Alicja Brozek
I am a first year PhD student from Imperial College London modelling the gene network underlying stem cell differentiation and robustness in C. elegans. When I’m not in the lab, I enjoy horseback riding, climbing and dancing.
I am a first year PhD student from Imperial College London modelling the gene network underlying stem cell differentiation and robustness in C. elegans. When I’m not in the lab, I enjoy horseback riding, climbing and dancing.
Please join us at a virtual workshop to assess the Data Science: Data Integration, Modeling, and Automation technical theme of Engineering Biology.
Friday, June 11, 2021
12:00pm – 3:00pm Eastern | 9:00am – 12:00pm Pacific
AgendaRegister Here
This workshop will draw upon our survey results to assess technical progress in the context of the research milestones predicted in Engineering Biology, the 2019 EBRC technical research roadmap. This information will shape a narrative of the most pressing needs and challenges still facing engineering biology over the next decade.
The workshop will inventory the degree of completion for each milestone, discuss technical and nontechnical barriers inhibiting progress, highlight new directions and avenues of research, and review social science dimensions associated with the development of technical goals. This will come in the form of a published report that will be adapted for public viewing.
These virtual writing workshops for each technical theme (3 hours each) are focused on drafting and revising the assessment. They are organized as followed:
The report will be invaluable to policymakers and funders to understand the continued challenges faced by our researchers; to researchers to learn the critical research gaps preventing engineering biology progression; and for our EBRC community to assess the utility of our roadmaps.
Hosted by Adam Arkin (UC Berkeley) and Nathan Hillson (LBNL)
Please join us at a virtual workshop to assess the Host Engineering: Host and Consortia Engineering technical theme of Engineering Biology.
Thursday, June 10, 2021
11:00am – 2:00pm Eastern | 8:00am – 11:00am Pacific
AgendaRegister Here
This workshop will draw upon our survey results to assess technical progress in the context of the research milestones predicted in Engineering Biology, the 2019 EBRC technical research roadmap. This information will shape a narrative of the most pressing needs and challenges still facing engineering biology over the next decade.
The workshop will inventory the degree of completion for each milestone, discuss technical and nontechnical barriers inhibiting progress, highlight new directions and avenues of research, and review social science dimensions associated with the development of technical goals. This will come in the form of a published report that will be adapted for public viewing.
These virtual writing workshops for each technical theme (3 hours each) are focused on drafting and revising the assessment. They are organized as followed:
The report will be invaluable to policymakers and funders to understand the continued challenges faced by our researchers; to researchers to learn the critical research gaps preventing engineering biology progression; and for our EBRC community to assess the utility of our roadmaps.
Hosted by James Carothers (UW) and Ute Galm (Zymergen)
Please join us at a virtual workshop to assess the Engineering DNA: Gene Editing, Synthesis, and Assembly technical theme of Engineering Biology.
Thursday, June 3, 2021
2:00pm – 5:00pm Eastern | 11:00am – 2:00pm Pacific
Agenda
Registration for this event is now closed. For more information, please contact helix@ebrc.org.
This workshop will draw upon our survey results to assess technical progress in the context of the research milestones predicted in Engineering Biology, the 2019 EBRC technical research roadmap. This information will shape a narrative of the most pressing needs and challenges still facing engineering biology over the next decade.
The workshop will inventory the degree of completion for each milestone, discuss technical and nontechnical barriers inhibiting progress, highlight new directions and avenues of research, and review social science dimensions associated with the development of technical goals. This will come in the form of a published report that will be adapted for public viewing.
These virtual writing workshops for each technical theme (3 hours each) are focused on drafting and revising the assessment. They are organized as followed:
The report will be invaluable to policymakers and funders to understand the continued challenges faced by our researchers; to researchers to learn the critical research gaps preventing engineering biology progression; and for our EBRC community to assess the utility of our roadmaps.
Hosted by Rebecca Nugent (Twist Bioscience) and Howard Salis (Penn State)
Please join us at a virtual workshop to assess the Biomolecular Engineering: Biomolecule, Pathway, and Circuit Engineering technical theme of Engineering Biology.
Tuesday, June 8, 2021
11:00am – 2:00pm Eastern | 8:00am – 11:00pm Pacific
Agenda
Registration for this event is now closed. For more information, please contact helix@ebrc.org.
This workshop will draw upon our survey results to assess technical progress in the context of the research milestones predicted in Engineering Biology, the 2019 EBRC technical research roadmap. This information will shape a narrative of the most pressing needs and challenges still facing engineering biology over the next decade.
The workshop will inventory the degree of completion for each milestone, discuss technical and nontechnical barriers inhibiting progress, highlight new directions and avenues of research, and review social science dimensions associated with the development of technical goals. This will come in the form of a published report that will be adapted for public viewing.
These virtual writing workshops for each technical theme (3 hours each) are focused on drafting and revising the assessment. They are organized as followed:
The report will be invaluable to policymakers and funders to understand the continued challenges faced by our researchers; to researchers to learn the critical research gaps preventing engineering biology progression; and for our EBRC community to assess the utility of our roadmaps.
Hosted by Chang Liu (UC Irvine) and Jesse Zalatan (UW)
Please join us for an exciting seminar on May 21, 2021, from 4-5:00 PM ET. This is the final seminar in the 2021 EBRC Seminar Series.
Speaker abstracts are below. The seminar is open to all, so please feel free to share this information with your colleagues.
The seminar will be held on Zoom using the following link for all sessions:
Zoom link: https://berkeley.zoom.us/j/97626552307?pwd=alVlS3dXM0lZYklYeE9zVXljWUI0UT09
Meeting ID: 976 2655 2307
Passcode: EBRC2021
“Sequence-function analysis helps identify multiple pathways to enhance Phenylalanine Ammonia-Lyase (PAL) activity”
Nikhil Unni Nair (Tufts University)
Phenylalanine ammonia-lyases (PALs) non-oxidatively deaminate L-phenylalanine to trans-cinnamic acid (tCA) and are widely found associated with secondary metabolism in plants, bacteria, and fungi. Biocatalytic applications for natural product and fine chemical synthesis has driven the discovery, expression, characterization, and engineering of PALs. More recently, development of PALs for phenylketonuria (PKU) management and cancer therapy has further increased interest in engineering this class of enzymes. While there is a general understanding of how residues in the substrate-binding pocket contribute to specificity and turnover, led by rational mutagenesis studies, there is generally a poor understanding of how distal residues affect function. In general, outcomes from directed evolution can identify distal hotspots but there have only been two such studies with this enzyme. The first study resulted in modest improvement in activity whereas the other, conducted by us, identified residues within the active site only. Deep mutational scanning (DMS) can identify functional hotspots, and when coupled with directed
evolution can accelerate engineering campaigns, although there are few examples of this approach. Further, DMS can provide a comprehensive map of sequence–function relationships to explore the protein fitness landscapes, uncover functionally relevant sites, improve molecular energy functions, and identify beneficial combinations of mutations for protein engineering. Though extensive body of research exists on function, structure, mechanism of PAL, a systematic study exploring the sequence-function space has not been attempted.
Previously, we developed a growth-coupled enrichment for rapid screening of high-activity variants of AvPAL* (used to formulate the PKU drug Pegvaliase®) in E. coli. After a single round of directed evolution using this growth-coupled enrichment, we identified 2 active site mutations improved kcat < 2-fold. However, the sequence-function fitness landscape of AvPAL* remains to be explored. In this study, we achieve several outcomes. First, we obtained the detailed sequence-function landscape of PAL, to date, using DMS, identifying >60 mutational hotspots. Next, we picked seven sites for comprehensive single and multi-site saturation mutagenesis and we identified multi-site mutations with ~2.5-fold improvement in the kcat (and >3-fold increase in catalytic efficiency). We then explored the epistatic effect of these mutations, uncovering positive, neutral, and negative interactions among distal and proximal sites. Finally, to understand the mechanistic role of key mutations in hyperactive variants, we performed modelling studies and concluded that there are multiple pathways to enhance PAL catalytic activity, including, decreased root mean square fluctuation (RMSF) of substrate in the active site, greater proximity of the substrate to catalytic residues, and facilitated diffusion of the substrate to the active site, among others. In summary, this study significantly advances basic and applied enzymology of PALs, a heretofore understudied class of enzymes with a wide array of applications.
Development of a yeast-based assay for bioavailable phosphorous
Heather Shepherd (University of Notre Dame)
Preventing eutrophication of inland freshwater ecosystems requires quantifying the phosphorous (P) content of the streams and rivers that feed them. Typical methods for measuring P assess soluble reactive P (SRP) or total P (TP) and require expensive analytical techniques that produce hazardous waste. Here we present a novel method for measuring the more relevant bioavailable P (BAP); this assay utilizes the growth of familiar baker’s yeast, avoids production of hazardous waste, and reduces cost relative to measurements of SRP and TP. The yeast BAP (yBAP) assay takes advantage of the observation that yeast density at saturating growth increases linearly with provided P. We show that this relationship can be used to measure P in freshwater in concentration ranges relevant to eutrophication. In addition, we measured yBAP in water containing known amount of fertilizer and in samples from agricultural waterways. We observed that the majority of yBAP values were between those obtained from standard SRP and TP measurements, demonstrating that the assay is compatible with real-world settings. The cost-effective and nonhazardous nature of the yeast-based assay suggests that it could have utility in a range of settings, offering added insight to identify water systems at risk of eutrophication from excess phosphorus.
Engineering alternative degradation tags for synthetic circuits
Prajakta Jadhav (South Dakota State University)
The goal in synthetic biology is to build robust synthetic circuits in bacteria that are dynamic, highly regulated, and result in a unified response. In the last 20 years, the synthetic biology field has effectively leveraged transcriptional (RNA production) and translational (protein production) controls. However, protein degradation plays an important role in determining the half-life of proteins and regulating biological systems. Amino acid degradation tags are exploited to avoid a reliance on cell division for protein dilution and to build dynamic circuits. The most leveraged
degradation tag in E. coli is the ssrA-tag, which is an 11-amino acid sequence that primarily target proteins for degradation by the ClpXP proteolytic system. However, the use of the ssrA-tag limits scalability and complexity especially in synthetic oscillators in bacteria due to itsmultiple proteolytic target recognition signals. Our goal is to build orthogonal oscillators that utilize degradation tags targeting to multiple proteases with no crosstalk. In this study, we aremodifying the ssrA-tag to change the substrate affinity and degradation rate and produce new
synthetic oscillators. We hypothesize that changing the degradation rate can alter the output signal of the oscillator. We have tailored the ssrA-tag to reduce crosstalk between proteolytic systems to increase robustness and developed a variety of degradation tags. While many of these tags exhibited decreased or no change in degradation rates, two depicted significant increase. We further tested interesting candidates for crosstalk between proteolytic systems and identified a tag that display minimum to no crosstalk with other proteolytic systems. We aim to build and compare the output signals of different oscillators with novel tags in batch cultures and at the single-cell level. The design and implementation of these novel degradation tags will enable development of biological building blocks for increased complexity in synthetic circuits.
Please join us for an exciting seminar on May 13, 2021, from 2-3:30 PM ET. This is the second of three seminars in the 2021 EBRC Seminar Series.
Speaker abstracts are below. The seminar is open to all, so please feel free to share this information with your colleagues.
The seminar will be held on Zoom using the following link for all sessions:
Zoom link: https://berkeley.zoom.us/j/97626552307?pwd=alVlS3dXM0lZYklYeE9zVXljWUI0UT09
Meeting ID: 976 2655 2307
Passcode: EBRC2021
TBD
Aindrila Mukhopadhyay (Lawrence Berkeley National Lab)
TBD
“Non-canonical crRNAs derived from host transcripts enable multiplexable RNA detection by Cas9”
Chunlei Jiao (Helmholtz Institute for RNA-based Infection Research)
CRISPR nucleases are guided by CRISPR RNAs (crRNAs) that are naturally derived from CRISPR arrays. Here, we discovered that crRNAs could derive from cellular transcripts outside of the CRISPR-Cas locus in Campylobacter jejuni, and we exploit this discovery to achieve a novel means of multiplexed RNA detection. The discovery came from sequencing RNAs that preferentially bound to the Cas9 in C. jejuni (CjeCas9), revealing an unexpected set of RNAs that shared a motif complementary to the anti-repeat of the system’s tracrRNA. The size and composition of these bound RNAs resembled that of crRNAs, indicating that the full-length version of these RNAs base pair with the anti-repeat portion of the tracrRNA and undergo processing to form crRNA-like RNAs. We call these processed RNAs non-canonical crRNAs (ncrRNAs). Using a cell-free transcription-translation (TXTL) system, we found that some ncrRNAs could drive efficient and sequence-specific DNA cleavage by CjeCas9 and its natural tracrRNA. Given the known sequence flexibility within the repeat:anti-repeat stem for single- guide RNAs, we hypothesized that the anti-repeat portion of the tracRNA could be reprogrammed to convert any RNA-of-interest into a functional ncrRNA that guides Cas9 to its DNA target. Using DNA cleavage assays in TXTL and in E. coli, we found that reprogrammed
tracrRNAs(Rptrs) designed to pair with different regions of an mRNA yielded efficient DNA cleavage not only for the CjeCas9 but also for the S. pyogenes Cas9 and the Streptococcus thermophilus CRISPR1 Cas9. Finally, based on the capability of Rptrs to link any RNA-of-interest to sequence-specific DNA cleavage, we established a multiplexed RNA diagnostic platform called LEOPARD (Leveraging Engineered tracrRNAs and On-target DNAs for PArallel RNA Detection). With LEOPARD, we achieved multiplexed detection of RNAs from different viruses including SARS-CoV-2 and other respiratory viruses in one reaction. We further distinguished SARS-CoV-2 and its D614G variant with single-nucleotide specificity in patient samples. These findings establish a previously unknown source of crRNAs and demonstrate the practical utility of LEOPARD for detecting numerous biomarkers in one test.
“Developing a mathematical framework for controlling complex biological systems”
Marcella Gomez (UC Santa Cruz)
In this talk, we refer to the achievement of an intended and predicted response in a biological system as controlling biology. Such efforts are often guided by classical mechanistic models from first principles. The level of complexity of these systems makes it extremely difficult to develop mechanistic models that can account for all possible interactions and predict biological response. To overcome these challenges, we propose to move away from first principal methods and instead identify key leverage points in the targeted biological pathways that can be directly up or down regulated by external signaling molecules. These molecules can be controlled by a feedback algorithm and delivered in situ by a bioelectronic device. In recent work, we successfully implemented feedback control on human‐induced pluripotent stem cells (hiPSCs) to regulate the cell’s resting potential known to affect cell physiology and functions such as proliferation, differentiation, migration, and apoptosis, as well as cell–cell communication and large‐scale morphogenesis. This was achieved without use of a model nor training data. We further outline an approach to extend the method to more complex biological systems. In particular, we consider the task of accelerating wound healing.
“The Promoter Calculator – A Sequence-to-Function Biophysical Model of Transcriptional Initiation for Sigma70 Promoters with Any Sequence”
Travis La Fleur (Penn State)
Engineering synthetic promoters with precision control has remained a challenge due to our inability to predict how a promoter’s sequence and DNA context controls its function and mRNA output. Here, we developed an accurate sequence-to-function model of transcriptional initiation that enables the automated design of synthetic promoters and the a priori prediction of cryptic promoters within natural systems – both of which are needed to advance Synthetic Biology towards genome-scale functional design. To do this, we combined oligopool synthesis, library-
based cloning, and next-generation sequencing to construct and characterize 14,206 rationally designed sigma70 promoters in vitro. Measurements include transcriptional start site frequencies and overall mRNA levels. This approach enables highly-parallel characterization of constitutive promoter activity without confounding factors such as unintentional transcriptional regulation, non-sigma70 activity, and mRNA decay. These measurements, in combination with machine learning, were used to parameterize a thermodynamic model quantifying the
interactions controlling transcription rate. For demonstration, the “Promoter Calculator” was used to accomplish three major tasks – accurate prediction of thousands of sigma70 promoters across various conditions, the de novo design of novel sigma70 promoters, and the identification of cryptic promoters internal to a genetic circuit. 4,350 highly non-repetitive promoters and 6,165 genome-integrated promoters characterized in vivo were accurately predicted by the model with Spearman Rank-Order Coefficients of .68 and .78, respectively. Promoters designed de novo using the Promoter Calculator were characterized in vivo exhibiting a 683-fold range in expression resulting in a Pearson Correlation Coefficient of .85. The Promoter Calculator was used to analyze a genetic circuit containing 11 circuit promoters and 29 cryptic promoters. The model identified 29 sigma70 promoters out of 40 observed promoters (72.5% accurate), including 10/11 circuit promoters and 19/29 cryptic promoters. The Promoter Calculator facilitates context-aware, rational promoter design without relying on a fixed table of pre-characterized sequences while serving as a powerful tool in promoter identification.
Please join us for an exciting seminar on May 4, 2021, from 3-4:00 PM ET. This is the first of three seminars in the 2021 EBRC Seminar Series.
Speaker abstracts are below. The seminar is open to all, so please feel free to share this information with your colleagues.
The seminar will be held on Zoom using the following link for all sessions:
Zoom link: https://berkeley.zoom.us/j/97626552307?pwd=alVlS3dXM0lZYklYeE9zVXljWUI0UT09
Meeting ID: 976 2655 2307
Passcode: EBRC2021
“Engineering multilevel CRISPR-based kill-switches for probiotic Escherichia coli“
Austin Rottinghaus (Washington University in St. Louis)
Probiotic microbes have become an effective framework for diagnostic and therapeutic technologies. However, there are safety concerns associated with using genetically engineered organisms for medical applications. Probiotic microbes have the potential to evolve growth advantages over natural microbes and characteristics that are harmful to the host or to the outside environment. To mitigate these concerns, we engineered the probiotic Escherichia coli Nissle 1917 to survive only when and where it is needed using CRISPR-based kill-switches (CRISPRks). We first designed a CRISPRks that induces cell death by expressing Cas9 and genome-targeting guide RNAs in response to the chemical inducer anhydrotetracycline. This design allows cell killing to occur while the microbe is in the gut in response to oral administration of the chemical. We optimized the efficiency and stability of the CRISPRks by combining four genomic Cas9 expression cassettes with three plasmid-based guide RNA expression cassettes, removing the antibiotic dependence for maintenance of the guide RNA plasmid, and knocking out genes involved in DNA recombination and mutagenesis. Using this optimized circuit in vitro, we achieved more than a 9-log reduction in cell number and demonstrated genetic stability for up to 28 days of continuous growth. This high killing efficiency was maintained in vivo, where we achieved complete elimination of the probiotic 24 hours after oral administration of the inducer. This is the first time on-demand elimination of an engineered microbe has been demonstrated in
vivo. We next modified our chemically inducible-CRISPRks to also induce cell death in response to ambient temperatures below 33’C. This two-input design induces cell killing either in response to oral administration of the chemical or when the microbe is excreted from the body in response to the reduced environmental temperature. This two-input circuit achieved more than a 9-log and 7-log reduction in cell number in vitro after exposure to the chemical inducer and temperature downshift, respectively. Future directions will include incorporating the CRISPRks in microbes engineered to diagnose and treat diverse medical conditions. Our CRISPRks strategy provides a template for future microbial biocontainment circuits. The sensor and killing mechanism employed in the kill-switch are well characterized and functional in many microbes, allowing the CRISPRks design to be broadly utilized. In addition, the temperature-sensing module can be easily replaced with sensors that recognize alternative signals, allowing comparable kill-switches to be created for applications beyond medicine.
“Substrate-Activated Expression of a Biosynthetic Pathway in Escherichia coli”
Cynthia Ni (MIT)
Microbial production leverages endogenous and heterologous enzymes to produce value-added chemicals. Overexpression of the genes encoding pathway enzymes can impose a metabolic burden to the host. There are many approaches to alleviating this burden, including using chemical inducers, such as IPTG, to delay expression, expressing genes from stationary phase promoters, or using feedback controllers that activate expression in response to a pathway intermediate. We developed a substrate-activated, feed-forward expression control strategy in which the necessary substrate of the pathway doubles as the inducer of heterologous pathway gene expression. We demonstrated this strategy on a D-glyceric acid pathway that utilizes galacturonate as a feed substrate. A galacturonate-responsive transcription factor was used to construct a galacturonate responsive biosensor. We constructed variants of the biosensor and selected the best performer through fluorescence characterization. The selected biosensor variant was used to control the heterologous gene expression of the D-glyceric acid biosynthetic pathway. We confirmed that expression was induced in presence of the substrate through qRT-PCR.
Production via substrate-induction with our expression control circuit was comparable to IPTG- controlled induction and significantly outperformed constitutive expression. Our work demonstrates that substrate-activated pathway expression is an attractive control strategy for microbial production.
“Robust direct digital-to-biological data storage in living cells”
Sung Sun Yim (Columbia University)
DNA has been the predominant information storage medium for biology and holds great promise as a next-generation high-density data medium in the digital era. Currently, the vast majority of DNA-based data storage approaches rely on in vitro DNA synthesis. As such, there are limited methods to encode digital data into the chromosomes of living cells in a single step. In this talk, I will describe a new electrogenetic framework for direct storage of digital data in living cells using an engineered redox-responsive CRISPR adaptation system. We demonstrated multiplex data encoding into barcoded cell populations to yield meaningful information storage and capacity up to 72 bits, which can be maintained over many generations in natural open environments. In addition, I will share our recent effort on directed evolution of CRISPR adaptation machineries to improve data storage capacity and port the system into other bacteria, thus enabling new applications of DNA-based cellular recording.