Synthesizing proteins inside liposomes and other micro-compartments is today a well-established practice. However, the origin of this research is not distant in time, dating back to the 1999-2004 period, where the first successful attempts were published. Protein synthesis inside artificial compartments is now under strong expansion in the context of synthetic biology (in the “bottom up” approaches), and in particular it strongly contributes to the construction of artificial cell-like systems. These systems, often called “synthetic cells”, can be used to model cellular processes, including membrane-centred ones. They are very innovative models that complement traditional studies and promise future applications. This review does not discuss all current directions in synthetic cell research; in particular it does not include all kind of artificial compartments. Instead, it is uniquely dedicated to the analysis of historical and technical developments of protein synthesis inside liposomes, highlighting a selected list of open questions. One of the goals is remarking the importance of mastering liposome technology together to cell-free systems for the successful realization of this specific type of synthetic cells. At this aim, four currently employed protocols are compared and commented, with major emphasis on the droplet transfer method, which is attractive due to its simplicity and encapsulation efficiency.

Cell-free transcription-translation (TXTL) has recently emerged as a versatile technology to engineer biological systems. In this chapter, we show how an all E. coli TXTL system can be used to build synthetic cell prototypes. We describe methods to encapsulate TXTL reactions in cell-sized liposomes, with an emphasis on the composition of the external solution and lipid bilayer. Cell-free expression is quantitatively described in bulk reactions and liposomes for three proteins: the soluble reporter protein eGFP, the membrane proteins alpha-hemolysin (AH) from Staphylococcus aureus, and the mechanosensitive channel of large conductance (MscL) from E. coli.

As expressed “God made the bulk; the surface was invented by the devil” by W. Pauli, the surface has remarkable properties because broken symmetry in surface alters the material properties. In biological systems, the smallest functional and structural unit, which has a functional bulk space enclosed by a thin interface, is a cell. Cells contain inner cytosolic soup in which genetic information stored in DNA can be expressed through transcription (TX) and translation (TL). The exploration of cell-sized confinement has been recently investigated by using micron-scale droplets and microfluidic devices. In the first part of this review article, we describe recent developments of cell-free bioreactors where bacterial TX-TL machinery and DNA are encapsulated in these cell-sized compartments. Since synthetic biology and microfluidics meet toward the bottom-up assembly of cell-free bioreactors, the interplay between cellular geometry and TX-TL advances better control of biological structure and dynamics in vitro system. Furthermore, biological systems that show self-organization in confined space are not limited to a single cell, but are also involved in the collective behavior of motile cells, named active matter. In the second part, we describe recent studies where collectively ordered patterns of active matter, from bacterial suspensions to active cytoskeleton, are self-organized. Since geometry and topology are vital concepts to understand the ordered phase of active matter, a microfluidic device with designed compartments allows one to explore geometric principles behind self-organization across the molecular scale to cellular scale. Finally, we discuss the future perspectives of a microfluidic approach to explore the further understanding of biological systems from geometric and topological aspects.

Cell-free transcription-translation provides a simplified prototyping environment to rapidly design and study synthetic networks. Despite the presence of a well characterised toolbox of genetic elements, examples of genetic networks that exhibit complex temporal behaviour are scarce. Here, we present a genetic oscillator implemented in an E.coli based cell-free system under steady-state conditions using microfluidic flow reactors. The oscillator has an activator-repressor motif which utilizes the native transcriptional machinery of E.coli; the RNAP and its associated sigma factors. We optimized a kinetic model with experimental data using an evolutionary algorithm to quantify the key regulatory model parameters. The functional modulation of the RNAP was investigated by coupling two oscillators driven by competing sigma factors, allowing the modification of network properties by means of passive transcriptional regulation.

The fast growing bacterium Vibrio natriegens is an emerging microbial host for biotechnology. Harnessing its productive cellular components may offer a compelling platform for rapid protein production and prototyping of metabolic pathways or genetic circuits. Here, we report the development of a V. natriegens cell-free expression system. We devised a simplified crude extract preparation protocol and achieved >260 μg/mL of superfolder GFP in a small-scale batch reaction after 3 h. Culturing conditions, including growth media and cell density, significantly affect translation kinetics and protein yield of extracts. We observed maximal protein yield at incubation temperatures of 26 or 30 °C, and show improved yield by tuning ions crucial for ribosomal stability. This work establishes an initial V. natriegens cell-free expression system, enables probing of V. natriegens biology, and will serve as a platform to accelerate metabolic engineering and synthetic biology applications.

The Class 2 Type V-A CRISPR effector protein Cas12a/Cpf1 has gained widespread attention in part because of the ease in achieving multiplexed genome editing, gene regulation, and DNA detection. Multiplexing derives from the ability of Cas12a alone to generate multiple guide RNAs from a transcribed CRISPR array encoding alternating conserved repeats and targeting spacers. While array design has focused on how to optimize guide-RNA sequences, little attention has been paid to sequences outside of the CRISPR array. Here, we show that a structured hairpin located immediately downstream of the 3ʹ repeat interferes with utilization of the adjacent encoded guide RNA by Francisella novicida (Fn)Cas12a. We first observed that a synthetic Rho-independent terminator immediately downstream of an array impaired DNA cleavage based on plasmid clearance in E. coli and DNA cleavage in a cell-free transcription-translation (TXTL) system. TXTL-based cleavage assays further revealed that inhibition was associated with incomplete processing of the transcribed CRISPR array and could be attributed to the stable hairpin formed by the terminator. We also found that the inhibitory effect partially extended to upstream spacers in a multi-spacer array. Finally, we found that removing the terminal repeat from the array increased the inhibitory effect, while replacing this repeat with an unprocessable terminal repeat from a native FnCas12a array restored cleavage activity directed by the adjacent encoded guide RNA. Our study thus revealed that sequences surrounding a CRISPR array can interfere with the function of a CRISPR nuclease, with implications for the design and evolution of CRISPR arrays.

Cas12a (Cpf1) is a CRISPR-associated nuclease with broad utility for synthetic genome engineering, agricultural genomics, and biomedical applications. While bacteria harboring CRISPR-Cas9 or CRISPR-Cas3 adaptive immune systems sometimes acquire mobile genetic elements encoding anti-CRISPR proteins that inhibit Cas9, Cas3, or the DNA-binding Cascade complex, no such inhibitors have been found for CRISPR-Cas12a. Here we employ a comprehensive bioinformatic and experimental screening approach to identify three different inhibitors that block or diminish CRISPR-Cas12a-mediated genome editing in human cells. We also find a widespread connection between CRISPR self-targeting and inhibitor prevalence in prokaryotic genomes, suggesting a straightforward path to the discovery of many more anti-CRISPRs from the microbial world.

Chromatin is a system of nuclear proteins and nucleic acids that plays a pivotal role in gene expression and cell behavior and is therefore the subject of intense study for cell development and cancer research. Biochemistry, crystallography, and reverse genetics have elucidated the macromolecular interactions that drive chromatin regulation. One of the central mechanisms is the recognition of post-translational modifications (PTMs) on histone proteins by a family of nuclear proteins known as “readers”. This knowledge has launched a wave of activity around the rational design of proteins that interact with histone PTMs. Useful molecular tools have emerged from this work, enabling researchers to probe and manipulate chromatin states in live cells. Chromatin-based proteins represent a vast design space that remains underexplored. Therefore, we have developed a rapid prototyping platform to identify engineered fusion proteins that bind histone PTMs in vitro and regulate genes near the same histone PTMs in living cells. We have used our system to build gene activators with strong avidity for the gene silencing-associated histone PTM H3K27me3. Here, we describe procedures and data for cell-free production of fluorescently tagged fusion proteins, enzyme-linked immunosorbent assay-based measurement of histone PTM binding, and a live cell assay to demonstrate that the fusion proteins modulate transcriptional activation at a site that carries the target histone PTM. This pipeline will be useful for synthetic biologists who are interested in designing novel histone PTM-binding actuators and probes.

CRISPR-Cas systems inherently multiplex through their CRISPR arrays–whether to confer immunity against multiple invaders or by mediating multi-target editing, regulation, imaging, and sensing. However, arrays remain difficult to generate due to their reoccurring repeat sequences. Here, we report an efficient, one-step scheme called CRATES to construct large CRISPR arrays through defined assembly junctions within the trimmed portion of array spacers. We show that the constructed arrays function with the single-effector nucleases Cas9, Cas12a, and Cas13a for multiplexed DNA/RNA cleavage and gene regulation in cell-free systems, bacteria, and yeast. We also applied CRATES to assemble composite arrays utilized by multiple Cas nucleases, where these arrays enhanced DNA targeting specificity by blocking off-target sites. Finally, array characterization revealed context-dependent loss of spacer activity and processing of unintended guide RNAs derived from Cas12a terminal repeats. CRATES thus can facilitate diverse applications requiring CRISPR multiplexing and help elucidate critical factors influencing array function.

The RNA-guided nucleases derived from the CRISPR-Cas systems in bacteria and archaea have found numerous applications in biotechnology, including genome editing, imaging, and gene regulation. However, the discovery of novel Cas nucleases has outpaced their characterization and subsequent exploitation. A key step in characterizing Cas nucleases is determining which protospacer-adjacent motif (PAM) sequences they recognize. Here, we report advances to an in vitro method based on an E. coli cell-free transcription-translation system (TXTL) to rapidly elucidate PAMs recognized by Cas nucleases. The method obviates the need for cloning Cas nucleases or gRNAs, does not require the purification of protein or RNA, and can be performed in less than a day. To advance our previously published method, we incorporated an internal GFP cleavage control to assess the extent of library cleavage as well as Sanger sequencing of the cleaved library to assess PAM depletion prior to next-generation sequencing. We also detail the methods needed to construct all relevant DNA constructs, and how to troubleshoot the assay. We finally demonstrate the technique by determining PAM sequences recognized by the Neisseria meningitidis Cas9, revealing subtle sequence requirements of this highly specific PAM. The overall method offers a rapid means to identify PAMs recognized by diverse CRISPR nucleases, with the potential to greatly accelerate our ability to characterize and harness novel CRISPR nucleases across their many uses.