Determing the plasmid copy number

The foundation of multi-plasmid framework is the ability to determine plasmid copy number (PCN) per cell. Our approach to count the PCN in the cell is built upon absolute quantitative PCR.

Why we chose a specific set of primers for PCN determination?

Commonly, gene used for PCN evaluations in qPCR is an ampicillin resistance gene (bla). In our case, the bla gene for plasmid number determination was not used, as the SynORI multi-plasmid framework employs a number of plasmids with different selection system gene circuit parts. Instead, the origin of replication was barcoded with distinct sets of primers (named qPCR) for different groups of origin copy number determination. This enables to determine the desired plasmid group copy number, when working with multi-plasmid systems.

Using cell lysates instead of extracted DNA

Since working with extracted genomic, plasmid or total DNA incorporates an error which depends on the extraction efficiency, we decided to work with specially prepared cell lysate suspensions, skipping the extraction step. Furthermore, dxs gene from chromosome was ligated into pUC19 plasmid for less complicated standard preparation.

Results

First of all, it was concluded that primers, designed for calculations, are appropriate for absolute quantitative PCR by measuring the efficiency of PCR amplification reaction. Both genes have almost ideal amplification efficiency and required no further optimization.

Figure 1.

Amplified qPCR product were verified using agarose gel electrophoresis.

Figure 2. Verification of products amplified during qPCR.

Figure 3. Melting curves of amplified qPCR products used for PCN determination.

Next, standard curves were generated for dxs and qPCR plasmid gene copy number determination according to the protocol written by our team (click here?).

Figure 4.

The fit of the linear regression model was satisfactory; the coefficient of determination (R2) was more than 0.999 for all standard curves. As amplification efficiencies and generated standard curves were ideal for both genes, the curves and earlier mentioned protocol were used for all further PCN determination experiments.

Refference:

Plotka M, Wozniak M, Kaczorowski T. Quantification of Plasmid Copy Number with Single Colour Droplet Digital PCR. Doi H, ed. PLoS ONE. 2017;12(1):e0169846. doi:10.1371/journal.pone.0169846.

Anindyajati, Artarini AA, Riani C, Retnoningrum DS. Plasmid Copy Number Determination by Quantitative Polymerase Chain Reaction. Scientia Pharmaceutica. 2016;84(1):89-101. doi:10.3797/scipharm.ISP.2015.02.

Plasmid copy number control

Flexible copy number control is the core of our framework, which is based on re-engineered ColE1 origin of replication.

Base of SynORI framework - ColE1 replicon

ColE1 plasmid replicon is based on two antisense RNA molecules: RNA I and RNA II.

Transcript of RNA II forms a RNA-DNA duplex and acts as a primer for DNA polymerase and for that reason is often called replication initiator.

During the transcription of RNA II several different secondary structures can form. Part of the structures are susceptible to the binding of RNA I – a shorter antisense version of RNA II. The interaction between RNA I and RNA II start upon formation of kissing-loop pairs between their anti-complementary secondary structures. If the kissing complex persists 3’ end of RNA I starts forming a zipper-like duplex with complementary single strand RNA II region. This results in replication inhibition, because primer cannot be formed anymore, which is why RNA I is often called replication inhibitor.

The main reasons why we have chosen ColE1 as base for SynORI framework was:

• It is a light system consisting of only two regulatory RNA molecules
• It is biochemically and mathematically well characterized
• Kissing-loop complex formation kinetics allows to predict plasmid group compatibility.

Picking the control type

It immediately becomes clear that in order to control the copy number of a plasmid one could simply change RNA I promoter. But there is a reason why it was never done before!

As RNA I and RNA II are two antisense molecules, changes made to sequence will affect both of them. Location of RNA I promoter coincides with the RNA II secondary structures, which are crucial to replication primer formation.

Even if one could somehow manage to change the RNA I promoter to another one without disabling replication initiation, it would still not be a convenient because picking another promoter would require a large pool of resources every time.

For that reason we have decided not to change or modify RNA I promoter inside the wild type ColE1 origin of replication, but rather to disable it completely and place a copy of it next to RNA II.

Disabling the RNA I promoter

The main problem of inactivating RNA I promoter is that precautions must be taken in order not to change critical secondary structures of RNA II.

We have first acquired a priority mutation list from literature which analyses RNA polymerase binding affinity to -10 and -35 promoter structures and its dependence on point mutations, with mutations causing the largest decrease in affinity being in the top of the list.

Then, we compiled a simulative algorithm which compared every possible combination of -10, -35 mutations and then compared them to predicted RNA II secondary structures made by CoFold, a thermodynamics-based RNA secondary structure folding algorithm that takes co-transcriptional folding into account. We have picked replicon mutants prioritizing:

• Mutants that have unchanged RNA II secondary structures.
• Mutants that are highest in mutation priority list (lowest RNA polymerase affinity).

Tailoring the copy number control

Once RNA I promoter is disabled in the ColE1 origin of replication, it can be moved to another plasmid location and used as a separate unit. Also, RNA I promoter can now be changed without damaging the replication initiation.

RNA I, and consequently, the copy number of a plasmid can now be placed under virtually any signal pattern required.

We have first showed this by placing RNA I under a series of constitutive Anderson promoters and an inducible Rhamnose promoter.

We can now flexibly control the copy number of a plasmid! What comes next?

Multiple plasmid groups

Multi-plasmid framework would not be much without multiple plasmids. We have equipped our synthetic origin of replication with specific sequences to create unique plasmid groups.

PAs RNA I and RNA II interact mainly with the three stem loops that form kissing complexes, we have decided to use this feature to our advantage in order to engineer different plasmid groups by adding unique, group-specific sequences to RNA I and RNA II stem loops.

The specific sequences were acquired from Grabow et al., where they have screened a large number of different RNA kissing stem loop complex combinations. They have derived a table of different loop sequences that only bind with each other but do not have any cross interaction to the following loop sequences in the list.

The inactivation and transfer of RNA I gene away from RNA II allows us to use different sequences for RNA I and RNA II molecules that are not necessarily ideal complements of each other.

Since there are three stem loops responsible for RNA I–RNA II interaction for each of the plasmid group we have decided to:

1. Use two different unique sequences in the first two RNR I and RNR II stem loops, in order to maximize same group specificity.
2. Keep RNA II unchanged for the third loop but change it in the RNR I by adding either G/C mutations to RNA I (GC type RNA I) or making RNA I completely non-complement to RNA II (NC type RNA I).

According to literature RNA II secondary structures at third loop are very sensitive to any mutations and has a high chance of ruining the replication initiation. For this reason we did not want to introduce new specific alterations into the third loop of RNA II sequence. Just because we chose not to interfere with the third loop of RNA II, we could not leave RNA I gene unchanged. If every group would have the fully compatible third loop, the background cross-group inhibition would be too large.

So now we have 5 different RNA II genes corresponding to groups A, B, C, D and E.

Also, we have 10 different RNA I alternatives: A , B, C, D, E with each having a version of either G/C or NC mutations.

These different plasmid groups can then be co-maintained in cell with a specific, pre-selected copy number. Copy number control principle is the same for every group, but each group is only specific to its own group.

Foto aprasymas it anim id est laborum. Sed ut perspiciatis unde omnis iste natus error sit voluptatem.

Global copy number regulation

Rop protein

Now that we have figured out and engineered a way to regulate plasmid copy number in a group - specific fashion the only control element that is missing is a way to control every group at the same time.

Recall, we have used ColE1 replicon as base of our system. And it has given us a perfect hint on how to achieve our current objective. Wild type ColE1 replicon codes a small homodimeric, four-helix bundle protein called Rop (also known as repressor of primer).

Foto aprasymas it anim id est laborum. Sed ut perspiciatis unde omnis iste natus error sit voluptatem.

Literature shows that Rop secondary structure specific, rather than sequence specific. What that means is that rop recognises RNA I - RNA II kissing loop complex.

Foto aprasymas it anim id est laborum. Sed ut perspiciatis unde omnis iste natus error sit voluptatem.

When Rop binds to secondary structures, it increases the binding affinity of RNA I and RNA II and consequently - replication inhibition.

As our framework approach is based on specific RNA sequence binding, having a Rop protein in our system is equivalent of having a global copy normal regulator. In theory, the protein should bind to every complex despite the specific interactions of each group, because the binding geometry should stay similar in each case.

Foto aprasymas it anim id est laborum. Sed ut perspiciatis unde omnis iste natus error sit voluptatem.

We have designed Rop protein with an anderson promoter and showed that it can reduce the copy number of single plasmid, and multiple plasmids non-specifically.

Selection system

Split antibiotic – 2 plasmids system

One of the essential parts of synthetic biology are plasmids. However, bacterial plasmid systems require a unique selection, usually an antibiotic resistance gene, to stably maintain the plasmids. As the number of different plasmid groups used in a single cell rise, the need for more selection markers grows. In addition to raising the issue of biosafety, the use of multiple antibiotic resistance genes destabilizes the functionality of the cells. To address this problem a protein granting the resistance to aminoglycoside family antibiotics, called amino 3'-glycosyl phosphotransferase (APH(3')), was split into two subunits by Calvin M. Schmidt et al.

According to the obscure guidelines we split an unmodified neo gene sequence between 59 and 60 amino acid residues. Two subunits were termed α-neo and β-neo. Furthermore, we added additional termination codon at the end of an α-neo fragment for the translation to stop. No other start codons were included into the β-neo subunit as the gene was designed for toehold switch system. Despite the fact that β-neo subunit had no start codon, the split antibiotic system worked perfectly when coupled with a standard promoter and a ribosome binding site (BBa_K608002). Consequently, a split antibiotic resistance gene provides a selection system to stably maintain two different plasmids.

Figure 1. Split neo gene principle scheme by M. Schmidt et al.

Toehold switches – 4 plasmids system

In order to increase the capability of our selection system, we reasoned that a split antibiotic system should be put under a transcriptional or translational control. A. A. Green et al. presented wide range of de novo synthesized dynamic riboregulators, called toehold switches, which take advantage of RNA-mediated linear interaction to initiate RNA strand displacement. A toehold switch contains two parts: a ribosome binding site and a linker sequence, both of which are sequestered by a secondary RNA stem loop structure. The linker sequence has a start codon and functions as a link.

Figure 2. Principal toehold switch scheme by A. A. Green et al.

Although the linker sequence adds additional 10 amino acid residues to the peptide, we reasoned that it will not affect the function of split antibiotic. Toehold switches are unlocked when an RNA trigger binds to the 5’ end of the toehold and initiates RNA duplex formation, which releases the locked RBS and reveals linker start codon. We concluded, that if the toehold sequences were added in front of α- and β-neo gene fragments, the translation would require trigger RNA to initiate protein synthesis.

Toeholds and their corresponding triggers design sequences were used as described by A. A. Green et al. with some modifications. First of all, it is important to note, that a “scar” which is made between biobrick prefix for protein coding sequences and suffix, contains a termination codon at the 3’ end. Therefore, it was necessary to use the other form of prefix for α- and β-neo genes, as the translation proceeds from one biobrick to another. Furthermore, seeing that the “scar” produced when joining two biobricks is 8 base pairs, we included an additional T nucleotide at the end of linker sequence to ensure the translation stays in frame to the α- and β-neo genes.

KA PARASYTI

We constructed a system, which includes two toehold riboregulators (termed toehold 1 and toehold 2) upstream of α- and β-neo genes in two different plasmids. The corresponding activating RNA triggers (name trigger 1 and trigger 2) were placed into additional two plasmids under constant expression. All the parts used together complete a 4-plasmid selection system - two distinct trigger RNAs are expressed by two different plasmids in order to unlock the translation of toehold controlled α- and β-neo peptides to form a complete amino 3'-glycosyl phosphotransferase. For this reason, if one plasmid is lost, the end product – α/β dimer APH(3') is not formed, therefore bacteria lose their antibiotic resistance.

Phage control – 5 plasmids system

The SynOri selection system circuit could be expanded by including additional transcription factor which induced the transcription of previously described RNA triggers. The fifth plasmid would house a transcription factor for the initiation of whole system. Phage modified promoter is perfect for this task, as it is activated by cI lambda peptide and repressed by cI 434 peptide with minimal leakage. Both of the RNA triggers - 1 and 2 - were placed under control of phage modified promoter. Furthermore, downstream of the trigger gene we included cI 434 repressor under constant expression to ensure minimal leakage of the promoter. The fifth plasmid was built to constantly express cI lambda – the activator of phage promoter. In the absence of this plasmid, the gene circuit cannot function, as the trigger RNA transcription is repressed by constant cI 434 expression and toehold switches lock the translation of α/β APH(3'). When the final component of the circuit is present, the cI lambda activator enhances the transcription of both RNA triggers. The transcribed triggers then unlock the translation of α/β neo peptides which form an active protein and confer the resistance to aminoglycoside family antibiotics.

Figure 4. The schematic representation of SynORI 5 plasmid selection gene circuit.

Results

2 plasmids selection system

The minimal SynORI selection system, designed for maintaining two plasmids in a cell, was validated by coupling two parts of the split antibiotic coding sequences - α-neo BBa_K2259018 and β-neo BBa_K2259019 - to constant expression promoter in two pSB1C3 plasmids.

Figure 5. Results of two plasmid co-transformation containing constantly expressed α and β split kanamycin antibiotic resistance gene. The control groups only had one or no antibiotic resistance gene subunits expressed.

4 plasmids selection system

In addition to split antibiotic system, we were able to co-transform 4 plasmids using only kanamycin as the selection marker by implementing a riboregulatory system upstream of the previously described divided neo gene. We employed toehold switches, an orthogonal translational regulation devices, to lock the protein synthesis of the corresponding split subunits (parts: BBa_K2259034 and BBa_K2259035). Furthermore, constantly expressed RNA trigger sequences (parts: BBa_K2259038 and BBa_K2259040) were added into additional two plasmids to complete the 4 plasmid genetic circuit. Our team has developed and optimized an electroporation protocol to carry out our transformation needs.

Figure 6. Results of four plasmid co-transformation containing constantly expressed toehold locked α and β split kanamycin antibiotic resistance gene and trigger RNA sequences.

It is important to note that, in contrast to our SynORI framework, all four plasmids were the same (pSB1C3) - it had identical replicons. As a result, the selection system became unstable after long period of growth due to replicon cross-interaction. Nevertheless, as you can see from Table 1., carrying out similar experiments with 4 different antibiotics produced zero bacterial growth!

Table 1. Experimental data of 4 plasmid electroporation results.

5 plasmids selection system

The hallmark of SynORI selection framework is genetic circuit which enables 5 plasmids to be maintained in a single cell using only one selection marker. That goal has been reached by employing modified phage promoter (BBa_I12006) to construct signal activated trigger RNA transcription devices (BBa_K2259042 and BBa_K2259043) that are controlled from the fifth plasmid.

Figure 7. Result of SynORI 5 plasmid selection system electroporation. The 4th field includes all the framework parts. 1 - No trigger 1 (control). 2 - No trigger 2 (control). 3 - No lambda activator plasmid (control). 4 - Full System: lambda activator plasmid; toehold 1 α-neo; toehold 2 β-neo; trigger 1; trigger 2.

The co-transformants were grown for 24 hours for the first colonies to appear, as all the plasmids had identical replicons (pSB1C3). Cell were sensitive and could not be inoculated into liquid LB medium containing standard kanamycin concentrations. Further optimizations with lower antibiotic quantities and different replicons are needed. Nevertheless, SynORI selection framework enabled bacteria to sustain 5 different plasmids with identical origins of replication in the presence of one selection marker, contrary to using a batch of different antibiotics as described earlier.

References

Stable Maintenance of Multiple Plasmids in E. coli Using a Single Selective Marker

Calvin M. Schmidt, David L. Shis, Truong D. Nguyen-Huu, and Matthew R. Bennett. ACS Synthetic Biology 2012 1 (10), 445-450 DOI: 10.1021/sb3000589

Active partitioning system

SynORI framework gives the opportunity to have low copy plasmid groups, yet in order for them not to be lost during cell division, there must be a mechanism that actively keeps plasmids in the cell.

We have looked into different active partitioning systems and first chose to characterize and apply a Staphylococcus aureus type II plasmid segregation system to our framework. Yet, after a lot of consideration we have decided to search for alternatives. The main reason was that partitioning system of S. aureus is rather large, almost 49 kDa, as it codes two large proteins for segregation.

We have then stumbled upon a described pSC101 plasmid stability region which is a lot different from its counterpart. It does not seem to encode any protein but rather contains a binding site for DNA gyrase (Wahle and Kornberg, 1988). Most importantly, the regulatory region is only 400 base pairs long.

It has been showed that pSC101 plasmids with partial deletions of stability region have decreased supercoiling and are extremely unstable. This has led to the proposal that gyrase-generated negative supercoiling establishes a DNA conformation which enables plasmids to interact with E. coli structures and distribute them to daughter cells during cell division (Miller et al., 1990)