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Reprogramming the genetic code

2011; Springer Nature; Volume: 30; Issue: 12 Linguagem: Inglês

10.1038/emboj.2011.160

1460-2075

Jason W. Chin,

Bacterial Genetics and Biotechnology

Medal Review20 May 2011free access Reprogramming the genetic code Jason W Chin Corresponding Author Jason W Chin Medical Research Council Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author Jason W Chin Corresponding Author Jason W Chin Medical Research Council Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author Author Information Jason W Chin 1 1Medical Research Council Laboratory of Molecular Biology, Cambridge, UK *Corresponding author. Medical Research Council Laboratory of Molecular Biology, Hills Road, Cambridge CB2 0QH, UK. Tel.: +440 122 340 2115; Fax: +440 122 341 2178; E-mail: [email protected] The EMBO Journal (2011)30:2312-2324https://doi.org/10.1038/emboj.2011.160 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Introduction When Maria Leptin called to say that I had been awarded the 2010 EMBO Gold Medal, I was delighted and surprised. The roll call of past winners is impressive, and I had been at the Laboratory of Molecular Biology (LMB) when Jan Löwe had received the award in 2007 for his elegant work on the bacterial cytoskeleton (Michie and Löwe, 2006). I also knew that several other LMB scientists, including Barbara Pearse, Hugh Pelham and Matthew Freeman, had previously won this prestigious award for their seminal contributions to molecular biology (Pelham, 1989; Freeman, 2002). Like many scientists that I know I invariably find the experiments we are about to do the most compelling, and our curiosity and optimism about the future, and the limitless possibilities that it offers, is perhaps part of what drives us continually forward into the unknown, but makes it difficult for us to sit down and write reviews about what we have already done. However, the recognition of our laboratory's work by this medal provides a ‘still point’ in which I can reflect, and allows me the opportunity to provide a personal account that recognizes the contributions to science made by the many talented and generous people that have educated and mentored me, and that I have had the privilege to work with. I hope that this review can honour, in a small way, the debt of gratitude I feel for the ways in which they have enriched my life in science. I was naturally drawn to chemistry at school because it provided a systematic explanation for how all matter behaves in terms of simple sets of rules governed by invisible particles and captured in the periodic table. I had fantastic teaching in both chemistry and physics, and was taught by Mr Liasis, Mr Manthorpe, Mr Teh and Ms Pountney over the years. It was clear to me that I wanted to do chemistry at University and I was lucky enough to be accepted to Oxford. At Oxford I was tutored by Peter Atkins and Gordon Lowe, who appeared to take the view that they were teaching us to be scientists and that the examinations at the end of the degree were incidental distractions that bright people would somehow get through. Their tutorials were aimed at getting us to think, sometimes in unusual ways, and I have not yet forgotten the surreal image of Peter Atkins reclining on his office chaise longue, a glass of dry sherry in hand and his leather trousers creaking, while he shone a lamp on my face and demanded that I explain how I would establish the laws of thermodynamics on a desert island using only a coconut (of course I had a truly marvellous proof, but there is insufficient space for it here!). While tutorials were fantastic entertainment, the lecture courses were generally more prosaic, traditional and thorough. Later in the course I specialized in organic chemistry and was given tutorials with John Sutherland who, like Gordon Lowe, was continually able to relate the chemistry we were learning to biological processes. I decided to do a part II research year with John, whose laboratory then worked on both engineering penicillin biosynthesis to make new antibiotics and the chemical origins of life (Powner et al, 2009). I worked on engineering an enzyme that naturally expands the five-membered ring of penicillin to the six-membered ring in a cephalosporin, so that it would accept new substrates and make new types of cephalosporin antibiotics. This experience got me hooked on a combination of chemistry and molecular biology and the idea of doing more research. In 1996, I moved to Yale to study for a PhD, since a US PhD allowed me to complement a thorough training in chemistry I had received at Oxford with the opportunity to take biology classes. After the first year at Yale, I had taken most of the biology courses and felt equally comfortable with both chemistry and biology. I decided to work for Alanna Schepartz at Yale, who had a great project that had been pioneered by a graduate student in the laboratory, Neal Zondlo. Neal had shown that it was possible to dissect out the DNA-binding residues of a helical protein and transfer these residues in register onto a small stable scaffold protein to generate a new functional chimeric protein with exquisite DNA-binding affinity and specificity (Zondlo and Schepartz, 1999). With Robert Grotzfeld, I developed combinatorial approaches, using phage display, to extend the approach Neal had developed. I went on to show that we could make high-affinity binders for protein and DNA targets using this approach (Chin et al, 2001; Chin and Schepartz, 2001a, 2001b). Since the approaches we developed were new to the laboratory, I learned a lot about how to do experiments and how to get things to work from scratch, which has been very valuable. In 2001, I finished my PhD and went off to Scripps for a postdoc with Pete Schultz. From then on my research has focussed on engineering the translational machinery of cells for incorporating new amino acids. I, therefore, describe our current overall strategy for reprogramming translation first (below), and then go on to describe how my postdoctoral work fits in and provides one key part of the foundation for our current programme. Reprogramming translation Protein translation is the process by which cells decode genetic information to build functional polymers of amino acids. While natural protein translation synthesizes proteins composed of the natural 20 amino acids, the process by which these polymers are made provides the ultimate paradigm for the synthesis of proteins containing unnatural amino acids beyond the canonical 20, and for the synthesis of entirely unnatural evolvable polymers of genetically determined length, composition and sequence (Figure 1). Figure 1.Reprogramming the genetic code. (A) A central paradigm in molecular biology for the synthesis of proteins containing natural amino acids (coloured circles) may be engineered for the synthesis of proteins containing unnatural amino acids (coloured stars), and by extension of the synthesis of completely unnatural polymers. (B) Progress in reprogramming the genetic code. Natural amino acids are represented by coloured circles and unnatural amino acids by coloured stars. The vertical axis shows progress in incorporating unnatural amino acids into proteins, while the horizontal axis shows progress in incorporating unnatural amino acids in increasingly complex organisms. Red arrows represent steps that have been experimentally demonstrated. Download figure Download PowerPoint Over the last 7 years, we have engineered protein translation for several purposes. First, we have engineered protein translation to create foundational approaches for the encoded and evolvable synthesis of new polymers (Rackham and Chin, 2005a; Wang et al, 2007; Neumann et al, 2010b) (Figure 2). Second, we have developed methods for site specifically installing several key post-translational modifications into recombinant proteins, and used these methods to provide previously unattainable insight into the role of these modifications in regulating biological function (Neumann et al, 2008a, 2008b, 2009; Nguyen et al, 2009a, 2009b; Lammers et al, 2010; Virdee et al, 2010; Zhao et al, 2010; Akutsu et al, 2011; Arbely et al, 2011) (Figure 3). Third, we have developed ‘photochemical genetic’ methods to rapidly control the activity of proteins in living cells, providing insight into the dynamics of elementary steps in biological processes, as well as insight into the regulation of intracellular network connectivity in space and time (Figure 4) (Gautier et al, 2010, 2011); with these methods, we hope to understand molecular processes inside cells and organisms with the level of precision more commonly associated with in vitro biochemistry or biophysics. Figure 2.Engineering the translational machinery to reprogramme the genetic code. (A) Creating orthogonal amber suppressor aminoacyl-tRNA synthetase/tRNA pairs. Cells contain natural synthetases (grey) that use natural amino acids (black oval) to aminoacylate natural tRNAs (black trident). Expanding the genetic code requires the addition of an orthogonal synthetase, tRNA, and amino acid (star) shown in blue. (B) Creation of orthogonal ribosome mRNA pairs by duplication and specialization. The natural cellular ribosome (grey) recognizes natural messages, while the new orthogonal ribosome (green) recognizes orthogonal messages (purple). (C) Evolving the orthogonal ribosome for quadruplet decoding for a parallel genetic code. Download figure Download PowerPoint Figure 3.Genetically encoding lysine acetylation, methylation and ubiquitination. (A) The chemical structures of the post-translationally modified amino acids. (B) Omit map of acetyl lysine density in acetylated cyclophilin A, contoured at 1σ. (C) Structural comparison of the acetylated and unacetylated cyclophilin A–cyclosporine complexes. Cyclosporine is in yellow. The backbone structure of cyclophilin and acetylated cyclophilin are very similar. The backbone of cyclophilin A from a previous structure (PBB 2CPL) is shown in grey and the acetyllysine from our structure is in green. Waters belonging to the unacetylated complex are in blue, waters belonging to the acetylated complex are in green. Acetylation leads to a reorganization of the water network at the interface. This rationalizes why acetylating cyclophilin leads to a 20-fold lower affinity for cyclosporine that may antagonize the immunosuppressive effects of cyclosporine. Download figure Download PowerPoint Figure 4.Controlling specific molecular events inside living cells with light. (A) Caging a near-universally conserved lysine (K97) in the MEK1 active site inactivates the enzyme by sterically blocking ATP binding. Decaging with light rapidly removes the caging group and activates the kinase (figures created using Pymol and MEK1 structure PDB: 1S9J). (B) Isolating a synthetic photoactivateable subnetwork in MAP kinase signalling, via genetically encoding a photocaged lysine in the MEK1 active site. Download figure Download PowerPoint The fidelity of natural translation is primarily set by two processes: (1) aminoacylation of the correct tRNA, and no other tRNA by an aminoacyl-tRNA synthetase (Ling et al, 2009) and (2) the correct decoding and translocation of each tRNA by the ribosome in response to its cognate triplet codon on the mRNA to direct peptide bond formation (Ramakrishnan, 2002). As the ribosome uses tRNA adapter molecules, the chemical identity of the monomers polymerized is chemically independent of the template; this is distinct from the case in nucleic acid-dependent nucleic acid polymerases (DNA polymerases and RNA polymerases), where the substrates for polymerization must directly pair with the template. As the ribosome uses a single set of active sites for polymerization, coupled to a translocation activity, it is—unlike NRPSs, PKSs or fatty acid synthetases—able to synthesize very long polymers of defined and arbitrarily programmed sequence. We realized that in order to reprogramme protein translation to incorporate new amino acids into proteins, and ultimately to synthesize completely unnatural polymers, there are at least three challenges we need to address. First, we need to uniquely attach a new amino acid to a new tRNA. This requires the creation of orthogonal aminoacyl-tRNA synthetase/tRNA pairs in which the aminoacyl-tRNA synthetase is able to uniquely recognize a new amino acid that is not a substrate for endogenous synthetases in the host organism and specifically load the new amino acid onto a cognate tRNA that is not a substrate for endogenous synthetases. Next, we need a codon with which we can uniquely encode the incorporation of the new amino acid. Each of the 64 triplet codons is used in encoding the synthesis of natural proteins, but we have demonstrated that it is possible to evolve the ribosome itself to decode additional genetic information. Finally, the chemical scope of natural protein translation is limited to the synthesis of polypeptides from α-L amino acids; to synthesize a full range of new polymers will require alteration of the ribosome's peptidyl-transferase centre (Dedkova et al, 2003) as well as, potentially, alterations to other parts of the translational machinery. Scripps, La Jolla and orthogonal aminoacyl-tRNA synthetase/tRNA pairs While there are no blank codons in the genetic code, it is well known that the amber stop codon can be decoded, using amber suppressor tRNAs, in a variety of cells and organisms. Amber suppression is inherently inefficient because the amber stop codon is normally read as a termination signal by protein release factors that bind to the A site in the ribosome and hydrolyse the nascent polypeptide chain attached to the P-site tRNA (Capecchi, 1967; Scolnick et al, 1968; Petry et al, 2005). When both an amber suppressor tRNA and the release factor are present in cells, the tRNA and release factor compete for A-site binding. Under these conditions, 80% of protein synthesis that is initiated typically terminates in response to the amber codon, while 20% is decoded by the amber suppressor tRNA and continues to produce the full-length protein (Wang et al, 2007). Though amber suppression is inefficient, it provides a codon that we can use as an initial insertion signal for unnatural amino-acid incorporation. This allows us to focus our attention on the first problem in reprogramming the genetic code—discovering aminoacyl-tRNA synthetase/tRNACUA pairs that are orthogonal in the host and that direct the incorporation of new amino acids. In fact, my interest in incorporating unnatural amino acids into proteins preceded any notion of wholesale genetic code reprogramming, and in 2001, I went to Pete Schultz's laboratory at Scripps to work on creating amber suppressor synthetase/tRNA pairs for incorporating unnatural amino acids into proteins in cells. Pete's laboratory had a long-term interest in incorporating unnatural amino acids into proteins. Indeed, when I applied to work with Pete, his laboratory had already pioneered in vitro methods for incorporating unnatural amino acids into proteins (Noren et al, 1989; Mendel et al, 1995) by combining cell extracts with methods for the chemical aminoacylation of amber suppressor tRNAs (Hecht et al, 1978; Heckler et al, 1984). With Dennis Dougherty and Henry Lester at Caltech, they had extended these approaches, using microinjection of aminoacylated tRNAs, into Xenopus oocytes (Dougherty, 2008). This allowed the introduction of unnatural amino acids into channel proteins expressed in the oocyte. The Dougherty laboratory extended these unnatural amino-acid mutagenesis strategies and has performed elegant studies that probed and defined the dynamics of nicotinic receptor and the role of pi-cation interactions in protein interactions (Dougherty, 2008). Early experiments from the Schultz and Dougherty laboratories beautifully demonstrated how the ability to tailor the properties of individual amino acids atom by atom at defined sites in proteins allow new biological insights to be revealed in complicated systems, via the application of physical organic chemistry principles. However, because in vitro methods for aminoacylating tRNAs are inherently inefficient and do not allow re-acylation of the tRNA in the translation reaction, the in vitro aminoacylation and translation methods yielded small amounts of protein and were technically very challenging. The Schultz laboratory was, therefore, working hard on developing in vivo methods for incorporating unnatural amino acids by engineering aminoacyl-tRNA synthetases and tRNAs (Figure 2A). Early indications that it might be possible to site specifically add new amino acids to proteins produced in cells came from experiments reported by Furter (1998). These experiments demonstrated that a fluorinated analogue of phenylalanine could be incorporated into a protein in Escherichia coli in response to the amber codon using the yeast phenylalanyl-tRNA synthetase tRNACUA pair. Since it is known that fluorinated analogues of phenylalanine are substrates for phenylalanine synthetases, these experiments used a strain of E. coli normally resistant to fluorinated phenylalanine, to avoid incorporation of fluorinated phenylalanine at sense codons via the endogenous E. coli PheRS/tRNAs. Since the yeast synthetase recognizes both phenylalanine and the fluorinated phenylalanine added to the cells, a mixture of fluorinated phenylalanine and natural amino acids were incorporated into the protein in response to the amber codon. David Liu, Thomas Magliery, Miro Pasternak and Peter Schultz articulated that the discovery of aminoacyl-tRNA synthetase/tRNACUA pairs that are orthogonal in a host organism, and that direct the site specific and quantitative incorporation of new amino acids might be achieved by breaking the problem down into two sub-problems (Liu et al, 1997; Liu and Schultz, 1999): (1) discovering aminoacyl-tRNA synthetase/tRNACUA, where the synthetase uses a natural amino acid but does not aminoacylate any tRNAs in the host organism, and the tRNACUA is not a substrate for any endogenous synthetases and (2) reprogramming the synthetase enzyme so that it uniquely recognizes a new unnatural amino acid added to the cell and no natural amino acids. The first sub-problem was addressed by importing aminoacyl-tRNA synthetase/tRNA pairs from heterologous organisms, taking advantage of the evolutionary divergence of synthetase and tRNA sequence and structure between domains of life. The second sub-problem was addressed by creating large libraries (109 variants) of aminoacyl-tRNA synthetase mutants in which the mutations are targeted, using structural information, and performing a two-step genetic selection on this library to identify synthetases that specifically use an unnatural amino acid and no natural amino acids. In Pete's laboratory, I addressed the incorporation of several of the first unnatural amino acids into proteins in response to the amber codon in E. coli using this strategy (Chin et al, 2002a, 2002b). This work took advantage of an amber suppressor derivative of the Methanococcus janaschii tyrosyl-tRNA synthetase (MjTyrRS)/tRNA pair, which is orthogonal in E. coli (Xie and Schultz, 2006). We showed that this pair could be evolved to direct the incorporation of a range of unnatural amino acids with useful properties in response to the amber codon. In particular, I demonstrated that it was possible to evolve this pair to incorporate photocrosslinking amino acids into proteins in response to the amber codon in E. coli (Chin et al, 2002a, 2002b; Chin and Schultz, 2002). This allowed the sites of protein interactions to be mapped both in vitro and in vivo by simply shining light on cells. Unlike non-covalent methods of investigating protein interactions in vivo, such as TAP tagging, this method traps the protein interaction in the cell before purification, and gives direct information about the sites within the proteins that are involved in interactions. The methods we developed have been used to obtain direct information about protein interactions in environments that are difficult to probe by other methods, for example for proteins at or in membranes. In addition, the method may be used to trap some of the most interesting weak or transient interactions that may be systematically lost in non-covalent approaches. Numerous laboratories have used the crosslinking methods we developed to provide unique insights into protein interactions in diverse systems, including the interactions of chaperones (trigger factor, ClpB and GroEL) with substrates, protein interactions important in cell-cycle regulation, conformational changes in RNAP and the topology of transcriptional initiation complexes, protein interactions at the inner and outer membrane of E. coli, protein interactions in the mitochondrial and ER membranes in yeast and protein interactions at the plasma membrane in mammalian cells (Schlieker et al, 2004; Weibezahn et al, 2004; Farrell et al, 2005; Kaiser et al, 2006; Mori and Ito, 2006; Chen et al, 2007; Haslberger et al, 2007; Lakshmipathy et al, 2007; Boos et al, 2008; Kimata et al, 2008; Mohibullah and Hahn, 2008; Panahandeh et al, 2008; Braig et al, 2009; Ieva and Bernstein, 2009; Okuda and Tokuda, 2009; Raschle et al, 2009; Tamura et al, 2009; Carvalho et al, 2010; Jensen et al, 2010; Liu et al, 2010; Tagami et al, 2010; Yamano et al, 2010). The initial methods for incorporating unnatural amino acids into proteins could only be applied in E. coli. I was interested in incorporating unnatural amino acids into eukaryotic cells and organisms because of the enormous potential I saw in being able to make atomic perturbations at specific sites in a specific protein within complex organisms. I realized that such approaches might allow us to dissect, follow and manipulate complex biological processes in space and time directly in vivo. However, the MjTyrRS/tRNACUA pair that we had used in E. coli could not be used in eukaryotic cells because it is not orthogonal with respect to eukaryotic synthetases and tRNAs. It was clear that to expand the genetic code of eukaryotic cells, we would need (1) new synthetase tRNA pairs and (2) new methods to evolve the specificity of these pairs directly in a eukaryotic host. Schimmel's laboratory and others had shown that tyrosyl-tRNA synthetase/tRNACUA pair and the leucyl-tRNA synthetase/tRNACUA pair may be orthogonal in eukaryotes (Edwards and Schimmel, 1990), and so I created a strategy to evolve these pairs to incorporate unnatural amino acids into proteins in yeast. I not only saw yeast as both interesting in its own right for genetic code expansion but also realized that the synthetases we evolved in this system might be directly transplanted to other eukaryotic hosts, including mammalian cells, where the direct transformation with large libraries of synthetase genes and the rapid selections and deconvolution methods we developed in yeast would not have been possible. Eric Meggers, Chris Anderson and Ashton Cropp worked with me on this project, and we successfully developed a method for evolving aminoacyl-tRNA synthetase/tRNA pairs for the incorporating unnatural amino acids, including photocrosslinkers, heavy atoms, biophysical probes and bio-orthogonal labels for protein labelling, into proteins in eukaryotic cells for the first time (Chin et al, 2003a, 2003b). The synthetases we developed in this work are widely used to probe processes in yeast and mammalian cells (Hino et al, 2005, 2011; Chen et al, 2007; Huang et al, 2008; Mohibullah and Hahn, 2008; Ye et al, 2009, 2010; Carvalho et al, 2010). Cambridge, reprogramming translation The Cambridge laboratory began in the summer of 2003. In 2002, shortly after finishing my PhD and moving to Scripps, I had contacted Greg Winter, whose pioneering work on protein engineering I knew well. Indeed, Greg's seminal work on antibody engineering (Jones et al, 1986; Riechmann et al, 1988; Winter and Milstein, 1991) had been an inspiration for my PhD work and his work, along with Alan Fersht and others, on defining the functional centres of tyrosyl-tRNA synthetase through early site-directed mutagenesis experiments had formed a foundation for my postdoctoral work on engineering these enzymes (Winter et al, 1982; Fersht et al, 1985; Bedouelle and Winter, 1986). Greg invited me to visit the Medical Research Council LMB and this eventually led to the offer of an independent position at LMB, with the suggestion that I go away and think of something ambitious and important to do in my independent career and the promise that I would have reasonable resources to get started. I accepted with the proviso that I would stay a year and a half to finish my postdoctoral projects at Scripps. I am very fortunate to be part of a community and environment at LMB, where there are few barriers to doing science. From my postdoctoral work with Pete, I was convinced of the enormous potential of encoding unnatural amino acids into proteins, but felt that we had only begun to scratch the surface of what might be possible. When I began to think about how we might systematically reprogramme translation in cells, I realized that we needed to take control of the engine of translation—the ribosome—and make a version of the ribosome that we could alter or evolve to do what we wanted. The ribosome is large and complicated. But exciting progress in structural biology of the ribosome had begun to provide a detailed picture of the subunits and the functional centres of the ribosome. An electrifying talk by Venki Ramakrishnan on 9 May 2003 at the Skirball Institute at NYU convinced me that we were now entering an era in which the ribosome could be understood in molecular detail and—potentially—engineered. Indeed, the molecular insights that Venki and his group at LMB have provided, along with many insights provided by the rest of the ribosome field, have turned out to be invaluable to our work on engineering the ribosome. However, I realized that even if we understood in molecular detail how to engineer the ribosome, altering the cellular ribosome—which is the ultimate cellular hub and responsible for making every protein in the cell—would be problematic. Indeed, it is well known that many mutations in the ribosome are dominant negative or lethal, since they interfere with the synthesis of the entire proteome. I realized that if we could create a new ‘orthogonal’ ribosome that was uncoupled from the requirement to synthesize the proteome, and decoded a message that was not read by the endogenous ribosome, then this new ribosome—which would be non-essential to the cell—should, in principle, be evolvable in the laboratory. Moreover, since the genetic code is a correspondence between amino acids and codons, set by the translational machinery, I realized that the selective delivery of tRNAs aminoacylated with unnatural amino acids to the orthogonal ribosome could form the basis for a parallel and independent, or orthogonal, genetic code for the synthesis of unnatural polymers. Oliver Rackham, who was the first postdoc in the Cambridge laboratory, began work on creating the orthogonal ribosome in E. coli. He first developed a genetic selection through which we could select for or against the expression of a single gene fusion and then showed that he could use this to select mRNA leader sequences, containing alternative Shine Dalgarno sequences (Hui and de Boer, 1987; Rackham and Chin, 2005a), that were not recognized by the endogenous ribosome, but are specifically and efficiently read by a new orthogonal ribosome (Rackham and Chin, 2005a) (Figure 2B). Oliver Rackham began to take advantage of this new non-essential orthogonal ribosome and showed that it is possible to use different orthogonal ribosomes to produce Boolean logic in gene expression (Rackham and Chin, 2005b). More recently, Wenlin An has shown that it is possible to select genetic elements that direct orthogonal transcription by T7 RNAP and orthogonal translation by an orthogonal ribosome (An and Chin, 2009). This provides an orthogonal gene expression pathway in the cell that is entirely insulated from that of normal gene expression. We have suggested that the synthesis of orthogonal, parallel and independent systems, that are released from the constraints that are frozen in natural biology by the evolutionary process, will allow the synthetic evolution of the most fundamental systems in biology. Furthermore, the selective insulation of orthogonal systems from cellular regulation may provide foundational technologies for making biology more amenable to engineering. Orthogonal systems may, therefore, provide a key to the creation of scalable, complex dynamic synthetic biology systems constructed from a large number of biological parts (Kwok, 2010). Wenlin demonstrated that the orthogonal gene expression pathway can be used to set up regulatory circuits that cannot be created using the endogenous, essential transcription and translation machinery (An and Chin, 2009). For example, Wenlin showed that it is possible to create a variety of transcription–translation networks, including transcription–translation feed forward loops, which would be impossible to create using the endogenous machinery. This allowed Wenlin to control the timing of gene expression in new ways and introduce information processing delays into gene expression on the order of hours. In the process of this work, Wenlin was also able to define a minimal transcript that is correctly transcribed and processed to produce a functional 16S rRNA in the ribosome small subunit. This allowed Wenlin to provide insights—into the minimal requirements for rRNA processing—that