Using a dual-fluorescence dual-plasmid quantification system to rapidly and robustly measure the performance of synthetic transcription factor - promoter pair

Introduction

Transcription factors and promoters are the basis of the genetic circuit. Compared with prokaryotic promoters, the mechanism of action of eukaryotic promoters is complex(Struhl, 1999), which increases the difficulty of new design and transformation. A limited number of eukaryotic transcription factors and promoters are currently bottlenecks in the design and implementation of complex gene circuits in mammalian cells to meet a variety of clinical or industrial applications.

In order to obtain a sufficient number of orthogonal synthetic transcription factor-promoter pairs, it is often necessary to carry out a certain number of constructions and select candidates with an excellent performance from the complex construction, which requires us to develop a robust, fast and cost-controllable method suitable for large-scale screening. Therefore, testing using transient transfection methods is an effective method.

However, since transient transfection experiments will introduce a heterogeneous number of plasmid copies into cells(Cohen et al., 2009), the differences between experiments will be expanded compared to stable expression cell lines. We found that a reasonable way to distinguish successfully transfected cells is the key to maintaining test robustness while quantifying the promoter,. As an example of a group of experiments in which a transcription factor and its corresponding promoter controlling a downstream reporter gene, are co-transfected, transient transfection will result in a high degree of heterogeneity in the fluorescent expression of the cells. If it is not possible to gate out cells that have not been successfully transfected by some means, the transfection efficiency of each change in each experiment will make these untransfected cells have a catastrophic effect on the statistical results (Fig. 1a) however, by introducing an additional fluorescent signal, we could clearly distinguish cells that have been successfully transfected (Fig. 1b) . This method has important implementation value

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However, in transient transfection, the introduction of additional plasmids may result in not all cells being able to take up all of the components of the plasmid set. Thus, the use of such a method does not effectively distinguish whether those cells are successfully transferred to all of the plasmid groups based on the presence or absence of internal reference fluorescence. However, the success rate of co-transfection can be significantly improved by reducing the amount of plasmid used.

So we developed a dual-fluorescence dual-plasmid (DFDP) quantification system (DFDP), keeping only two plasmids that allow us to confidently perform a co-transfection experiment. Simultaneously, the P2A element couples the expression of the internal reference fluorescence with transcription factor (TF), so that we can gate out the cells that have not been successfully transferred into the plasmid by distinguishing the presence or absence of one of the fluorescences. Thereby we can minimize the interference caused by the difference in transfection efficiency between the repeats. The DFDP quantification system thus helps us measure transcription factor-promoter pair strength quickly and robustly.

Here we summarize the comparison of using a 2A element to construct the DFDP system relative to co-transfecting an internal reference plasmid (Table. 1).

Table

Method

The Desing of DFDP system

Based on theoretical assumptions (see Supplementary Materials for details), we designed a dual-fluorescence-double plasmid (DFDP) system containing two plasmids (Fig. 2). Spectral overlap problems in multicolor fluorescence signal measurements can interfere with the effectiveness of the measurement. Thus, we used mCherry and EGFP here as two dual fluorescent signals because of the minimal overlap of their spectra.

figure2

Optimize DHDP plasmid vector backbone easy molecular cloning

Since in our design, the P-TF plasmid requires the construction of a fusion protein, we specifically designed p2Y-C-mCherry-P2A as the basic skeleton for P-TF construction. We designed multiple cloning sites (MCS) at the mCherry-P2A backend. We inserted a BamHI site after the last Pro on the C-terminus of P2A. TF can be inserted directly after digestion with BamHI and HindIII. In addition to the restriction enzyme digestion method, we inserted two BbsI sites after the 3' end of BamHI and 5' of HindIII to meet the potential requirements of the Golden Gate cloning method, and the restriction sites were specially designed. Make the same as the cleavage site of BamHI and HindIII. A stop codon was also inserted between the two BbsI sites, such that the p2Y-C-mCherry-P2A backbone without the insertion of TF was normally terminated after transcription initiation. Therefore, the p2Y-C-mCherry-P2A plasmid can be used as a control plasmid (P-Ctrl). We also optimized the base sequence after the 5' end of BamHI and the 3' end of HindIII to reduce the hairpin structure and make it very easy to use when using ClonExpress or Gibson assembly. Although insertion of the BamHI site will result in three amino acids of the N-terminal residual Pro-Gly-Ser of TF, as far as we know, it does not affect the function of SynTF. You can also use the ClonExpress or NEBuilder cloning method to remove the BamHI site during assembly, leaving only one Pro.

Molecular cloning of the plasmids based on this specially optimized backbone system enables DFDP system has never been easier and more convenient.

We use the ClonExpress (Vazyme #C115) or the standard Biobrick assembly for all plasmid assembly.

Download the sequence file to view our MCS design

Cell line

293T (ATCC #CRL-3216) were cultured in DMEM (Gibco #11965092) supplemented with 10% fetal bovine serum (Gibco # 10437036), 1x Penicillin-Streptomycin.

Transient transfection

Transfection was performed using the cost-effective cationic polymer polyethyleneimine Max (Polysciences #24765). All cells transient transfection experiments were performed in 24-well plates. Cells were plated one day earlier and transfected the next day at 80% cell confluence. For a well, 1.5 µl 1 µg/µl were added into 23.5 µl Opti-MEM (Thermo Fisher Scientific #11058021). Total 500 or 450 ng plasmid was added into 25 µl Opti-MEM. 5 min later PEI-Opti-MEM and DNA-Opti-MEM mixture were mixed and place at room temperature for 10 min before adding into the well.

The amount of plasmid used for activating-, silencing-, NIMPLY-form promoter tests are slightly different (Table. 2).

Flow Cytometry (FCM)

All the flow cytometry data is acquired by FACSJazz (BD Biosciences). After 48 hours of transfection, the cells were fixed with 4% paraformaldehyde for 15 min, and washed twice with PBS before analysis, keeping dark conditions as much as possible during the operation. d2EGFP are recorded under the excitation laser of 488 nm with a 530/40 filter. mCherry is recorded under the excitation laser of 561 nm with a 610/20 filter. The photomultiplier tube (PMT) amplification values for FSC, SSC, EGFP, mCherry channel are 28, 28, 28 and 44 separately. The spectral overlap between EGFP and mCherry is very small, meanwhile EGFP and mCherry were activated by 2 lasers separately, no compensation is needed.

Data analysis

Three biological replicates were performed for each experiment. FCM data are analyzed by FlowJo 10 (TreeStar). 3 basic hierarchical populations are gated: FSC-SSC → FSC-Trigger pulse width → 488_530/40-561_610/20. Successfully transfected cells are gated by mCherry positive for the same group. MFI represents the median fluorescence intensity of EGFP in the top 50% of cells with the strongest expression of d2EGFP. The different biological replicates were normalized using the fluorescence intensity of mCherry. The Fold value is calculated according to the formula in the theoretical hypothesis (see theoretical assumptions for details). All statistical analysis was performed with Prism 7 (Graphpad).

Results

Testing activating-form promoters using DFDP system

We used the artificial zinc finger protein (Synthetic Zinc finger, SynZF) (Khalil et al., 2012; Lohmueller et al., 2012) to construct three mammalian-adapted DNA binding domain (DBD) of ZF21.16, ZF42.10, and ZF43.8, based on an optimized iGEM 2017 Fudan's standardized synthetic transcription factors (SynTF) – synthetic promoter (SynPro) construction paradigm. The testing using DFDP system showed that the ZF21.16-, ZF42.10-, ZF43.8-VP64 had good activation characteristics (Fig. 3), and the activation magnification ratio ranged from 51 to 454 fold. Among them, 8×ZF21.16-minCMV had the largest activation ratio (454 fold), while 8×ZF43.8-minCMV had the smallest activation ratio (51 fold). The difference between them is mainly due to the relatively high background expression of 8×ZF43.8-minCMV. Thus we believe that 8×ZF21.16-minCMV is a relatively better promoter (in our opinion on excellent promoters, see the discussion section).

figure3

Testing the orthogonality between synthetic DBDs using cross-paired DFDP system

Cross-paired DFDP can be used to verify promoter orthogonality by using the co-transfection strategy of P-TF with both corresponding or uncorresponding P-Pro. Since there are 5 aSynTF-aSynPro pairs, 5 x 5 = 25 sets of experiments are required. The intensity of d2EGFP under different conditions was examined, and it was observed that the group on the diagonal (i.e., the correctly paired group) had the most significant expression (Fig. 4), thus indicating that the 5 pairs of aSynTF-sSynProdui pairs were orthogonal to each other. The advantage of using the activated promoter for DNA binding domain orthogonality testing over the use of sSynTF-sSynPro testing is that there is a difference in the basal expression of sSynPro, so more normalization is required to rule out differences in basic expression. Abnormal results.

figure4

Testing silencing-form promoters using DFDP system

Based on the iGEM 2017 Fudan (see the improvement page), we adjust the position of the sSynTF corresponding response element (sTF REs) from the 3' end of the promoter to the 5' end, and replace the original pSV40 with a stronger CMV promoter to serve as the conPro. Then we constructed a new generation of inhibitory transcription factors (Fig. 5). Such a construction helps to attenuate the interference of sTF REs on the under expression of conPro. Tests using the DFDP system show that ZF21.16-, ZF42.10-, ZF43.8-KRAB have good inhibition characteristics, and the magnification ranges from 9 to 13 times (Fig. 5). The reason why aSynTF-aSynPro differs by a hundred fold and the difference of only about tenfold here does not mean that our sSynPro function is not good. Rather, the basic expression of aSynPro is particularly weak, and it is close to the minimum of our flow cytometry detection range at the time of testing.

figure5

Testing NIMPLY-form promoters using DFDP system

In addition to traditional activating- and silencing-from promoters, we have also designed a novel NIMPLY-form promoter. Adding multiple REs corresponding to the activating- or silencing-form transcription factors at the 5' and 3' ends of minCMV, respectively. The dominant inhibition can be achieved by (1) KRAB recruitment of transcriptional inhibitors can inhibit promoter expression (Margolin et al., 1994) (2) Since sTF REs is located downstream of the promoter, when the inhibitory transcription factor binds to the REs, it can cause steric hindrance and enhance the ability of transcriptional inhibition by inhibiting the forward movement of RNA polymerase. This type of promoter exhibits the logical selectivity of NIMPLY under conditions of overexpression of aTFs and sTFs. That is, it can be expressed only in the presence of aTFs and the absence of sTFs. 8xZF21.16-minCMV-2xZF43.8 can only be activated in the presence of ZF21.16-VP64 and in the absence of ZF43.8-KRAB. 8xZF43.8-minCMV-2xZF21.16 can only be activated in the presence of ZF43.8-VP64 and in the absence of ZF21.16-KRAB (Fig. 5). It can be observed that in the group transfected with both aTF and sTF, the expression of NIMPLY-form promoter is increased a little because although sTF is dominant, it can slightly reverse sTF in the condition of overexpressed aTF.

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Discussion

What kind of promoter is a good promoter?

A promoter is not always the higher expression the better performance. Just as in some experiments that require well control of protein expression. Scienctist would even insert no promoter on the viral vector and rely solely on the LTR element to maintain the protein at a weak expression(Cai et al., 2007). It was also observed in our SynNotch experiment that comparing with using CMV to control SynNotch, by using a wea promoter of PGK can significantly reduce the basal expression (data not shown). For inducible promoters, we always expect that the activating-form promoter maintains very low background expression when not activated, and produces differential expression when activated. Note that we do not believe that an activating-form promoter is activated as much as possible after being induced, indicating that it works well, and more importantly is whether the background expression of its background is low enough and whether it has sufficient tunability. Similarly, for silencing-form promoters, we always want to maintain stable expression under uninhibited conditions and can be shut down as much as possible after being inhibited. Similarly, we are more concerned about the stability of expression and the degree of inhibition.

Factors that interfere with the DFDP system

The d2EGFP intensity was used as an indirect characterization of promoter strength in our test system, but it should be noted that d2GFP stability may vary by cell type. Therefore, the comparison of d2EGFP intensity across cell types may not truly reflect differences in promoter strength in different cell types. Only the same cell type can be used as a common chassis for the promoter test. Under these conditions, the d2EGFP intensity driven by different promoters can be compared to reflect the difference between the strengths of the promoters. At the same time, for mammals, even the same constitutive promoter, its differential performance in different cell lines is still very significant, for example, the CMV promoter is very active in some cell types (such as 293T cells). But in some other cell types (such as mesenchymal stem cells) it may be quite weak (Qin et al., 2010). In our measurement protocol, we used 293T cells as chassis cells to screen for functional synthetic transcription factor-promoter pairs. If you need to migrate the potential synthetic transcription factor-promoter pair obtained from the initial screening to other cell lines, we strongly recommend you to use a commonly used transcription factor-promoter that works well in this cell line as a positive control. And then repeat the DFDP test on your target cell line to qualitatively verify the synthetic transcription factor-promoter pair again.

The success of multicolor flow cytometry depends on many key hardware factors: first, the shape and position of the excitation laser beam; second, the choice and quality of the optical filter; and third, the sensitivity and resolution of photoelectron detection (Perfetto et al., 2012). The intensity and sensitivity of the excitation fluorescence of different flow cytometers have a certain degree of difference. To ensure that the data between each test is quantitatively comparable, the same flow cytometer should be used for the test. Quality control is required prior to each analysis even if using the same flow cytometer. If a non-fixed flow system is used, automatic/manual calibration of the flow is required. Make it the optimal position. Dual fluorescent signals should be received using the same optical channel (filter configuration) and should ensure that the sensor-photomultiplying tubes (PMTs) for each channel set the same gain for each test. An unfixed PMT setting will destroy your experiment. If a flow cytometer with non-fixed PMT is used, it is important to select and set an appropriate PMT value for the subsequent experimental needs during the initial experiment. We recommend the following method. When you first start your synthetic transcription factor-promoter pair measurement, you could set up a strong double positive cell to determin the proper PMT setting. For Example, we use CMV-mCherry and CMV-d2EGFP in our experiment and add just the two channels to ensure that the brightest group of cells are within the maximum detecting range of the flow cytometer.

Transient transfection efficiency will interfere with comparability between different experiments. Different transfection reagents, reagent dosages, as well as cell density at the time of transfection, cell status, and the duration of transfection can significantly affect the efficiency of transfection. So you should control these variables as much as possible during the test. For example, try to use the same batch of transfection reagents; optimize the transfection efficiency and then use the same amount of transfection reagent and fix the ratio between the DNA and the transfection reagent; use the same transfection density; use the cells with low passage number; and fix the cell at the same time after transfection for future Flow analysis.