Transfection

Mammalian cell transfection is used to introduce foreign genetic sequences into cells. Depending on cell line and desired effect, there are different ways to transfect mammalian cells. These methods can be broadly classified into two groups: transient and stable transfections. Normally, a selection marker is needed to identify cells that are transfected.

Transient Transfection

Transient transfections generate short-term impact of alterations in gene and protein expression. In this case, the expression of the introduced gene is temporary because the foreign nucleic acids are not integrated into the host cell genome and therefore will not be passed on by cell division. Transient transfection of cells was achieved by two different methods:

Stable transfection

The aim of stable transfection is to introduce a gene of interest into the genome of cells, which results in permanent expression of foreign genes in target cells. Lentiviral transduction is used in this project to generate stable cell lines.

1. Make packaging mix:
   Mix packaging and envelope plasmids, fill up with sterile filtered 150 mM NaCl to an end volume of 990.4 µl and incubate for 10 min at room temperature
2. Make PEI mix:
   Mix 115.8 µl PEI (7.5 mM) with 884.2 µl sterile filtered 150 mM NaCl and incubate for 10 min at room temperature
3. Make construct mix:
   Add 983,9 µl of the packaging master mix and 1000 µl of the PEI mix to the transfer plasmid
4. Incubate for 15 min
5. Pipette 2000 µl of the construct mix dropwise to the HEK293T cells
6. After 24 hours and 48 hours of incubation, collect the media and add fresh medium to the cells
7. Supernatant of the collected medium contains virus, which can be directly stored or centrifuged to concentrate virus

Cryo Stocks

To store cells for a long time cryo stocks were made:

HEK cells: HEK cells were washed with PBS and trypsinized. Trypsinization was stopped with RPMI media and the sample was centrifuged
(100 g, 3 min). The supernatant was discarded and the pellet was resuspended in RPMI and the cells counted according to our protocol (LINK?).

After another centrifugation step (100 g, 3 min) the supernatant was discarded and the pellet resuspended in 1 ml of 90% FBS and 10% DMSO.

The cells were transferred to a cryo tube, which was stored for 2 days at -80 °C in an ice box to ensure gradual freezing of the cells. The tubes were then placed into a liquid nitrogen tank for long term storage.

Jurkat cells: Jurkat cells from a full 10 cm dish were transferred to a 15 ml falcon tube and centrifuged (100 g, 3 min). The supernatant was discarded and the pellet resuspended in 2 ml of 90% FBS and 10% DMSO. Aliquots of 1 ml cell suspension were transferred into a cryo tube and stored for 2 days at -80 °C in an ice box to ensure gradual freezing of the cells. Afterwards the tubes were placed into a liquid nitrogen tank for long term storage.

Thawing of the samples:

To thaw samples, they were taken out of the liquid nitrogen tank and placed into a 37 °C incubator. 15 ml falcon tubes with 5 ml RPMI were also placed in the 37 °C incubator to heat up. Afterwards the thawed sample was transferred to the warmed 15 ml falcon tube with RPMI and centrifuged (100 g, 3 min). The supernatant was discarded and the pellet resuspended in RPMI. The sample was then transferred to a 6-well plate and could be placed back into the incubator.

Fluorescence-Activated Cell Sorting (FACS)

Fluorescence-activated cell sorting was developed by Leonard Herzenberg in 1970 (Dangl & Lanier, 2013) and was a remarkable breakthrough in immunology and cancer biology. Relying on basic principles of flow cytometry, FACS enables separation of cell populations into sub-populations based on fluorescent labelling. To take the advantages of FACS, vectors containing one constitutively expressed reporter gene were cloned, so that cells transfected with these vectors could be sorted. Cell Sorting service was provided at BIOSS from the Toolbox team using Bio-Rad S3eTM Cell Sorter.

Cloning

Cloning, being being the fundament and one of the most important tool of synthetic biology, also vastly contributed to this year’s project.

If not otherwise stated all methods were done according to the protocol of the manufacturer.

Cloning methods

Gibson Assembly

Plasmids, that were judged to be assembled in the best way via Gibson Assembly, were generated using NEBuilder HiFi DNA Assembly Kit.

T4 Ligation

Plasmids, that were judged to be assembled in the best way via Ligation, were generated using Qiagen T4 Ligase.

Digestion for cloning

To extract desired DNA fragments for the cloning process the following protocol has been used.

If two restriction enzymes were used simultaneously the right conditions were determined by using “NEB Double Digest Finder” or “Thermo scientific double digest calculator” online tool.

The reaction was stopped after 1-4 hours by adding 10x Orange G, 6x NEB Purple or 6x Thermo Scientific loading dye to the sample. After this the sample was analyzed by agarose gel electrophoresis.

Gel extraction for cloning

Extraction of amplified DNA fragments or digested fragments from agarose gels was performed using the QIAquick gel extraction kit from Qiagen.

PCR

Amplification of DNA fragments was performed by using NEB Q5 High Fidelity or NEB One Taq polymerase.

Extension PCR

To generate the DNA fragments containing overlapping sequences for Gibson assembly an extension PCR was performed using NEB Q5 High Fidelity polymerase.

Preparation and Handling

Culture conditions

Liquid overnight cultures of E. coli strains, regardless of the fact if there were containing plasmids or not, were incubated for 12 – 16 hours in LB medium at 37°C and 200 rpm. The only exception were E. coli strains containing lentiviral transfer plasmids, these were incubated at 30°C to ensure higher plasmid concentrations. Solid cultures were incubated on 1.5% LB Agar plates at 37°C.

Selection of E. coli was performed by adding the appropriate antibiotic to the LB medium or LB plates. The following antibiotic concentrations were used: ampicillin at a final concentration of 100 µg/mL, chloramphenicol at 25 µg/mL and kanamycin at 50 µg/mL.

Davis, Leonard. Basic methods in molecular biology. Elsevier, 2012.

Glycerol stocks

To store bacterial strains 500 µL of liquid overnight culture were mixed with 500 µL of 50 % (v/v) glycerol diluted in ddH₂O. Afterwards the cryotube was inverted 4 to 6 times and placed in a -80°C freezer for longterm storage. For overnight cultures, the stock were scraped at the surface with an pipette tip and medium inocculated with it.

Davis, Leonard. Basic methods in molecular biology. Elsevier, 2012.

Preparation of chemically competent E. coli

Competent E. coli were produced using the Zymo Research Mix & Go E. coli Transformation Kit.

Transformation

Chemically competent E. coli strains were transformed by adding 2 – 10 µL of T4 ligation or Gibson assembly reaction mix containing the appropriate plasmids. Afterwards the transformed cells were incubated for 1 hour in 1000 µL LB medium containing no antibiotics, at 37°C and 300 rpm, followed by spreading on a LB agar plate containing the appropriate antibiotic resistance. The spread cells were incubated overnight at 37°C.

Subsequently, 3 - 10 colonies were picked per plate and inoculated in 3 - 5 mL of LB medium again containing the appropriate antibiotic and incubated overnight.

Davis, Leonard. Basic methods in molecular biology. Elsevier, 2012.

Plasmid preparation

The preparation of plasmids was performed using kits from Qiagen,Jena Biosience, Promega and Zymo Research, according to their provided protocol.

Agarose gels

Amplified or digested DNA samples were supplemented with the appropriate amount of 10x Orange G, 6x NEB Purple or 6x Thermo Scientific loading dye and loaded on a 1 - 2% (w/v) agarose TAE gel.

TAE agarose gels were produced by mixing agarose with 1x TAE buffer. Afterwards the mixture was microwaved until the agarose was completely dissolved. Staining of the DNA was performed by the PeqGreen DNA dye at the recommended concentration.

The gel was subjected to electrophoresis at 100 – 130 V for 15 - 30 minutes or until a sufficient separation of the bands was observable. Visualization of the separated DNA fragments in the gel was performed by an ultraviolet light imager. A NEB 2-log DNA ladder or a Thermo Scientific 1kb+ DNA ladder were included as molecular size marker.

Davis, Leonard. Basic methods in molecular biology. Elsevier, 2012.

Screening methods

Test Digestion

To screen for clones containing the right plasmid, the prepared plasmid was treated for the following protocol.

Afterwards the mixture was analyzed by gel electrophoresis and clones, that showed the right band sizes were further analyzed by sequencing.

Davis, Leonard. Basic methods in molecular biology. Elsevier, 2012.

Colony PCR

To perform a colony PCR a colony was chosen and transferred to a new LB plate, containing the appropriate antibiotic, with a pipet tip.

Afterwards the pipet tip was dipped into nuclease free water and the water was used to perform a PCR.

Innis, Michael A., et al., eds. PCR protocols: a guide to methods and applications. Academic press, 2012.Colony PCR for screening

Sequencing

Sequencing was performed for the definitively verification of plasmids. Therefore 20 µl of preperated plasmid DNA with a concentration of 30 – 100 ng and 20 µl of the appropriate primer with a concentration of 10 µM were send to GATC-Biotech. Sequencing results were analyzed using Geneious version 10.0.9.

Plate Reader

Plate reader measurements are used to quantify for example the relative expression of fluorescent proteins under different promoters. Another application is tracking and quantification of enzyme activity, which is used for Luciferase or SEAP assays (Mohamed-Bassem et al., 1987). In this project, the plate reader was used to analyze the spatial and temporal resolution of the promoter activity.

In this method, a laser is used to excite fluorescent proteins at their absorption maximum. The emitted light is measured by a photosensitive sensor at the emission maximum either detecting from the top or the bottom of the microplate. The latter is prefered for tissue cells as they adhere on the ground of a plate well. Since the measured fluorescence intensity is proportional to the amount of expressed fluorescence proteins, this method is highly suitable for promoter characterizations.

For plate reader measurements, cells were incubated under different conditions on a black 96 well plate with clear bottom in 200 µl medium. The fluorescence intensity was measured after different incubation times with a Synergy H4 from Biotek.

SEAP Assay

SEAP (secreted embryonic alkaline phosphatase) assays were performed to measure the induction of the HRE promoter.
Firstly, 200 µl of cell supernatant was transferred to 1.5 ml Eppendorf tubes. These were heated for 30 min at 65°C, followed by a centrifugation step (1250 x g, 1 min).
2 µl of 2x SEAP buffer (20 mM homoarginine, 1 mM MgCl2, 21 % (v/v) diethanolamine, pH 9.8) were filled into wells of a 96 well plate. 40 µl of the heated cell supernatant were added to the wells with 2x SEAP buffer.
The addition of 68 µl of the substrate pNPP (120 mM para-nitrophenyl phosphate dissolved in H2O) was directly followed by spectroscopic measurement for 180 min, every 30 s at a wavelength of 405 nm in a microplate reader (Biotek Synergy H4 Hybrid Reader).

SEAP Assay Analysis

The averages of absorbance were calculated for all triplicates. Afterwards, the corresponding untransfected control was subtracted from each sample and the controls were set to 0.
The slope was determined for the samples and added in the following formula to determine the relative enzyme activity:
((slope/(18600*0.5))*(10^6)*20)/80

Flow Cytometry

Flow cytometry was used to measure the induction of all promoters. All experiments were performed in triplicates.
For the analysis, cells were resuspended and collected in Eppendorf tubes. Following a centrifugation step (200 x g, 5 min) the supernatant was discarded and the cells were resuspended in 300 µl FACS buffer (10 ml PBS, 100 µl FCS).
Untransfected cells and cells with either CFP or mCherry were used for the compensation of the channels.
Measurement was performed until either a time limit of 5 min, a total event number of 1 mio or 20,000 living cells was reached.
For the experiments a Gallios flow cytometer from Beckman Coulter was used, as well as the CyAn from Beckman Coulter.

Flow Cytometry Analysis

To analyze the data measured by the flow cytometer, FlowJoTM 10 was used.
First, gates were set up for living cells in the wild-type cells, which then were gated for mCherry positive cells and these for CFP positive cells. This was done for each treatment group independently, were the gating was always applied from the corresponding wild-type control to the test sample.

Wild-type cells that were treated in the same way as the cells containing the construct of interest were used for a gating of living cells and mCherry and CFP positive cells.
An exemplary gating strategy can be seen in figure 1 and 2

Figure 1: Gating strategy for flow cytometry analysis of mCherry+ CFP+ cells under the population of living, single cells. Exemplary gating for living wild-type Jurkat cells which were treated with Forskolin (100 µM) and IBMX (100 µM) for 24h. a) Gating for living cell population. b) Afterwards, the living population was gated for single cells. c) Followed by the gating of mCherry positive cells. d) Depicted is the gating for CFP positive cells under the population of mCherry positive cells. These were however gated underneath the single cell population, considering that the wild type cells do not express mCherry. The gate then was moved under the population of mCherry positive cells.

Figure 2: Applied gates from the corresponding wild type control to Jurkat cells which contain the CRE plasmid and were treated with IBMX (100 µM) and forskolin (100 µM). The gating of the wild-type control was shown in figure 1. a) Gating for living cell population. b) Afterwards, the living population was gated for single cells. c) The mCherry positive single cells were determined and then d) gated for CFP positive cells to obtain the cells with successful expression of the construct.

T7E1 Assay

To assess insertion of small mutations T7 endonuclease 1 (T7E1) assay was performed. T7E1 specifically cleaves mismatched dsDNA. From Cas9 treated clonal cells and non-transfected cells gDNA was extracted using KAPA Biosystems Express gDNA extraction kit according to the manufacturer's protocol. From this the Cas9 gRNA target sequences were amplified by PCR and subsequently purified using PCR purification kit (Quiagen).
For each Cas9 treated clone one pure sample and a 1:1 mix with wild type gDNA PCR fragment was analyzed. The samples were melted for 10 min at 95ºC in NEB2 buffer (1x) and slowly re-annealed by switching off the heat block and letting them cool down in it for at least 2.5 h to form mismatched double strands (Fig. 3). Then 1.5 μl 1:1 dilution of T7E1 enzyme (NEB) with NEB2 (2x) was added to 300 ng annealed PCR fragment in 12 µl NEB2 (1x). The reaction was incubated 20 min at 37ºC and stopped by addition of gel loading dye on ice for subsequent analysis on ethidium bromide stained agarose gel (2 %).
As positive control served the monoallelic CCR5Δ32 mutation of HEK293T cells. If only the mix of knockout and untreated fragments is cleaved by T7E1, the mutation is present in both alleles, however, if the pure treated sample is cleaved only one allele was mutated.