times - do not vortex
3. incubate until the cell suspension clears (clear ≠ colorlessly) (approximately 1–5 minutes)

4. Splitting proteins

1. add 10 μl of Alkaline Protease Solution
2. mix by inverting the tube 4 times - do not vortex
3. incubate for 5 minutes at room temperature

5. Neutralization

1. add 350 μl of Neutralization Solution
2. immediately mix by inverting the tube 4 times - do not vortex

6. Isolation of the plasmids

1. centrifuge the bacterial lysate at maximum speed (around 14,000 ×g) in a microcentrifuge for 10 minutes at room temperature

2. Isolation of the plasmid DNA

1. Transfer the cleared lysate (approximately 850 μl, Section 3.B, Step 6) to the prepared Spin Column by decanting. Avoid disturbing or transferring any of the white precipitate with the supernatant.

1. If the white precipitate is accidentally transferred to the Spin Column, pour the Spin Column contents back into a sterile 1.5ml microcentrifuge tube and centrifuge for another 5–10 minutes at maximum speed. Transfer the resulting supernatant into the same Spin Column that was used initially for this sample. The Spin Column can be reused but only for this sample.

2. Centrifuge the supernatant at maximum speed in a microcentrifuge for 1 minute at room temperature. Remove the Spin Column from the tube and discard the flowthrough from the Collection Tube. Reinsert the Spin Column into the Collection Tube.

3. Wash the plasmid DNA

1. Add 750 μl of Column Wash Solution.
2. Centrifuge at maximum speed in a microcentrifuge for 1 minute at room temperature.
3. Remove the Spin Column from the tube and discard the flowthrough.
4. Reinsert the Spin Column into the Collection Tube.

4. Wash again the plasmid DNA

1. Add 250 μl of Column Wash Solution.
2. Centrifuge at maximum speed in a microcentrifuge for 2 minutes at room temperature.
3. If the Spin Column has Column Wash Solution associated with it, centrifuge again for 1 minute at maximum speed.
4. Transfer the Spin Column to a new, sterile 1.5ml microcentrifuge tube, being careful not to transfer any of the Column Wash Solution with the Spin Column.

5. Elute the plasmid DNA

1. Add 50 μl of Nuclease-Free Water to the Spin Column, wait 5 minutes
2. Centrifuge at maximum speed for 1 minute at room temperature in a microcentrifuge.

6. After eluting the DNA, remove the assembly from the 1.5ml microcentrifuge tube and discard the Spin Column.

^[1]https://www.promega.de/-/media/files/resources/protocols/technical-bulletins/0/wizard-plus-sv-minipreps-dna-purification-system-protocol.pdf

@@ Line 709: / Line 709: @@
 <div class="inner" style="display:none;">
 <br>
-<b> 1. Aim </b><br>
+<b> 1. Aim </b><br><br>
 <div style="text-align: justify; margin-left:20px">
 A restriction digest is the division of DNA ina  specific area by the help of restriction enzymes. The aim afterwards could be to analyze and characterize (restriction maps) the DNA, to compare it to others or to clone it into e.g. a vector. Previous to the restriction, the DNA has to be isolated (see protocol miniprep).</div>
 <br>
-<b> 2. Test digest - What for?</b><br>
+<b> 2. Test digest - What for?</b><br><br>
 <div style="text-align: justify; margin-left:20px"><br>
 In a test digest the cleavage products are analyzed to verify that the used DNA has e.g. taken in a specific fragment. Only few of the DNA is digested, because the uncut DNA will be used in further steps. The whole DNA strand is often too long to analyze, therefore:</div><br>
@@ Line 725: / Line 725: @@
 In a preparative digest normally 1 U enzyme digests 1 µg DNA in one hour. </div><br>
-<b>3. Preparative digest - What for? </b><br>
+<b>3. Preparative digest - What for? </b><br><br>
 <div style="text-align: justify; margin-left:20px">
 In a preparative digest the entire available DNA is digested, because the cleavage products are used in further steps. The cut DNA can be extracted from a gel.
 It is important that as much DNA as possible is digested. Therefore 0,2 - 0,4 µl enzyme per µg DNA are applied and the reaction should run for ~2h.</div>
 <br>
-<b> 4. Procedure</b><br>
+<b> 4. Procedure</b><br><br>
 <div style="text-align: justify; margin-left:20px">
 exemplary pipetting scheme:</div>


If there is more than the wanted DNA band: A. Dissolving the Gel Slice	If there iss only one DNA band: B. Processing PCR Amplifications
1. Following electrophoresis, excise DNA band from gel (with scalpel) and place gel slice in a 1.5 ml microcentrifuge tube. 2. Add 10 μl Membrane Binding Solution per 10 mg of gel slice. Vortex and incubate at 50 – 65 °C until gel slice is completely dissolved.	You can just work with the rest of your PCR aliquote (when you did not use all of it for the gel electrophoresis). 1. Add an equal volume (so you have to know how much aliquote is still in your eppi!) of Membrane Binding Solution to the PCR amplification.

Component	25 μl Reaction	50 μl Reaction	Final Concentration
Q5 High-Fidelity 2X Master Mix	12.5 μl	25 μl	1X
10 μM Forward Primer	1.25 μl	2.5 μl	0.5 μM
10 μM Reverse Primer	1.25 μl	2.5 μl	0.5 μM
Template DNA	variable	variable	< 1,000 ng
Nuclease-Free Water	to 25 μl	to 50 μl

Step	Temperature	Time
Initial Denaturation	98°C	30 seconds
	98°C	5–10 seconds
25–35 Cycles	*50–72°C	10–30 seconds
	72°C	20–30 seconds/kb
Final Extension	72°C	2 minutes
Hold	4-10°C

	volume
T4 DNA Ligase Buffer (10 x)	2 µl
T4 DNA Ligase	1 µl
Vector DNA
Insert DNA
Nuclease-free water	to 20 µl

Difference between revisions of "Team:Potsdam/Protocols"

Revision as of 17:24, 31 October 2017

IGEM Potsdam