Team:Heidelberg/Notebook MaM

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=Materials & Methods=

Kits
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Marker
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Enzymes
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Restriction Enzymes
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Bacteria
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Bacteria Growth Media
Luria Broth (LB)
 * 10 g/L tryptone
 * 5 g/L yeast extract
 * 10 g/L NaCl

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Eucaryotic cell lines

 *  MCF-7 
 *  HeLa
 *  U2-OS

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DMEM+++

 * DMEM
 * 1% L-Glutamine
 * 1% Pen/Strep
 * 10% FCS

DMEM++

 * DMEM
 * 1% L-Glutamin
 * 1% Pen/Strep
 * 1% non-essential aminoacids

Zeocin-Medium

 * DMEM+++
 * 100 µl/ml Zeocin

Hygromycin-Medium

 * DMEM+++
 * 200 µl/ml Hygromycin

Minimal Medium

 * 2,1 g/l NaHCO3
 * 1µM CaCl2
 * 10 µM Hepes-Buffer
 * 1 % Pen/Strep
 * Krebs-Henseleit-Buffer (Sigma-Aldrich )

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Oligos
We used LabLife, a great, non-commercial lab management resource for lab management and especially for keeping track of Oligos we used. All Oligos therefore are numbered, and all Notebook entries describing PCR reactions refer to a certain number of Oligo. A complete list of all Oligos we had in our lab can be downloaded [[Media:HD09_complete_oligo_list.txt|as comma-separated values]]; find below a list of more interesting oligos we used for RA-PCR and LAM-PCR.

Oligos for real-time RT-PCR
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Plasmids
The following is a list of all used plasmids. Some of them are just working plasmids, others are of greater value for our project. The latter are bold in the table.

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Instruments
The following is a list of all used Instruments.

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Lab material
The following is a list of the used Lab material.

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= Methods =

Transformation of Bacteria
For enrichment of vectors, E. coli DH5α were used. For the transformation 100 µl of the competent cells were thawed on ice and 50 – 400 ng DNA solution added (depending on the concentration of the DNA solution). After a 30-60 minute incubation on ice, cells were made permeable for the DNA by heat shocking for 45 seconds at 42 °C and a further 3 minute incubation on ice. The samples were than rescued by adding 500 µl preheated antibiotic free LB-medium and incubated for one hour at 37 °C while shaking for induction of the antibiotic resistance. The selection for plasmid containing and therefore antibiotic resistant bacteria was conducted by plating them on antibiotic containing LB-agar plates.

Glycerol stock
To store bacteria for long term glycerol stocks were used. Therefore 1 ml of an over night culture were added to 150 µl of 80 % Glycerol into a cryo tube, vortexed and incubated at room temperature for 30 min. Afterwards the glycerole stock was stored at -80 °C.

Isolation of plasmid DNA by alkaline lysis (mini and maxiprep)
For analysis of ligations and transformations QIAprep Spin Kits (Qiagen, Hilden) were used following the manufacturer instructions. For miniprep a single colony was picked from a LB-agar plate or glycerol stock was used to inoculate 5 ml LB-medium with appropriate antibiotic for selection (100 µg/µl ampicillin, 50 µg/µl kanamycin, 35 µg/µl chloramphenicol). Bacteria were grown over night at 37 °C while shaking (200 rpm). By using 4 ml over night culture with this kit the yield was around 6-10 µg. For maxipreps the Qiagen CompactPrep Plasmid Maxi Kit was used following the instructions given by the instruction manual. In this case 250 ml LB-medium were inoculated and used for preparation of plasmid DNA. The routinely yield was 300-400 µg plasmid DNA. Purity and amount of DNA was analysed using a NanoDrop.

Preparing chemically competent cells
First, a 20 ml over night culture was inoculated in antibiotic free LB medium from a fresh single colony and transferred into 400 ml antibiotic free LB medium the next day. This culture was incubated at 37 °C while shacking until an OD600 of 0.5 – 0.6 was achieved. The culture was than cooled down on ice, centrifuged (8 min, 4 °C, 3500 rpm), the supernatant discarded and the pellet resuspended in 10 ml 100 mM CaCl2. After addition of further 190 ml 100 mM CaCl2 the suspension was incubated on ice for 30 min. The suspension was than again centrifuged (8 min, 4 °C, 3500 rpm), the supernatant discarded, the pellet resuspended in 20 ml 82.5 mM CaCl2 with 17.5 % glycerol and aliquoted. The aliquots were flash frozen in liquid nitrogen and than stored at -80 °C until usage.

Ligation
Standard Ligation Protocol

T4 ligase joins the 5' phosphate and the 3'-hydroxyl groups of DOUBLE stranded DNA molecules. First step for ligation is to estimate the vector and insert concentrations:if insert is from PCR, assume that 50% is recovered, typically 5 mg, and resuspended in 10 ml of H20; same with vector, assume half is recovered in purification/precipitation and resuspend in 10-20 ml of H20 if it is not re-suspended already. Second step is to plan control reactions: One reaction with no insert, one reaction with no vector (if enough insert is available). Then Molar Ratio of Insert to Vector is determined where a ratio of Insert:Vector 3:1 (100-150ng Vector DNA) is tried to be achieved. After this, ligation mixure is setup in the following way: Samples are incubated according to the guidelines below After incubation the ligase is heat inactivated by placing tube in 65 °C water bath for 10 minutes. 2 ml of the ligation mixture is used for transformation. Transformed cells are plated on selective media and incubated overnight.
 * 10X Ligase buffer (Promega #M1801)
 * 1 ml T4 Ligase (Promega #M1801, 3 U/ml)
 * Bring volume to 10 (or 20 see note below) ml with nuclease-free water.
 * If doing a blunt-end ligation, it may be needed to add PEG (up to 15%) to increase the efficiency.
 * Incubate sticky end ligation reactions at room temperature for 3 hours (or at 4 to 8 °C, overnight).
 * Incubate blunt-end ligation reactions at 17 °C for 4 to 18 hours.

Site-directed mutagenesis
For removal of unwanted Restriction Sites, a PCR-based site direct mutagenesis protocol was adapted from Stratgene. Oligos were designed to have a high (>78°C) Tm by applying the formula The following scheme was used for pipetting:

The PCR procedure was as follows:


 * Initial denaturation, 1 cycle
 * 95 °C, 30 sec
 * Amplification, 16 cycles
 * denaturation 95 °C, 50 sec
 * annealing, 60 °C, 50 sec
 * extension, 72 °C, 40sec/kb + 30sec
 * Final Extension,1 cycle
 * 72 °C, 5 – 10 min
 * 4 °C hold, forever

After completion, 1 µL of DpnI (NEB) was added and the mix was incubated for 1h at 37°C. Then, DH5alpha cells were transformed as described above.

DNA synthesis
Oligos were designed using GeneDesign. 15Bp were chosen as an overhang and 56 °C as an annealing temperature. Oligos were ordered at a concentration of 100 µM. 1:10 dilutions of the first and last oligo were prepared. Then, all oligos (including first and last) were pooled at 1 µl each. Water was added to 10x the original volume. (So if a gene is synthesized from x = 14 oligos, water was added to 10*x = 140 µl). This pool was then diluted 1:10. x µl of the dilution were put into a PCR reaction with 25 µl Phusion master mix and 25-x µl water. PCR was conducted as following:


 * Initial denaturation, 1 cycle
 * 95 °C, 5 min
 * Amplification, 7 cycles
 * denaturation 95 °C, 30 sec
 * annealing, 58 °C, 30 sec
 * extension, 72 °C, 1 minute
 * 4 °C hold, forever

Afterwards, 1 µl of the 1:10 dilutions of the innermost and outermost primer were added. The same PCR protocol was then repeated, but with 25 instead of 7 cycles. PCR products were run on a 2% agarose gel and gel purified.

Purification of DNA from PCR reactions
PCR products were purified by the QIAquick PCR Purification Kit from Qiagen following the instructions of the Qiagen Handbook. To check the purity and amount of extracted DNA an aliquot was analysed using a NanoDrop.

Enzymatic hydrolysis of DNA by restriction enzymes
Restriction digest Restriction digest of DNA was used for analysis of purified DNA form mini or maxiprep or for isolation of specific DNA fragments for further cloning. Analytical digestions were routinely conducted in 20 µl volume,preparative digestions were routinely conducted in 50 µl volume. In all digestions a minimum of 2 Units restriction enzyme(s) was used per microgram DNA. Optimised buffer conditions were secured by using NEB buffer system. The final reaction volume was achieved by adding H2Odest. The sample was incubated at optimal temperature for the restriction enzyme(s) (usually 37 °C). Analysis and preparation of DNA were either done by gelelectrophoreses (loading dye was added to the samples and they were loaded on a agarose gel) or alternatively PCR Purification (PCR Purification QIAquick Kit from Qiagen)

Phosphatasing with Shrimp Alkaline Phosphatase SAP is used for Dephosphorylation of 5'-ends of DNA in Restriction Enzyme Reaction. After restriction digest 1 unit of SAP is added for every 1 pmol of DNA ends (about 1 μg of a 3 kb plasmid) and incubated at 37°C for 30-60 min. The reaction is stopped by heating at 65°C for 15 min (this completely inactivates SAP) The minimum effective amount of SAP for dephosphorylation of 1 pmol of DNA termini in 1 hr at 37°C is:
 * 0.05 units for 5'-protruding termini
 * 0.05 units for blunt termini
 * 0.1 units for 5'-recessed termini

Agarose gel electrophoresis for separating DNA
In the agarose gel electrophoresis a mixture of DNA fragments with different sizes are separated in an electrical field by their size. This is achieved by moving the negatively charged DNA through an agarose matrix while shorter fragments will run faster. The size of the pores can be controlled by agarose concentration. The higher the agarose concentration the smaller the pores are and the smaller fragments can be separated. Agarose concentrations between 0.7 and 1.5 % agarose in 0.5x TE buffer were used. The agarose was dissolved completely by heating up and 0.1 µg/ml ethidium bromide was added. The DNA fragments were separated using a constant voltage between 80 and 130 V. Under UV light (λ = 254 nm) DNA is visible through the unspecific intercalated ethidium bromide and can be documented or cut out and extracted from the gel.

Isolation of DNA fragments from an agarose gel
Plasmid DNA and DNA fragments were extracted using the Gel Extraction Kit from Qiagen following the manufacture instructions. To check the purity and amount of extracted DNA an aliquot was analysed using a NanoDrop.

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Splitting cells
Cultures of HeLa, MCF7 and U2OS cells are splitted 1:10 every 3-4 days or 1:3 every day: DMEM+++ (see Material) covering the cells is removed and the cells are washed with 3 ml PBS. The PBS is removed after the washing step and 3 ml of trypsin are added to the flask; almost all of the Trypsin is removed again until the cells are only slightly covered with Trypsin. The trypsinized cells are then incubated for 5 min at 37 °C and 5% CO2. After this incubation time, the cells are detached from the bottom and the flask is filled with 5 ml of DMEM+++. Cells are resuspended in medium by pipetting up and down. To achieve a diluton of 1:10, 0,5 ml of the suspension is transferred to another flask and 5 ml of DMEM+++ are added. This dilution is then incubated for 3-4 days at 37°C and 5% CO2. The rest of the cell suspension (about 4 ml) is transferred to a 50 ml falcon tube and the same volume of DMEM+++ is added to the falcon. 9,6 µl of this dilution are transferred to a counting chamber and the number of cells is determined (x*104).
 * Note 1: HeLa and U2-OS seem to grow faster than MCF-7; check notebook for information on splitting ratios

Transfection of Mammalian cells
1.Fugene (Roche) 105 cells were moved to 6-well plates. 2,5 ml of DMEM medium plus addiditives were added. The following day, 97 µl pure DMEM was incubated for 5 minutes with 3 µl Fugene6 reagent at room temperature. Afterwards, the mixture was added dropwise to 500 ng of plasmid and allowed to incubate for 15 minutes at RT. The mixture was then added dropwise to the cells. Cells were incubated for 24 hours without selection pressure, afterwards, medium was changed and Zeocin was added to a concentration of 350 ng/ml.

2. Effectene Transfection (Qiagen) Move x (for apropriate amounts see table below) cells to 6-well plates/ LabTek 8 chamber and add 2 ml/0,5 ml DMEM+++. Incubate over night at 37 °C and 5 % CO2.The following day, dilute x µg of DNA in Buffer EC to a total volume of x µl (note: Yara just adds 100 µl/50 µl). Add x µl of Enhancer and mix by tapping the tube (or vortex for 1 sec). Incubate for 2-5 min at RT. Add x µl of Effectene Transfection reagent to the DNA-Enhancer Mixture. Mix by tapping the tube (or vortexing for 10 s or pipetting up and down 5 times). Incubate samples for 5- 10 min at RT to allow for transfection-complex formation. While complex formation takes place, aspire medium and add x ml of fresh growth medium. After incubation time add the transfection -complexes drop-wise to the cells on the well. Gently swirl the plate to ensure uniform distribution of the complexes.
 * transient transfection: remove Effectene-DNA complexes after 6-18 hours, wash cells 1x with sterile PBS, add fresh medium. Incubate cells with the complexes under normal growth conditions for an appropriate time for expression.
 * stable transfection: change medium after 24 hours (wash with PBS); check confluency after 48 hours - (if required) split to 25 % confluency, add fresh medium and incubate 2-3 hours until cells have attached to culture dish. Then, remove medium and add fresh DMEM+++/Zeocin.

Freeze cells
Cells to be frozen should be confluent, but not too dense. Medium covering the cells is removed and cells are washed once with PBS. Afterwards cells are rinsed with 3 ml Trypsin; Trypsin is removed until the cells are only slightly  covered (about 1 ml) and cells are incubated for about 5 min. After incubation, cells are resuspended in medium (at least about 4-5 times as much as Trypsin to inactivate that;  used 5 ml). The suspension is transferred to 15 ml falcon and centrifuged at 2000 rpm for 3-5 minutes. The supernatant is discarded and the remaining pellet is resuspended in DMEM+++ (here: 7,5 ml for 5 aliquots, 1,5 ml each). 1,5 ml of the suspension is distributed into freezer vials. After adding of 150 µl DMSO to each vial, cells are put on ice and stored at -80 °C.

Picking cells with cloning disks
Foci of cells, which were under selection pressure (zeocin) are marked on the bottom of the petri dish (use pen from underneath the microscope). As many cloning disks as foci available are prepared by incubating them 3-5 minutes in trypsin (in a petri dish). Medium covering the cells is removed and cells are washed with PBS. PBS is removed till the cells are only slightly covered with it. The cloning disks are placed on the previousely marked spots and incubated for 3 minutes at 37 °C. During incubation a 96-well plate is prepared with 0,7 ml DMEM+++/Zeocin per well. The clonig disks are picked up with pressure (or swipe a little) to remove the entire foci off the plate and the cloning disk is transferred to the 96-well plate (one disk per well). Medium is pipetted up and down to loosen the cells from the cloning disks.

Fix cells for microscopy
The medium is removed from the cells on cover slips. 500 µl of 4% Paraformaldehyde (in PBS) is added to the cells and incubated for 10 minutes. PFA is taken off and cells are washed with 500 µl of PBS. Slides are labeled and a small drop of fluoromount is added with a pipet tip. Cover slips are washed in dH20, by picking them up with tweezers and dipping them into the water. Afterwards cover slips are dried with filter paper without the paper touching the cover slip side containing the cells. Cover slips are layed down on the drop of fluoromount with the cells facing downwards. Fixed cells are dried overnight and can then be used for fluorescence microscopy.

Induction
To induce the respective pathways specifically, we searched literature and used knowledge of groups working with the drugs. The following table is an overview of the pathways, the used drugs and the conditions for activation.

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RA-PCR
For a RA-PCR protocol, please refer to Project page

We used the following Oligo concentrations:

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Screening by TECAN
Cells were grown in black 96 well plates (PerkinElmer) and induced as described for the various pathways. Outer wells were left free. After induction, media was removed and replaced by PBS. Fluorescence was then measured using a TECAN infiniteM200 automated plate reader (excitation = 488 nm, emission=518 nm). In order to account for variations in cell number, cells were stained by Hoechst 33342 dye. PBS was removed and replaced by PBS containing Hoechst 33342 dye at a concentration of 1µg/mL. Cells were allowed to incubate at room temperature for 30 minutes. Afterwards, medium was replaced by PBS again and fluorescence (excitation = 355nm, emission=455nm) was measured again by TECAN. Fluoresence values from the first reading were divided by fluorescence values of the second reading and multiplied by the lowest value of the second reading. Clones showing induction were considered further.

Flow Cytometry (FC)
Before preparing samples for flow cytometry the flow cytometer has to be turned on at least 20 min before the run. Samples are then prepared in the following way (this is for a 96 well plate): As much DMEM+++ as possible is removed from the cells. 60 µl Trypsin are added and the cells are incubated at 37 C for 10-15 min. Then, 160 µl of 1xPBS + 1% BSA are added. Samples are now ready for FC. At the FC the correct plate format is chosen. The plate is placed in the machine and the lid is taken off. When starting the flow cytometry measurement we first adjusted the gates and the background signal to the negative control. Measurement parameters such as plateformat, wells, and the desired protocol are chosen from the list. Volume and Mix iterations are are also chosen and saved as... .The discriminator is set to increase the threshold(2-10). The measurement is started with forward vs. side scatter in order to find cells. When cells are found, a gate (line) is added around the cells or cell candidates. Selected cells are then measured. Cells were prepared in 96-well format with 10^4 cells/well and transfected with the promoter of interest and if necessary induction drug was added for inducible promoters.

Microscopy/ Image Analysis
Microscopy images of our samples fixed with 4% formaldehyde were taken using the Nikon Eclipse 90i upright automated widefield microscope in the Nikon Imaging Center at the University of Heidelberg. Each image was taken in the GFP and mCherry channel and the exposure time had to be adjusted for over- or underexposed images. The conversion factor was experimentally determined by imaging a fluorescent plate with exposure time varying from 10 to 70 ms (the estimated range of our promoters) and plotting the mean grey values of these images versus time. A linear relationship between the exposure times and grey values was observed for this time frame and the conversion factor was determined to be 0.998. For image analysis we used the ImageJ (Image Processing and Analysis in Java) software measuring the mean grey values of each cell containing the promoter of interest. ImageJ is an open source software developed at the National Institutes of Health (http://rsbweb.nih.gov/ij/). The measurement is performed by setting the boundaries around the cells using the threshold in the mCherry image and generating a mask that is then applied to the image of the GFP channel in the following steps:


 * Open image and select split channels
 * Select mCherry image (all images should be 8-bit)
 * Go to Image> Adjust> Threshold
 * Set lower bar to maximum and adjust upper bar so that the cells are still visible
 * Remove unwanted spots such as dead cell using the drawing tools
 * Apply the threshold (image should appear in all white now with black spots indicating the cells)
 * Go to Process> Math> Divide and divide the image by 255 (image should be all white now)
 * Next go to Process> Image Calculator and multiply the GFP channel with the mCherry channel pictures.
 * Now select the result image and adjust the threshold so that the lower bar is set to the maximum (255) and the upper bar is set to 1 (only GFP containing cells should be visible through the mask)
 * Go to Analyze> Set Measurement and make sure the measurement is limited to the threshold and the result image of the multiplication is selected.
 * Go to Analyze> Measure.
 * Mean grey values should show up in the 'Results' window

Alternatively the images can be analyzed using the ROI manager where the cell have to be selected by hand using the drawing tools and added to the ROI list. In comparison the threshold method is much quicker and the results are comparable. We preselect the mCherry positive cells which are very likely to contain GFP. The mean grey values of 5-10 images per sample were averaged and adjusted to the same exposure time. The results were given in REU relative to JeT, which was always prepared as a reference sample for each measurement.

Real-time RT-PCR
Cells were collected as a pellet. Cell number must excede 1*105cells/tube and no more than 6*106/tube due to column capacity. In order to obtain reliable results and reduce the variance during the extraction, reverse transcription and PCR, we did the measurement for each promoter with 12 replicates from an identical sample. Resuspend the pellet with 600µl RLT buffer containing ß-ME (1:100). QIAcube follows protocol "RNeasy Mini Animal Cell QIAshredder Version 3" using RNeasy Plant Mini Kit. mRNA from untransfected HeLa cells was extracted to be "Standard" to perform plate-to-plate correction. After RNA is extracted, the extracts were put on ice immediately. 2 tubes of 5 µl aliquots were taken, One for measuring RNA concentration and one for gel electrophoresis. The RNA concentration was measured using NanoDrop under 230 nm. Each sample was measured 3 times. The gel electrophoresis was performed to confirm the RNA integrity because RNA degrade under many conditions. We used 1% agarose gel and 0.5x TAE buffer. Next step is the Real-time RT-PCR. We had different setup for different kits. In each plate, the 5 housekeeping genes were crossed with 12 replicates as well as Standard. eGFP was mixed with 12 replicates and RNase-free water as negative control. After the program finished, data was collected from OneStep software 2.0 associated to the StepOnePlus system. With a certain threshold (0.05), a series of Ct values were determined, then analyzed with MatLab.
 * For SensiMix One-Step Kit (Quantace):
 * 96-well reaction setup (25 µl/reaction):
 * 2X MasterMix:               12.5 µl
 * forward primer (5 µM):    1.0 µl
 * reverse primer (5 µM):    1.0 µl
 * probe (1.67 µM):             1.0 µl
 * RNase inhibitor:               0.5 µl
 * MgCl:                               0.5 µl
 * RNase-free water:           3.5 µl
 * RNA template (20 ng/µl): 5.0 µl
 * Program setup:
 * Reverse transcription: 49°C, 30 min
 * Enzyme activation: 95°C, 10 min
 * Amplification: 35 cycles:
 * 95°C, 10 s
 * 60°C, 1 min
 * For QuantiTect Probe RT-PCR Kit (QIAGEN):
 * 96-well reaction setup (25 µl/reaction):
 * 2X MasterMix: 12.5 µl
 * forward primer (5 µM): 2 µl
 * reverse primer (5 µM): 2 µl
 * probe (1.67 µM): 2 µl
 * RNase-free water: 1.25µl
 * RT Mix: 0.25µl
 * RNA template (20 ng/µl): 5.0 µl
 * Program setup:
 * Reverse transcription: 50°C, 30 min
 * Enzyme activation: 95°C, 15 min
 * Amplification: 45 cycles:
 * 94°C, 15 s
 * 60°C, 1 min

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