CRISPR Gene Editing in Yeast: An Experimental Protocol for an Upper-Division Undergraduate Laboratory Course

An undergraduate research experience in CRISPR-Cas9 mediated eukaryotic genome editing to teach fundamental biochemistry techniques

CRISPR-Cas9 technology is an established, powerful tool for genome editing through the ability to target specific DNA sequences of interest for introduction of desired genetic modifications. CRISPR-Cas9 is utilized for a variety of purposes, ranging from a research molecular biology tool to treatment for human diseases. Due to its prominence across a variety of applications, it is critical that undergraduates in the life sciences are educated on CRISPR-Cas9 technology. To this end, we created an intensive eight-week long course-based undergraduate research experience (CURE) designed for students to understand CRISPR-Cas9 genome editing and perform it in Saccharomyces cerevisiae. Students enrolled in the CURE participate in 2, 3-h sessions a week and are engaged in the entire process of CRISPR-Cas9 genome editing, from preparation of genome editing reagents to characterization of mutant yeast strains. During the process, students master fundamental techniques in the life sciences, including sterile technique, Polymerase Chain Reaction (PCR), primer design, sequencing requirements, and data analysis. The course is developed with flexibility in the schedule for repetition of techniques in the event of a failed experiment, providing an authentic research experience for the students. Additionally, we have developed the course to be easily modified for the editing of any yeast gene, offering the potential to expand the course in research-driven classroom or laboratory settings.

A genetic screen of transcription factors in the Drosophila melanogaster abdomen identifies novel pigmentation genes

Gene regulatory networks specify the gene expression patterns needed for traits to develop. Differences in these networks can result in phenotypic differences between organisms. Although loss-of-function genetic screens can identify genes necessary for trait formation, gain-of-function screens can overcome genetic redundancy and identify loci whose expression is sufficient to alter trait formation. Here, we leveraged transgenic lines from the Transgenic RNAi Project at Harvard Medical School to perform both gain- and loss-of-function CRISPR/Cas9 screens for abdominal pigmentation phenotypes. We identified measurable effects on pigmentation patterns in the Drosophila melanogaster abdomen for 21 of 55 transcription factors in gain-of-function experiments and 7 of 16 tested by loss-of-function experiments. These included well-characterized pigmentation genes, such as bab1 and dsx, and transcription factors that had no known role in pigmentation, such as slp2. Finally, this screen was partially conducted by undergraduate students in a Genetics Laboratory course during the spring semesters of 2021 and 2022. We found this screen to be a successful model for student engagement in research in an undergraduate laboratory course that can be readily adapted to evaluate the effect of hundreds of genes on many different Drosophila traits, with minimal resources.

CRISPR/Cas9 gene targeting plus nanopore DNA sequencing with the plasmid pBR322 in the classroom

Both nanopore-based DNA sequencing and CRISPR/Cas-based gene editing represent groundbreaking innovations in molecular biology and genomics, offering unprecedented insights into and tools for working with genetic information. For students, reading, editing, and even writing DNA will be part of their everyday life. We have developed a laboratory procedure that includes (i) the biosynthesis of a guide RNA for, (ii) targeting Cas9 to specifically linearize the pBR322 plasmid, and (iii) the identification of the cutting site through nanopore DNA sequencing. The protocol is intentionally kept simple and requires neither living organisms nor biosafety laboratories. We divided the experimental procedures into separate activities to facilitate customization. Assuming access to a well-equipped molecular biology laboratory, an initial investment of approximately $2,700 is necessary. The material costs for each experiment group amount to around $130. Furthermore, we have developed a freely accessible website (https://dnalesen.hs-mittweida.de) for sequence read analysis and visualization, lowering the required computational skills to a minimum. For those with strong computational skills, we provide instructions for terminal-based data processing. With the presented activities, we aim to provide a hands-on experiment that engages students in modern molecular genetics and motivates them to discuss potential implications. The complete experiment can be accomplished within half a day and has been successfully implemented by us at high schools, in teacher training, and at universities. Our tip is to combine CRISPR/Cas gene targeting with nanopore-based DNA sequencing. As a tool, we provide a website that facilitates sequence data analysis and visualization.

Overexpression of the FBA and TPI genes promotes high production of HDMF in Zygosaccharomyces rouxii

4-Hydroxy-2,5-dimethyl-3 (2H)-furanone (HDMF) is widely used in the food industry as a spice and flavoring agent with high market demand. In this study, fructose-1,6-bisphosphate aldolase (FBA) and triose phosphate isomerase (TPI) were overexpressed in Zygosaccharomyces rouxii in the form of single and double genes, respectively, via electroporation. High-yield HDMF-engineered yeast strains were constructed by combining the analysis of gene expression levels obtained by real-time fluorescence quantitative PCR technology and HDMF production measured by HPLC. The results showed that there was a significant positive correlation between the production of HDMF and the expression levels of the FBA and TPI genes in yeast; the expression levels of the FBA and TPI genes were also positively correlated (p < 0.05). Compared with the wild type (WT), the engineered strains F10-D, T17-D, and TF15-A showed marked increases in HDMF production and FBA and TPI gene expression (p < 0.05) and exhibited great genetic stability with no obvious differences in biomass or colony morphology. In addition, the exogenous addition of d-fructose promoted the growth of Z. rouxii. Among the engineered strains, when fermented in YPD media supplemented with d-fructose for 5 days, TF15-A (overexpressing the FBA and TPI genes) generated the highest HDMF production of 13.39 mg/L, which is 1.91 times greater than that of the wild-type strain. The results above indicated that FBA and TPI, which are key enzymes involved in the process of HDMF biosynthesis by Z. rouxii, positively regulate the synthesis of HDMF at the transcriptional level. d-fructose can be used as a precursor for the biosynthesis of HDMF by engineered yeast in industrial production.

Long non-coding RNA and ribosomal protein genes in a yeast ageing model: an investigation for undergraduate research-based learning

The unicellular yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe are widely used eukaryotic model organisms. Research exploiting the tractability of these model systems has contributed significantly to our understanding of a wide range of fundamental processes. In this article, we outline the features of yeast that have similarly been exploited for undergraduate research training. We selected examples from published literature that demonstrate the utility of the yeast system for research-based learning embedded in the curriculum. We further describe a project which we designed for the team-based final-year dissertation projects module on our transnational joint programme, which investigates whether the expression and functions of the budding yeast RPL36 ribosomal protein paralogs are influenced by the overlapping long non-coding RNA genes. Students carry out the experimental procedures in a 2-week timetabled teaching block and exercise widely applicable biochemical techniques, including aseptic yeast cell culture and sample collection, RNA isolation, qRT-PCR quantitation, protein extraction and Western blot analysis, and cell cycle progression patterns using light microscopy and flow cytometry. It is challenging to design training programmes for undergraduates that are meaningful as well as practical and economical, but it is possible to transform active research projects into authentic research experiences. We consider yeast to be an ideal model organism for such projects. These can be adapted to the constraints of course schedules and explore fundamental biochemical topics which are evolutionarily conserved from yeast to mammals.

CRISPR in butterflies: An undergraduate lab experience to inactivate wing patterning genes during development

CRISPR is a technique increasingly used in the laboratory for both fundamental and applied research. We designed and implemented a lab experience for undergraduates to carry out CRISPR technology in the lab, and knockout the wing patterning genes optix and WntA in Vanessa cardui butterflies. Students obtained spectacular phenotypic mutants of butterfly wings color and patterns, awakening curiosity about how genomes encode morphology. In addition, students successfully used molecular techniques to genotype and screen wild-type caterpillar larvae and butterflies for CRISPR edits in genes. Student feedback suggests that they experienced a meaningful process of scientific inquiry by carrying out the whole CRISPR workflow process, from the design and delivery of CRISPR components through microinjection of butterfly eggs, the rearing of live animals through their complete life cycle, and molecular and phenotypic analyses of the resulting mutants. We discuss our experience using CRISP genome editing experiments in butterflies to expose students to hands-on research experiences probing gene-to-phenotype relationships in a charismatic and live organism.

An experimental protocol for teaching CRISPR/Cas9 in a post-graduate plant laboratory course: An analysis of mutant-edited plants without sequencing

The CRISPR/Cas9 system is widely used for editing genes in various organisms and is a very useful tool due to its versatility, simplicity, and efficiency. To teach its principles to post-graduate students we designed a laboratory activity to obtain and analyze PDS3 mutants in Arabidopsis thaliana plants consisting of: 1) Design of guide RNAs using bioinformatics tools; 2) plant transformation (which is optional depending on the length of the course); 3) observation and evaluation of the mutant's phenotypes in the Phytoene desaturase (PDS3) gene, which exhibit an albino phenotype and different degrees of mosaicism in the editing events we evaluated; 4) PCR amplification of a fragment that includes the mutated region followed by analysis of single-stranded DNA conformation polymorphisms (SSCP) using native polyacrylamide gel electrophoresis and silver nitrate staining to detect changes in the amplicon sequence due to gene editing. Through SSCP, the students were able to distinguish between homozygous and heterozygous edited plants. A highlight feature of this protocol is the visualization and detection of the mutation/edition without sequencing the edited fragment.

An experimental protocol for molecular biology lab at an Amazonian University

Laboratory-based practical classes are an essential component in teaching molecular biology for undergraduate students. Universidad Regional Amazonica Ikiam is a higher education institution located in the Ecuadorian Amazon rainforest, a high biodiversity place, including amphibians. Based on this, we have established a practical molecular biology program with eight sessions that contextualize the biodiverse surroundings of the University. This program stimulates synchronization of information between theory and practice and improves research skills. During these sessions, students are motivated to identify and characterize antimicrobial peptides from Ecuadorian frog skin secretions, using molecular biology techniques and biochemistry and microbiology knowledge. This practical course was held twice with a total of 56 students from the fifth semester of the biotechnology engineering. The evaluation of the practical program was carried out through a questionnaire applied to students using the Likert scale. Overall, this form of teaching had high receptivity and presented benefits for student learning. Interestingly, 80% of respondents strongly agreed that this course provided tools and knowledge for the development of their undergraduate dissertation. Therefore, practical courses tailored to the student's context can stimulate student learning and interest. Additionally, this experimental methodology is interdisciplinary and can be applied to other research fields and subjects.

CRISPR in Your Kitchen: an At-Home CRISPR Kit to Edit Genes in Saccharomyces cerevisiae Used during a Remote Lab Course

The use of CRISPR-based experiments in an undergraduate course is appealing because of the ease of editing, and the relevance of CRISPR to current research. Before the COVID-19 pandemic, we developed an in-person lab for a high-enrollment course that allowed students to design and conduct CRISPR editing experiments in budding yeast, Saccharomyces cerevisiae. Post pandemic, the lab course moved online, and we lost the hands-on component. We subsequently developed an at-home kit that contained all the necessary materials for students to grow and transform S. cerevisiae with the DNA molecules necessary for CRISPR-Cas9 induced editing. Our at-home kits cost approximately $70 each to produce and were shipped to over 600 students during the 2020 to 2021 academic year. By adding the at-home experimental work to our remote, online lab course, students were able to generate loss-of-function mutants in ADE2 (causing a red color phenotype). Students were able to send edited yeast samples back to campus for sequencing, allowing for characterization of the different mutations that can occur due to CRISPR-Cas9 induced editing. Here, we described the protocol to produce and use the kits and summarized the student experience of using the at-home kit in a large enrollment, remote lab course. These kits provided opportunities to engage students in hands-on experimentation during a remote course and could also be used to reach learners in other domains, such as high schools and outreach programs.

Implementing CRISPR-Cas9 Yeast Practicals into Biology Curricula

CRISPR-Cas9 is a genome-editing technique that has been widely adopted thanks to its simplicity, efficiency, and broad application potential. Due to its advantages and pervasive use, there have been attempts to include this method in the existing curricula for students majoring in various disciplines of biology. In this perspective, we summarize the existing CRISPR-Cas courses that harness a well-established model organism: baker's yeast, Saccharomyces cerevisiae. As an example, we present a detailed description of a fully hands-on, flexible, robust, and cost-efficient practical CRISPR-Cas9 course, where students participate in yeast genome editing at every stage-from the bioinformatic design of single-guide RNA, through molecular cloning and yeast transformation, to the final confirmation of the introduced mutation. Finally, we emphasize that in addition to providing experimental skills and theoretical knowledge, the practical courses on CRISPR-Cas represent ideal platforms for discussing the ethical implications of the democratization of biology.

CRISPR-Cas9 Gene Editing in Yeast: A Molecular Biology and Bioinformatics Laboratory Module for Undergraduate and High School Students

CRISPR-Cas9 genome editing technology is widely used in scientific research and biotechnology. As this technology becomes a staple tool in life sciences research, it is increasingly important to incorporate it into biology curricula to train future scientists. To demonstrate the molecular underpinnings and some limitations of CRISPR-based gene editing, we designed a laboratory module to accompany a discussion-based course on genome editing for college and advanced high school biology students. The laboratory module uses CRISPR-Cas9 to target and inactivate the ADE2 gene in Saccharomyces cerevisiae so as to give red colonies, employing an inexpensive yeast model system with a phenotypic readout that is easily detectable without specialized equipment. Students begin by accessing the yeast ADE2 sequence in a genome database, applying their understanding of Cas9 activity to design guide RNA (gRNA) sequences, using a CRISPR analysis tool to compare predicted on- and off-target effects of various gRNAs, and presenting and explaining their choice of an optimal gRNA to disrupt the ADE2 gene. They then conduct yeast transformations using Cas9 and preselected gRNA plasmids with or without donor templates to explore the importance of DNA repair pathways in genome editing. Lastly, they analyze the observed editing rates across different gRNAs targeting ADE2, leading to a discussion of editing efficiency. This module engages students in experimental design, provides hands-on experience with CRISPR-Cas9 gene editing and collaborative data analysis, and stimulates discussion on the uses and limitations of CRISPR-based gene editing technology.

The Challenges of Teaching Anatomy and Physiology Laboratory Online in the Time of COVID-19

In early spring 2020, New York's K-12 schools, colleges, and universities went into lockdown in response to the COVID-19 pandemic, and faculty began converting face-to-face classes into distance learning classes. Teachers and college faculty learned to use new technology to continue instruction for the remainder of the academic year. Learning curves were steep for some, as using technology proved to be overwhelming. Students who registered for in-person classes had to learn in an online environment. Pre-Health Science majors who rely heavily on practical, hands-on courses to gain the necessary skills required for their career participated in virtual anatomy and physiology laboratories. Clearly, face-to-face courses are not always possible; therefore, it is imperative to know the best practices of online teaching and learning.

"Designer babies?!" A CRISPR-based learning module for undergraduates built around the CCR5 gene

CRISPR-cas technology is being incorporated into undergraduate biology curriculum through lab experiences to immerse students in modern technology that is rapidly changing the landscape of science, medicine and agriculture. We developed and implemented an educational module that introduces students to CRISPR-cas technology in a Genetic course and an Advanced Genetics course. Our primary teaching objective was to immerse students in the design, strategy, conceptual modeling, and application of CRISPR-cas technology using the current research claim of the modification of the CCR5 gene in twin girls. This also allowed us to engage students in an open conversation about the bioethical implications of heritable germline and non-heritable somatic genomic editing. We assessed student-learning outcomes and conclude that this learning module is an effective strategy for teaching undergraduates the fundamentals and application of CRISPR-cas gene editing technology and can be adapted to other genes and diseases that are currently being treated with CRISPR-cas technology.

Using Metabolic Engineering to Connect Molecular Biology Techniques to Societal Challenges

Genetically modified organisms (GMOs) are a topic of broad interest and are discussed in classes ranging from introductory biology to bioethics to more advanced methods-focused molecular biology courses. In most cases, GMOs are discussed in the context of introducing a single protein-coding gene to produce a single desired trait in a crop. For example, a commercially available kit allows students to test whether food products contain GMOs by detecting the Bacillus thuringiensis delta-endotoxin gene, which confers resistance to European corn borers. We have developed an 8-week laboratory module for upper-division undergraduates and graduate students that builds upon students’ basic understanding of GMOs to introduce them to the techniques used to sustainably produce commercially valuable products in yeast through metabolic engineering. In this course, students use recombination-based methods to assemble genes encoding entire metabolic pathways in Saccharomyces cerevisiae, perform genetic screens to identify yeast genes that impact metabolite yield, and use error-prone PCR to optimize metabolic pathway function. In parallel to these laboratory-based activities, students engage with the societal impact of these approaches through case studies of products made via yeast metabolic engineering, such as opioids, omega-3 fatty acids, and the Impossible Burger. In this report, we focus on these case studies as well as an individual sustainability project assignment created for this course. This assignment, which spans the 8-week module, asks students to find examples of yeast metabolic engineering that could be used to address current sustainability challenges in their communities. By the end of the course, students synthesize this information to create a case study that could be used to teach concepts related to metabolic engineering and sustainability to their peers. Student approaches to this project have varied from literature reviews, to news searches, to directly contacting and interviewing researchers using novel metabolic engineering approaches. These student-produced projects are used as case studies in future semesters, amplifying student voices and contributing to student ownership. While developed in the context of this course, the sustainability project and case studies are broadly applicable and could be adapted for use in biology or bioethics courses at the undergraduate or graduate level. Through this report, we hope to gain collaborators interested in implementing a version of the course at their institutions, allowing for robust assessment of the impact of the course on a larger group of students.

Bringing immersive science to undergraduate laboratory courses using CRISPR gene knockouts in frogs and butterflies

The use of CRISPR/Cas9 for gene editing offers new opportunities for biology students to perform genuine research exploring the gene-to-phenotype relationship. It is important to introduce the next generation of scientists, health practitioners and other members of society to the technical and ethical aspects of gene editing. Here, we share our experience leading hands-on undergraduate laboratory classes, where students formulate hypotheses regarding the roles of candidate genes involved in development, perform loss-of-function experiments using programmable nucleases and analyze the phenotypic effects of mosaic mutant animals. This is enabled by the use of the amphibian Xenopus laevis and the butterfly Vanessa cardui, two organisms that reliably yield hundreds of large and freshly fertilized eggs in a scalable manner. Frogs and butterflies also present opportunities to teach key biological concepts about gene regulation and development. To complement these practical aspects, we describe learning activities aimed at equipping students with a broad understanding of genome editing techniques, their application in fundamental and translational research, and the bioethical challenges they raise. Overall, our work supports the introduction of CRISPR technology into undergraduate classrooms and, when coupled with classroom undergraduate research experiences, enables hypothesis-driven research by undergraduates.

A Scalable CURE Using a CRISPR/Cas9 Fluorescent Protein Knock-In Strategy in Caenorhabditis elegans

Genome editing with CRISPR/Cas9 technology has advanced from the lab bench to clinical application with multiple trials underway. This article introduces a course-based undergraduate experience (CURE) combining CRISPR/Cas9 genome editing (using a modified two-plasmid system) and the animal model Caenorhabditis elegans. This CURE is designed to be a scalable, semester-long laboratory that will introduce the students to literature searches, molecular biology, experiment planning, microscopy, CRISPR bioethics discussion, and scientific writing. Here, students challenged themselves to endogenously tag the C. elegans gene zmp-4, a matrix metalloproteinase enzyme, with a fluorescent protein marker and successfully generated a new worm strain. The knock-in was confirmed with genotyping and imaging and will be available for use by the entire worm community.

An undergraduate laboratory module that uses the CRISPR/Cas9 system to generate frameshift mutations in yeast

The CRISPR/Cas9 system is a powerful tool for gene editing and it has become increasingly important for biology students to understand this emerging technique. Most CRISPR laboratory teaching modules use complex metazoan systems or mammalian cell culture which can be expensive. Here, we present a lab module that engages students in learning the fundamentals of CRISPR/Cas9 methodology using the simple and inexpensive model system, Saccharomyces cerevisiae. Students use CRISPR/Cas9 and nonhomologous end joining to generate frameshift insertion and deletion mutations in the CAN1 gene, which are easily selected for using media plates that have canavanine. DNA sequencing is also performed to determine what type of mutation occurred in gene-edited cells. This easy to implement set of experiments has been run as both a 5-week and a shorter 3-week lab module. Learning assessments demonstrate increased understanding in CRISPR-related concepts as well as increased confidence using molecular techniques. Thus, this CRISPR/Cas9 lab module can be added to an existing Genetics, Microbiology, or Molecular Biology lab course to help undergraduate students learn current gene editing techniques with limited effort and cost. © 2019 International Union of Biochemistry and Molecular Biology, 47(5):573-580, 2019.