Cell‑free synthetic DNA is emerging as a compelling alternative to plasmid‑based DNA synthesis, for applications like mRNA therapeutics, viral vectors, and genome engineering. For this article, we spoke with Ribbon Bio’s product manager Lukas Leiendecker about the difference, and how teams can transition from plasmid‑centric pipelines to hybrid or cell‑free‑first methods.
Introduction
One of the differences between a traditional cloning workflow and cell-free DNA synthesis is the number of steps involved. With the cell-free approach we use at Ribbon Bio, cloning is eliminated and the DNA is in vitro transcription (IVT) ready. As covered in a previous article, cell-free DNA has unique advantages for mRNA therapeutics. We dive into more detail in the interview below. Transcripts from our interview with Lukas are below and are interspersed with additional commentary.
How do you explain the difference between cell‑free DNA and plasmid DNA to a scientist starting a new program?
Traditional plasmid workflows rely on cloning and amplification in bacteria, which introduces biological constraints into the manufacturing process. These constraints can show up as instability of your desired DNA molecule or toxicity of the encoded product to the bacterial host. They also lead to longer timelines from the initial design of your sequence of interest (SOI) to the final, sequence‑perfect clone you need for your program.
On top of that, working in a biological system introduces impurity risks. Endotoxins are a big concern later on for CMC, and residual proteins have been shown to increase the risk of downstream application failures.
💡Key point: For scientists, a key shift is to stop thinking of DNA as something that must come out of bacteria and to instead view it as a synthetic reagent that can be built entirely in vitro.
When you talk about toxicity, what kinds of molecules or proteins are problematic in plasmid‑based systems?
There are two main categories that come to mind. The first are the classic cytotoxic proteins, for example virus-derived proteins that vaccine developers might want to express. These can be harmful to the bacterial host and thus hard to assemble in bacterial systems.
The second category is more subtle: regulatory elements that are not toxic per se, but in the context of a bacterial cell can cause deregulation of gene regulation and ultimately bacterial cell death. These might be elements you need for genome editing or complex control of expression in mammalian systems, but they interfere with bacterial regulatory networks.
Our cell‑free DNA allows us to handle both classes, because there is no living host involved in the assembly process that bottlenecks the tolerated sequence space.
You mentioned downstream failures caused by impurities. Can you give an example of how that shows up in practice?
A very hot topic right now is mRNA therapies. To produce an mRNA therapy, you typically start with a DNA template that you feed into an in vitro transcription (IVT) reaction. IVT is delicate and can fail at several points.
One well‑described failure mode is when the DNA template is still occupied by DNA‑binding proteins that came from the plasmid production process. If DNA‑binding proteins are attached, the IVT reaction often works much less efficiently.
Because our cell‑free process eliminates DNA‑binding proteins from the start, we’ve seen and heard strong positive feedback: our linear, cell‑free DNA templates perform at least as well as, and in many cases better than, traditional plasmid templates in IVT reactions.
💡Key point: If you are developing mRNA therapeutics, template quality quickly becomes a limiting factor: protein adducts, DNA damage, and endotoxin carried over from plasmid preparation can depress IVT yields and complicate purification.
For a beginner, where do these DNA‑binding proteins come from, and why do they matter?
Think of a bacterial cell as an individual organism constantly regulating DNA accessibility and function in response to external stimuli like temperature, pH, and other environmental factors. It does this using DNA‑binding proteins, especially transcription factors, which bind to different spots on the DNA to regulate gene expression.
When you run plasmid fermentations, you operate at a specific temperature, pH, and set of conditions. Those conditions influence how the DNA is occupied by DNA‑binding proteins. After purification, some of those proteins can remain associated with the plasmid DNA.
If you remove the living organism entirely, as in our cell‑free process, there is simply nothing in the system that can bind in that way. That’s why the resulting DNA is much cleaner and more predictable for sensitive downstream steps like IVT.
💡 From a mechanistic standpoint, plasmid DNA emerging from E. coli is the product of a highly regulated chromosomal environment, in which transcription factors and nucleoid‑associated proteins constantly bind and unbind DNA; even after purification, some of these proteins remain attached and can interfere with enzymatic reactions that expect a “naked” DNA template.
What kind of controls do you use in cell‑free workflows to monitor variability and performance, for example in IVT?
A common strategy is to include a positive control construct that’s easy to read out in a downstream assay using fluorescence or luminescence. For instance, if you’re developing an mRNA‑based cancer therapy, you might have 10 candidate vaccine genes plus an 11th construct encoding luciferase.
You run this luciferase construct through the entire process, including our synthesis, IVT, and in vivo testing. If the mice you dose with the luciferase construct show a clear luminescence signal, you know the entire workflow functioned correctly. If that control fails, you go back and troubleshoot each step to identify where performance was lost.
This type of control can be used with both plasmid and cell‑free workflows, but having a cleaner DNA input makes interpreting these controls more straightforward.
For developers, pairing experimental templates with a robust reporter control—such as a luciferase mRNA whose in vivo signal integrates performance across synthesis, IVT, formulation, and delivery—offers a practical way to benchmark new template formats like cell‑free DNA against familiar plasmid‑derived standards.
Where do plasmid‑based approaches still make sense, and where do you see clear advantages for cell‑free DNA?
Plasmids still make sense in environments with deeply established infrastructure built around intracellular propagation in bacteria or other cells, or where workflows are explicitly configured around circular DNA molecules. Academia is a good example, where plasmid‑based tools are entrenched and adoption cycles are long.
In more cutting‑edge biotech and biopharma companies, we already see cell‑free DNA outperforming plasmids when sequence complexity is high and when teams need cleaner DNA inputs for downstream applications. Beyond the technical benefits, there are regulatory advantages too. One customer told us that switching to cell‑free DNA saved them 16 pages of documentation in their regulatory filings, which translates directly into time and cost savings.
So plasmid DNA is still the “gold standard” for many applications today, but there are now clear areas like for IVT templates, genome engineering, and complex constructs where cell‑free DNA provides very significant gains.
In practice, this means most organizations will continue using plasmids where tooling and costs are optimized, while selectively introducing cell‑free DNA where it clearly unlocks value. One expample is when regulatory teams want to avoid antibiotic resistance markers and host‑cell DNA, or when R&D needs difficult constructs that repeatedly fail in E. coli.
How does cell‑free DNA compare to plasmids when it comes to expression performance?
Customers often worry that they might have to trade expression performance for the benefits of cell‑free DNA. Our typical approach is to run a proof‑of‑concept (POC) study: we benchmark their established plasmid “gold standard” workflow head‑to‑head against our linear cell‑free DNA.
What we see in those POCs is that customers do not have to sacrifice performance. They retain comparable or better expression while gaining sequence flexibility and simpler workflows. They no longer have to design their entire process around the living organism as a bottleneck, and can instead implement the workflow as originally envisioned. That realization is often the tipping point that makes them comfortable switching to cell‑free approaches.
💡 Published comparisons of cloning‑free synthetic DNA, including rolling‑circle‑amplified templates, demonstrate that carefully designed linear or minicircle constructs can drive enzyme or protein expression comparable to plasmids, suggesting that performance concerns can often be addressed empirically with targeted benchmarking studies.
Why are certain sequence types more difficult in bacteria, and how does cell‑free DNA help?
The bacterial genome is relatively simple: one chromosome that the cell is optimized to replicate and maintain. When we use bacteria to produce plasmids, we’re essentially hijacking that machinery and asking it to maintain both its own genome and a foreign plasmid.
Modern biotech requires sequences with features that bacteria are not designed to handle: very large repeat structures, extreme GC content (high or low), complex regulatory elements, engineered promoters and UTRs, and bacteriotoxic elements derived from viruses or regulatory modules. These features are foreign to the bacterial cell, which often does not know how to cope and fails.
For customers, that can mean weeks of trial and error trying to get a synthetic construct to propagate in bacteria, only to discover that the host system is the real bottleneck. Our cell‑free process removes the organismal constraint from the outset, giving us an almost unlimited design “playground” for these complex sequences.
💡 When you move away from plasmid cloning, design is no longer constrained by what E. coli can maintain: inverted repeats, long homopolymers, extreme GC content, and arrays of regulatory elements can be synthesized directly, avoiding the deletions, rearrangements, and “sequence drift” that are common when such constructs are forced through bacterial propagation.
How does this apply to AAV genomes and other viral vectors?
In the AAV workflow, we typically produce the AAV genome, which is then transfected into mammalian cells that package and secrete AAV particles. Traditionally, these AAV genomes are cloned and propagated in bacteria as plasmids before moving into mammalian expression.
However, it is well known that bacteria “dislike” certain AAV elements, especially the inverted terminal repeats (ITRs). These sequences are hard to maintain stably in plasmids. Cell‑free processes offer a way to bypass this bottleneck by directly synthesizing the AAV genome without relying on bacterial propagation.
We see similar benefits for other viral vectors, like lentiviral constructs, where design flexibility and sequence complexity are crucial for next‑generation therapies.
If you work with AAV, you have likely encountered ITR instability during plasmid propagation; recent sequencing anlayses show that a large fraction of deposited ITR‑containing plasmids carry unintended deletions, underscoring why direct synthesis of AAV genomes in cell‑free systems is attractive for maintaining full‑length, sequence‑verified vectors.
What about highly complex genome engineering applications like CRISPR arrays?
In genome engineering, particularly with CRISPR arrays, researchers increasingly want to introduce large cassettes that modify multiple loci in the human genome simultaneously. You combine the inherent complexity of AAV or lentiviral vector backbones with the additional complexity of large CRISPR arrays, and the odds of getting a sequence‑perfect plasmid out of bacteria drop dramatically.
These are exactly the types of applications where we see cell‑free DNA at the forefront. By decoupling from bacterial constraints, we can directly synthesize and assemble these complex constructs, supporting more ambitious genome engineering strategies.
💡 As multiplex CRISPR tools evolve—from long guide RNA arrays to base‑ and prime‑editing cassettes—constructs routinely span multiple kilobases of repetitive, structured sequence, creating a natural fit for cell‑free synthetic DNA production where assembly and maintenance are not limited by bacterial repair and replication systems.
For companies deeply invested in plasmid workflows, what does a realistic hybrid roadmap look like?
We need to be honest: in its current state of maturity, cell‑free DNA is not the superior solution for every single application. The key is to identify where it adds immediate value.
We usually sit down with customers and map their process to pinpoint bottleneck steps—places where plasmid‑based methods cause delays, failures, or regulatory friction. We then propose targeted POC studies in those specific areas. It doesn’t make sense to push cell‑free DNA into trivial plasmid use cases where the benefit is marginal.
For now, the strongest strategy is selective bottleneck removal: use cell‑free DNA where it can clearly accelerate or de‑risk the platform, while keeping plasmids where they are still working well. Over time, that balance can shift as cell‑free technologies and regulatory familiarity advance.
💡 For some organizations, a slow transition to cell-free DNA allows them to test its precision. Here, it is effective when used to alleviate common pains, like slow IVT template generation, or in constructs that are not amenable to cloning, before scaling its use in larger programs.
Looking ahead, will plasmid DNA remain the gold standard, or can cell‑free DNA become the default?
Plasmid DNA will remain an important component of biopharma and biotech workflows; it has been established for 30–40 years, and for some applications there is little added value in switching.
However, I do expect cell‑free DNA to gradually become the default for many emerging applications. We already see this for IVT templates, CRISPR arrays, and increasingly for viral vectors where design flexibility and complexity are essential. Modern therapies demand modern molecule designs, and truly cell‑free synthesis is uniquely suited to deliver those.
Ribbon Bio’s role is to make cell‑free DNA more accessible and dependable, so that teams can unlock these next‑generation designs and accelerate the development of personalized therapies and precision medicine.
Plasmids will continue to be important in biomanufacturing, but we believe cell-free synthetic DNA will become the gold-standard in applications where complexity, speed, and purity are decisive for both scientific success and regulatory acceptance.
Conclusion
The industry is slowly moving away from an exclusive reliance on plasmid cloning in E. coli and toward enzymatic, cell‑free DNA synthesis. By eliminating the bacterial host, cell‑free workflows reduce endotoxin and host‑protein impurities, avoid instability of challenging sequences such as AAV ITRs, and support rapid, small‑batch mRNA and viral‑vector production.
As the field moves toward personalized therapies, cell‑free synthetic DNA appears increasingly central to enabling this next generation of genetic medicine and biomanufacturing.
