Publications

We describe a combinatorial CRISPRinterference (CRISPRi) screening platform for mapping genetic interactions in mammalian cells. We targeted 107 chromatin-regulation factors in human cells with pools of either single or double single guide RNAs (sgRNAs) to downregulate individual genes or gene pairs, respectively. Relative enrichment analysis of individual sgRNAs or sgRNApairs allowed for quantitative characterization of genetic interactions, and comparison with protein–protein-interaction data revealed a functional map of chromatin regulation.

Visualization of chromosome dynamics allows the investigation of spatiotemporal chromatin organization and its role in gene regulation and other cellular processes. However, current approaches to label multiple genomic loci in live cells have a fundamental limitation in the number of loci that can be labeled and uniquely identified. Here we describe an approach we call ‘‘track first and identify later’’ for multiplexed visualization of chromosome dynamics by combining two techniques: CRISPR imaging and DNA sequential fluorescence in situ hybridization. Our approach first labels and tracks chromosomal loci in live cells with the CRISPR-Cas9 system, then barcodes those loci by DNA sequential fluorescence in situ hybridization in fixed cells and resolves their identities. We demonstrate our approach by tracking telomere dynamics, identifying 12 unique subtelomeric regions with variable detection efficiencies, and tracking back the telomere dynamics of respective chromosomes in mouse embryonic stem cells.

We developed a systematic approach to map human genetic networks by combinatorial CRISPR–Cas9 perturbations coupled to robust analysis of growth kinetics. We targeted all pairs of 73 cancer genes with dual guide RNAs in three cell lines, comprising 141,912 tests of interaction. Numerous therapeutically relevant interactions were identified, and these patterns replicated with combinatorial drugs at 75% precision. From these results, we anticipate that cellular context will be critical to synthetic-lethal therapies.

The ability to dynamically manipulate the transcriptome is important for studying how gene networks direct cellular functions and how network perturbations cause disease. Nuclease-dead CRISPR-dCas9 transcriptional regulators, while offering an approach for controlling individual gene expression, remain incapable of dynamically coordinating complex transcriptional events. Here, we describe a flexible dCas9-based platform for chemical-inducible complex gene regulation. From a screen of chemical- and light-inducible dimerization systems, we identified two potent chemical inducers that mediate efficient gene activation and repression in mammalian cells. We combined these inducers with orthogonal dCas9 regulators to independently control expression of different genes within the same cell. Using this platform, we further devised AND, OR, NAND, and NOR dCas9 logic operators and a diametric regulator that activates gene expression with one inducer and represses with another. This work provides a robust CRISPR–dCas9-based platform for enacting complex transcription programs that is suitable for large-scale transcriptome engineering.

Recent advances in genome engineering are starting a revolution in biological research and translational applications. The clustered regularly interspaced short palindromic repeats (CRISPR)-associated RNA-guided endonuclease CRISPR associated protein 9 (Cas9) and its variants enable diverse manipu- lations of genome function. In this review, we describe the development of Cas9 tools for a variety of applications in cell biology research, including the study of functional genomics, the creation of transgenic animal models, and genomic imaging. Novel genome engineering methods offer a new avenue to understand the causality between the genome and phenotype, thus promising a fuller understanding of cell biology. 

Essential gene functions underpin the core reactions required for cell viability, but their contributions and relationships are poorly studied in vivo. Using CRISPR interference, we created knockdowns of every essential gene in Bacillus subtilis  and probed their phenotypes. Our high-confidence essential gene network, established using chemical genomics, showed extensive interconnections among distantly related processes and identified modes of action for uncharacterized antibiotics. Importantly, mild knockdown of essentialgene functions significantly reduced stationary-phase survival without affecting maximal growth rate, suggesting that essential protein levels are set to maximize outgrowth from stationary phase. Finally, high-throughput microscopy indicated that cell morphology is relatively insensitive tomild knockdown but profoundly affected by depletion of gene function, revealing intimate connections between cell growth and shape. Our results provide a framework for systematic investigation of essential gene functions in vivo broadly applicable to diverse microorganisms and amenable to comparative analysis. 

The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated 9 (Cas9) system, a versatile RNA-guided DNA targeting platform, has been revolutionizing our ability to modify, manipulate, and visualize the human genome, which greatly advances both biological research and therapeutics development. Here, we review the current development of CRISPR/Cas9 technologies for gene editing, transcription regulation, genome imaging, and epigenetic modification.We discuss the broad application of this system to the study of functional genomics, especially genome-wide genetic screening, and to therapeutics development, including establishing disease models, correcting defective genetic mutations, and treating diseases.

The Cas9 protein (CRISPR-associated protein 9), derived from type II CRISPR (clustered regularly interspaced short palindromic repeats) bacterial immune systems, is emerging as a powerful tool for engineering the genome in diverse organisms. As an RNA-guided DNA endonuclease, Cas9 can be easily programmed to target new sites by altering its guide RNA sequence, and its development as a tool has made sequence-specific gene editing several magnitudes easier. The nuclease-deactivated form of Cas9 further provides a versatile RNA-guided DNA-targeting platform for regulating and imaging the genome, as well as for rewriting the epigenetic status, all in a sequence-specific manner. With all of these advances, we have just begun to explore the possible applications of Cas9 in biomedical research and therapeutics. In this review, we describe the current models of Cas9 function and the structural and biochemical studies that support it. We focus on the applications of Cas9 for genome editing, regulation, and imaging, discuss other possible applications and some technical considerations, and highlight the many advantages that CRISPR/Cas9 technology offers.

Developing technologies for efficient and scalable disruption of gene expression will provide powerful tools for studying gene function, developmental pathways, and disease mechanisms. Here, we develop clustered regularly interspaced short palindromic repeat interference (CRISPRi) to repress gene expression in human induced pluripotent stem cells (iPSCs).CRISPRi, in which a doxycycline-inducible deactivated Cas9 is fused to a KRAB repression domain, can specifically and reversibly inhibit gene expression in iPSCs and iPSC-derived cardiac progenitors, cardiomyocytes, and T lymphocytes. This gene repression system is tunable and has the potential to silence single alleles. Compared with CRISPR nuclease (CRISPRn), CRISPRi gene repression is more efficient and homogenous across cell populations. The CRISPRi system in iPSCs provides a powerful platform to perform genome-scale screens in a wide range of iPSC-derived cell types, dissect developmental pathways, and model disease.

The human naive pluripotent stem cell (PSC) state, corresponding to a pre-implantation stage of development, has been difficult to capture and sustain in vitro. We report that the Hippo pathway effector YAP is nuclearly localized in the inner cell mass of human blastocysts. Overexpression of YAP in human embryonic stem cells (ESCs) and induced PSCs (iPSCs) promotes the generation of naive PSCs. Lysophosphatidic acid (LPA) can partially substitute for YAP to generate transgene-free human naive PSCs. YAP- or LPA-induced naive PSCs have a rapid clonal growth rate, a normal karyotype, the ability to form teratomas, transcriptional similarities to human pre-implantation embryos, reduced heterochromatin levels, and other hallmarks of the naive state. YAP/LPA act in part by suppressing differentiationinducing effects of GSK3 inhibition. CRISPR/Cas9-generated YAP-/- cells have an impaired ability to form colonies in naive but not primed conditions. These results uncover an unexpected role for YAP in the human naive state, with implications for early human embryology.

The bacterial CRISPR–Cas9 system has emerged as a multifunctional platform for sequence-specific regulation of gene expression. This Review describes the development of technologies based on nuclease-deactivated Cas9, termed dCas9, for RNA-guided genomic transcription regulation, both by repression through CRISPR interference (CRISPRi) and by activation through CRISPR activation (CRISPRa). We highlight different uses in diverse organisms, including bacterial and eukaryotic cells, and summarize current applications of harnessing CRISPR–dCas9 for multiplexed, inducible gene regulation, genome-wide screens and cell fate engineering. We also provide a perspective on future developments of the technology and its applications in biomedical research and clinical studies. 

Targeted modulation of transcription is necessary for understanding complex gene networks and has great potential for medical and industrial applications. Clustered regularly interspaced short palindromic repeats (CRISPR) is emerging as a powerful system for targeted genome activation and repression, in addition to its use in genome editing. This protocol describes how to design, construct, and experimentally validate the function of sequence-specific single guide RNAs (sgRNAs) for sequencespecific repression (CRISPRi) or activation (CRISPRa) of transcription in mammalian cells. In this technology, the CRISPR-associated protein Cas9 is catalytically deactivated (dCas9) to provide a general platform for RNA-guided DNA targeting of any locus in the genome. Fusion of dCas9 to effector domains with distinct regulatory functions enables stable and efficient transcriptional repression or activation in mammalian cells. Delivery of multiple sgRNAs further enables activation or repression of multiple genes. By using scaffold RNAs (scRNAs), different effectors can be recruited to different genes for simultaneous activation of some and repression of others. The CRISPRi and CRISPRa methods provide powerful tools for sequence-specific control of gene expression on a genome-wide scale to aid understanding gene functions and for engineering genetic regulatory systems.

CRISPR-Cas9 technology has rapidly changed the landscape for how biologists and bioengineers study and manipulate the genome.Derived from the bacterial adaptive immune system, CRISPR-Cas9 has been coopted and repurposed for a variety of new functions, including the activation or repression of gene expression (termed CRISPRa or CRISPRi, respectively). This represents an exciting alternative to previously used repression or activation technologies such as RNA interference (RNAi) or the use of gene overexpression vectors. We have only just begun exploring the possibilities that CRISPR technology offers for gene regulation and the control of cell identity and behavior. In this review, we describe the recent advances of CRISPR-Cas9 technology for gene regulation and outline advantages and disadvantages of CRISPRa and CRISPRi (CRISPRa/i) relative to alternative technologies.

In this review we briefly describe the development of CRISPR tools for genome editing and control of transcription in bacteria. We focus on the Type II CRISPR/Cas9 system, provide specific examples for use of the system, and highlight the advantages and disadvantages of CRISPR versus other techniques. We suggest potential strategies for combining CRISPR tools with high-throughput approaches to elucidate gene function in bacteria.

Stem cells occupy variable environments where they must distinguish stochastic fluctuations from developmental cues. Here, we use optogenetics to investigate how the pluripotency network in embryonic stem cells (ESCs) achieves a robust response to differentiation cues but not to gene expression fluctuations. We engineered mouse ESCs to allow quantitative control over the endogenous mechanism of neural differentiation through a light-inducible Brn2 transgene and monitored differentiation status through a genome-integrated Nanog-GFP reporter. By exposing cells to pulses of Brn2, we find that the pluripotency network rejects Brn2 inputs that are below specific magnitude or duration thresholds, but allows rapid differentiation when both thresholds are satisfied. The filtering properties of the network arise through its positive feedback architecture and the intrinsic half-life of Nanog, which determines the duration threshold in the network. Together our results suggest that the dynamic properties of positive feedback networks might determine how inputs are classified as signal or noise by stem cells.

The CRISPR/Cas9 system has recently been developed as a powerful and flexible technology for different purposes of tar-geted genome engineering, including genome editing (modifying the genome sequence) and gene regulation (without modifying the ge-nome sequence). These applications require design of single guide RNAs (sgRNAs) that are efficient and specific, which, however, remains challenging, as it requires consideration of many criteria. Several designer tools have been developed for gene editing, but currently there is no tool to design sgRNAs for gene regulation. With accumulating experimental data on the use of CRISPR/Cas for gene editing and regulation, we implement a comprehensive computa-tional tool based on a set of sgRNA design rules summarized from these published reports. We report a genome-wide sgRNA designer tool and provide an online user-friendly website for predicting sgR-NAs that are efficient and specific. We name the tool CRISPR-ERA, for CRISPR-mediated editing, repression, and activation (ERA). 

CRISPR interference (CRISPRi) is a powerful technology for sequence-specifically repressing gene expression in bacterial cells. CRISPRi requires only a single protein and a custom designed guide RNA for specific gene targeting. In Escherichia coli, CRISPRi repression efficiency is high (~300 fold), and there are no observable off-target effects. The method can be scaled up as a general strategy for the repression of many genes simultaneously using multiple designed guide RNAs. Here we provide a protocol for efficient guide RNA design, cloning and assay of the CRISPRi system in E. coli. In principle, this protocol can be used to construct CRISPRi systems for gene repression in other species of bacteria.

The bacterial CRISPR-Cas9 system has emerged as an effective tool for sequence-specific gene knockout through non-homologous end joining (NHEJ), but it remains inefficient for precise editing of genome sequences. Here we develop a reporter-based screening approach for high-throughput identification of chemical compounds that can modulate precise genome editing through homology-directed repair (HDR). Using our screening method, we have identified small molecules that can enhance CRISPR-mediated HDR efficiency, 3-fold for large fragment insertions and 9-fold for point mutations. Interestingly, we have also observed that a small molecule that inhibits HDR can enhance frame shift insertion and deletion (indel) mutations mediated by NHEJ. The identified small molecules function robustly in diverse cell types with minimal toxicity. The use of small molecules provides a simple and effective strategy to enhance precise genome engineering applications and facilitates the study of DNA repair mechanisms in mammalian cells.

Eukaryotic cells execute complex transcriptional programs in which specific loci throughout the genome are regulated in distinct ways by targeted regulatory assemblies. We have applied this principle to generate synthetic CRISPR-based transcriptional programs in yeast and human cells. By extending guide RNAs to include effector protein recruitment sites, we construct modular scaffold RNAs that encode both target locus and regulatory action. Sets of scaffold RNAs can be used to generate synthetic multigene transcriptional programs in which some genes are activated and others are repressed. We apply this approach to flexibly redirect flux through a complex branched metabolic pathway in yeast. Moreover, these programs can be executed by inducing expression of the dCas9 protein, which acts as a single master regulatory control point. CRISPR-associated RNA scaffolds provide a powerful way to construct synthetic gene expression programs for a wide range of applications, including rewiring cell fates or engineering metabolic pathways.

In microbial communities, bacterial populations are commonly controlled using indiscriminate, broad range antibiotics. There are few ways to target specific strains effectively without disrupting the entire microbiome and local environment. Here, we use conjugation, a natural DNA horizontal transfer process among bacterial species, to deliver an engineered CRISPR interference (CRISPRi) system for targeting specific genes in recipient Escherichia coli cells. We show that delivery of the CRISPRi system is successful and can specifically repress a reporter gene in recipient cells, thereby establishing a new tool for gene regulation across bacterial cells and potentially for bacterial population control.

Signals in many biological processes can be amplified by recruiting multiple copies of regulatory proteins to a site of action. Harnessing this principle, we have developed a protein scaffold, a repeating peptide array termed SunTag, which can recruit multiple copies of an antibody-fusion protein. We show that the SunTag can recruit up to 24 copies of GFP, thereby enabling long-term imaging of single protein molecules in living cells. We also use the SunTag to create a potent synthetic transcription factor by recruiting multiple copies of a transcriptional activation domain to a nuclease-deficient CRISPR/Cas9 protein and demonstrate strong activation of endogenous gene expression and re-engineered cell behavior with this system. Thus, the SunTag provides a versatile platform for multimerizing proteins on a target protein scaffold and is likely to have many applications in imaging and controlling biological outputs.

While the catalog of mammalian transcripts and their expression levels in different cell types and disease states is rapidly expanding, our understanding of transcript function lags behind. We present a robust technology enabling systematic investigation of the cellular consequences of repressing or inducing individual transcripts. We identify rules for specific targeting of transcriptional repressors (CRISPRi), typically achieving 90%–99% knockdown with minimal off-target effects, and activators (CRISPRa) to endogenous genes via endonuclease-deficient Cas9. Together they enable modulation of gene expression over a ∼1,000-fold range. Using these rules, we construct genome-scale CRISPRi and CRISPRa libraries, each of which we validate with two pooled screens. Growth-based screens identify essential genes, tumor suppressors, and regulators of differentiation. Screens for sensitivity to a cholera-diphtheria toxin provide broad insights into the mechanisms of pathogen entry, retrotranslocation and toxicity. Our results establish CRISPRi and CRISPRa as powerful tools that provide rich and complementary information for mapping complex pathways.

Synthetic non-coding RNAs have emerged as a versatile class of molecular devices that have a diverse range of programmable functions, including signal sensing, gene regulation and the modulation of molecular interactions. Owing to their small size and the central role of Watson–Crick base pairing in determining their structure, function and interactions, several distinct types of synthetic non-coding RNA regulators that are functional at the DNA, mRNA and protein levels have been experimentally characterized and computationally modelled. These engineered devices can be incorporated into genetic circuits, enabling the more efficient creation of complex synthetic biological systems. In this Review, we summarize recent progress in engineering synthetic non-coding RNA devices and their application to genetic and cellular engineering in a broad range of microorganisms.

The spatiotemporal organization and dynamics of chromatin play critical roles in regulating genome function. However, visualizing specific, endogenous genomic loci remains challenging in living cells. Here, we demonstrate such an imaging technique by repurposing the bacterial CRISPR/Cas system. Using an EGFP-tagged endonuclease-deficient Cas9 protein and a structurally optimized small guide (sg) RNA, we show robust imaging of repetitive elements in telomeres and coding genes in living cells. Furthermore, an array of sgRNAs tiling along the target locus enables the visualization of nonrepetitive genomic sequences. Using this method, we have studied telomere dynamics during elongation or disruption, the subnuclear localization of the MUC4 loci, the cohesion of replicated MUC4 loci on sister chromatids, and their dynamic behaviors during mitosis. This CRISPR imaging tool has potential to significantly improve the capacity to study the conformation and dynamics of native chromosomes in living human cells.

The genetic interrogation and reprogramming of cells requires methods for robust and precise targeting of genes for expression or repression. The CRISPR-associated catalytically inactive dCas9 protein offers a general platform for RNA-guided DNA targeting. Here, we show that fusion of dCas9 to effector domains with distinct regulatory functions enables stable and efficient transcriptional repression or activation in human and yeast cells, with the site of delivery determined solely by a coexpressed short guide (sg)RNA. Coupling of dCas9 to a transcriptional repressor domain can robustly silence expression of multiple endogenous genes. RNA-seq analysis indicates that CRISPR interference (CRISPRi)-mediated transcriptional repression is highly specific. Our results establish that the CRISPR system can be used as a modular and flexible DNA-binding platform for the recruitment of proteins to a target DNA sequence, revealing the potential of CRISPRi as a general tool for the precise regulation of gene expression in eukaryotic cells.

Sequence-specific control of gene expression on a genome-wide scale is an important approach for understanding gene functions and for engineering genetic regulatory systems. We have recently described an RNA-based method, CRISPR interference (CRISPRi), for targeted silencing of transcription in bacteria and human cells. The CRISPRi system is derived from the Streptococcus pyogenes CRISPR (clustered regularly interspaced palindromic repeats) pathway, requiring only the coexpression of a catalytically inactive Cas9 protein and a customizable single guide RNA (sgRNA). The Cas9-sgRNA complex binds to DNA elements complementary to the sgRNA and causes a steric block that halts transcript elongation by RNA polymerase, resulting in the repression of the target gene. Here we provide a protocol for the design, construction and expression of customized sgRNAs for transcriptional repression of any gene of interest. We also provide details for testing the repression activity of CRISPRi using quantitative fluorescence assays and native elongating transcript sequencing. CRISPRi provides a simplified approach for rapid gene repression within 1–2 weeks. The method can also be adapted for high-throughput interrogation of genome-wide gene functions and genetic interactions, thus providing a complementary approach to RNA interference, which can be used in a wider variety of organisms.

Targeted gene regulation on a genome-wide scale is a powerful strategy for interrogating, perturbing, and engineering cellular systems. Here, we develop a method for controlling gene expression based on Cas9, an RNA-guided DNA endonuclease from a type II CRISPR system. We show that a catalytically dead Cas9 lacking endonuclease activity, when coexpressed with a guide RNA, generates a DNA recognition complex that can specifically interfere with transcriptional elongation, RNA polymerase binding, or transcription factor binding. This system, which we call CRISPR interference (CRISPRi), can efficiently repress expression of targeted genes in Escherichia coli, with no detectable off-target effects. CRISPRi can be used to repress multiple target genes simultaneously, and its effects are reversible. We also show evidence that the system can be adapted for gene repression in mammalian cells. This RNA-guided DNA recognition platform provides a simple approach for selectively perturbing gene expression on a genome-wide scale.

Complex interactions among genetic components often result in variable systemic performance in designed multigene systems. Using the bacterial clustered regularly interspaced short palindromic repeat (CRISPR) pathway we develop a synthetic RNA-processing platform, and show that efficient and specific cleavage of precursor mRNA enables reliable and predictable regulation of multigene operons. Physical separation of linked genetic elements by CRISPR-mediated cleavage is an effective strategy to achieve assembly of promoters, ribosome binding sites, cis-regulatory elements, and riboregulators into single- and multigene operons with predictable functions in bacteria. We also demonstrate that CRISPR-based RNA cleavage is effective for regulation in bacteria, archaea and eukaryotes. Programmable RNA processing using CRISPR offers a general approach for creating context-free genetic elements and can be readily used in the bottom-up construction of increasingly complex biological systems in a plug-and-play manner.

Bacterial regulators of transcriptional elongation are versatile units for building custom genetic switches, as they control the expression of both coding and noncoding RNAs, act on multigene operons and can be predictably tethered into higher-order regulatory functions (a property called composability). Yet the less versatile bacterial regulators of translational initiation are substantially easier to engineer. To bypass this tradeoff, we have developed an adaptor that converts regulators of translational initiation into regulators of transcriptional elongation in Escherichia coli. We applied this adaptor to the construction of several transcriptional attenuators and activators, including a small molecule-triggered attenuator and a group of five mutually orthogonal riboregulators that we assembled into NOR gates of two, three or four RNA inputs. Continued application of our adaptor should produce large collections of transcriptional regulators whose inherent composability can facilitate the predictable engineering of complex synthetic circuits.

Non-coding RNAs (ncRNAs) are versatile regulators in cellular networks. While most trans-acting ncRNAs possess well-defined mechanisms that can regulate transcription or translation, they generally lack the ability to directly sense cellular signals. In this work, we describe a set of design principles for fusing ncRNAs to RNA aptamers to engineer allosteric RNA fusion molecules that modulate the activity of ncRNAs in a ligand-inducible way in Escherichia coli. We apply these principles to ncRNA regulators that can regulate translation (IS10 ncRNA) and transcription (pT181 ncRNA), and demonstrate that our design strategy exhibits high modularity between the aptamer ligand-sensing motif and the ncRNA target-recognition motif, which allows us to reconfigure these two motifs to engineer orthogonally acting fusion molecules that respond to different ligands and regulate different targets in the same cell. Finally, we show that the same ncRNA fused with different sensing domains results in a sensory-level NOR gate that integrates multiple input signals to perform genetic logic. These ligand-sensing ncRNA regulators provide useful tools to modulate the activity of structurally related families of ncRNAs, and building upon the growing body of RNA synthetic biology, our ability to design aptamer-ncRNA fusion molecules offers new ways to engineer ligand-sensing regulatory circuits.

Our ability to routinely engineer genetic networks for applications is limited by the scarcity of highly specific and non-cross-reacting (orthogonal) gene regulators with predictable behavior. Though antisense RNAs are attractive contenders for this purpose, quantitative understanding of their specificity and sequence-function relationship sufficient for their design has been limited. Here, we use rationally designed variants of the RNA-IN-RNA-OUT antisense RNA-mediated translation system from the insertion sequence IS10 to quantify >500 RNA-RNA interactions in Escherichia coli and integrate the data set with sequence-activity modeling to identify the thermodynamic stability of the duplex and the seed region as the key determinants of specificity. Applying this model, we predict the performance of an additional ~2,600 antisense-regulator pairs, forecast the possibility of large families of orthogonal mutants, and forward engineer and experimentally validate two RNA pairs orthogonal to an existing group of five from the training data set. We discuss the potential use of these regulators in next-generation synthetic biology applications.

The widespread natural ability of RNA to sense small molecules and regulate genes has become an important tool for synthetic biology in applications as diverse as environmental sensing and metabolic engineering. Previous work in RNA synthetic biology has engineered RNA mechanisms that independently regulate multiple targets and integrate regulatory signals. However, intracellular regulatory networks built with these systems have required proteins to propagate regulatory signals. In this work, we remove this requirement and expand the RNA synthetic biology toolkit by engineering three unique features of the plasmid pT181 antisense-RNA-mediated transcription attenuation mechanism. First, because the antisense RNA mechanism relies on RNA-RNA interactions, we show how the specificity of the natural system can be engineered to create variants that independently regulate multiple targets in the same cell. Second, because the pT181 mechanism controls transcription, we show how independently acting variants can be configured in tandem to integrate regulatory signals and perform genetic logic. Finally, because both the input and output of the attenuator is RNA, we show how these variants can be configured to directly propagate RNA regulatory signals by constructing an RNA-meditated transcriptional cascade. The combination of these three features within a single RNA-based regulatory mechanism has the potential to simplify the design and construction of genetic networks by directly propagating signals as RNA molecules.

Small-molecule regulation of gene expression is intrinsic to cellular function and indispensable to the construction of new biological sensing, control and expression systems. However, there are currently only a handful of strategies for engineering such regulatory components and fewer still that can give rise to an arbitrarily large set of inducible systems whose members respond to different small molecules, display uniformity and modularity in their mechanisms of regulation, and combine to actuate universal logics. Here we present an approach for small-molecule regulation of transcription based on the combination of cis-regulatory leader-peptide elements with genetically encoded unnatural amino acids (amino acids that have been artificially added to the genetic code). In our system, any genetically encoded unnatural amino acid (UAA) can be used as a small-molecule attenuator or activator of gene transcription, and the logics intrinsic to the network defined by expanded genetic codes can be actuated.

Our current ability to engineer biological circuits is hindered by design cycles that are costly in terms of time and money, with constructs failing to operate as desired, or evolving away from the desired function once deployed. Synthetic biologists seek to understand biological design principles and use them to create technologies that increase the efficiency of the genetic engineering design cycle. Central to the approach is the creation of biological parts — encapsulated functions that can be composited together to create new pathways with predictable behaviors. We define five desirable characteristics of biological parts — independence, reliability, tunability, orthogonality and composability, and review studies of small natural and synthetic biological circuits that provide insights into each of these characteristics. We propose that the creation of appropriate sets of families of parts with these properties is a prerequisite for efficient, predictable engineering of new function in cells and will enable a large increase in the sophistication of genetic engineering applications.