Author Profile

Using CRISPR/Cas9 genome editing technology as an educational tool to study primary cilia in mammalian cell culture

Jonathon Walsh, University of Georgia
Jonathan Eggenschwiler, PhD, University of Georgia
Brian Condie, PhD, University of Georgia
Karl Lechtreck, PhD, University of Georgia
Location: Georgia

Abstract

This is a three semester-long "project lab" that engages undergraduate students to conduct real, novel research to address fundamental questions in cell biology. The project lab is a team-driven effort that involves one overarching scientific goal with each student being responsible for part of the project to reach that goal. All students are trained in a common set of cutting-edge methods and techniques and each student applies this towards studying the role of an individual gene involved in assembly and maintenance of cilia.

Student Goals

  1. Students gain a general understanding of a fundamental cell biological process: the assembly of an organelle, the cilium, serving functions ranging from motility, movement of fluid, to sensing the extracellular environment
  2. Students gain firsthand experience using a systematic approach to studying basic biological questions and developing alternate strategies/questions based on results acquired throughout the process i.e. troubleshooting experiments.
  3. Students gain a deeper understanding of basic genetic concepts through firsthand experience such as genotype/phenotype relationships, mechanisms of DNA repair, homologous recombination, mutagenesis, and epistasis.

Research Goals

  1. Students construct the molecular tools required for generating mutations in genes involved in cilia biogenesis and regulation.
  2. Students generate mutations of target genes in cell culture and identify cilia related phenotypes using various imaging techniques.

Context

This is a 3-semester research course designed for students to fulfill the 3-semester research requirement for an honors thesis project at UGA. Normally, this is done with a single student under the mentorship of a faculty member with or without the aid of a graduate student. For this course, 6 students were recruited to pilot the experience. Students will need to have a basic understanding of molecular biology and genetics to be successful in the course. Some lab experience would be helpful but is not required. This course was specifically designed with 2nd or 3rd year undergraduate students in mind.

Target Audience:Major
CURE Duration:Multiple terms

CURE Design

A simple application of CRISPR/Cas9 mutagenesis is employed to generate mutations in a set of genes implicated in ciliogenesis. For some of these, the ciliary phenotypes can be predicted fairly well and for others, it is less clear. Mutations are generated in a cell line, mIMCD3, which generates long cilia in culture and which carries an Ift88::YFP reporter cassette (inserted into the genome) that will allow the use of microscopy to analyze phenotypes. The students work in pairs, or teams, with each pair working on a set of single or double mutations. There are specific 'checkpoints' in place that each pair needs to reach before moving on to the next step of the project. If one team reaches a checkpoint before the others, they help the other teams reach the checkpoint. Once all teams have reached the checkpoint, the next steps are discussed as a class and plans for reaching the next checkpoint are made.

Core Competencies:
Nature of Research:Basic Research, Wet Lab/Bench Research

Tasks that Align Student and Research Goals

Research Goals →
Student Goals ↓
Research Goal 1: Students construct the molecular tools required for generating mutations in genes involved in cilia biogenesis and regulation.
Research Goal 2: Students generate mutations of target genes in cell culture and identify cilia related phenotypes using various imaging techniques.

Student Goal 1: Students gain a general understanding of a fundamental cell biological process: the assembly of an organelle, the cilium, serving functions ranging from motility, movement of fluid, to sensing the extracellular environment

1) Conduct phenotype analysis using various microscopic techniques including fixed cell staining/fluorescent imaging and live-cell imaging of ciliary transport.
2) Use scientific literature to understand why specific genes were chosen for mutagenesis with respect to their roles in ciliogenesis.


Student Goal 2: Students gain firsthand experience using a systematic approach to studying basic biological questions and developing alternate strategies/questions based on results acquired throughout the process i.e. troubleshooting experiments.

1) Design the genome targeting sequence used to mutate their gene of interest using online bioinformatics tools.
2) Use molecular techniques such as restriction enzyme digests, gel electrophoresis, molecular cloning, and PCR to generate the necessary DNA constructs for utilizing CRISPR/Cas9 mutagenesis in cell culture.
3) Use Sanger DNA sequencing to validate successful generation of the targeting constructs.

1) Employ mammalian cell culture techniques for the maintenance and propagation of adherent stable cells in culture.
2) Carry out DNA transfection (lipofection) and chemical selection for transfection of a selectable marker, generation of clonal cell lines, and fluorescence activated cell sorting (FACS).


Student Goal 3: Students gain a deeper understanding of basic genetic concepts through firsthand experience such as genotype/phenotype relationships, mechanisms of DNA repair, homologous recombination, mutagenesis, and epistasis.

1) Describe what the CRISPR/Cas9 system is, how it is used, and where it comes from. 
2) Explain the utility of homology directed repair and how this concept relates to the design of the constructs for CRISPR/Cas9-mediated homologous recombination.
3) Summarize basic concepts related to genotypes/phenotypes in the context of creating mutations.

1) Conduct PCR genotyping and sequence analysis for the identification of mutations.
2) Generate double mutants (mutations in 2 separate genes in the same cell) and the concept of epistasis/genetic interactions. 
3) Explore off-target mutations and genetic complementation.


Instructional Materials

Student Syllabus (Microsoft Word 2007 (.docx) 17kB Jun27 18)
Semester Workflow (Microsoft Word 2007 (.docx) 14kB Jun27 18)

Assessment

Student Assessment and Survey (Microsoft Word 2007 (.docx) 75kB Jun27 18)

Instructional Staffing

The three faculty (Jonathan Eggenschwiler, Brian Condie, and Karl Lechtreck) were involved in the bi-weekly class meetings designed to introduce the students to reading literature related to the class topic and to discuss the research progress and troubleshooting experiments. Jonathon Walsh (graduate student) was involved in the day-to-day experiments and training students in the laboratory with assistance from Jonathan Eggenschwiler. Jonathon Walsh also developed the course lab manual based on protocols developed during his PhD research. Jonathon attended the bi-weekly meetings and often led the class discussions, literature reviews, and student progress reporting.

Author Experience

Jonathon Walsh, University of Georgia

CRISPR/Cas9 mutagenesis is an innovative tool for science research and has a huge potential for student learning in a laboratory setting. The range of techniques that can be employed to effectively use this tool in a biological system lends a unique, systematic approach to introducing students to multiple concepts and methodologies in fields of genetics and molecular biology.


Advice for Implementation

This was a large undertaking for a graduate student and it may not be advised unless it is a portion of their graduate thesis. This course was designed for the students to work in pairs or asynchronously throughout the three semesters, which made it a huge time commitment for the graduate student and faculty involved. It would be much less cumbersome if it could be designed where students had designated time to be in lab each day.


Working with undergraduate students on a project using mammalian cell culture as a model became a problem and we would suggest using a simpler model for future applications of this type of CURE. This would alter the methodologies quite a bit, but would allow for further student progress. Students were delayed for a large portion of the 2nd semester due to contamination issues, which did not allow them to progress to the final stage of the project (phenotype analysis).


Students are not used to the 'real science' experience and early on get very upset when experiments do not work and immediately blame themselves for not being good enough or experienced enough to avoid mistakes. When they start out, they do not yet realize that troubleshooting experiments is a huge part of research. Throughout the course, the instructors were always positive when addressing such concerns and constantly reinforced the idea that often it is not the researcher's fault when an experiment fails and that they need to think through the methodologies and determine where the problem occurred and how to approach solving that problem. With this approach, the students' confidence in the laboratory increased significantly throughout the course series.

Iteration

The open-ended nature of this project allowed the students to take part in troubleshooting/problem solving throughout the course series. This was both with respect to experiment failure because of experimenter mistakes, as well as needing to optimize experimental conditions. For example, a student pair failed to ligate a construct into a plasmid and needed assistance. We discussed basic troubleshooting strategies for optimizing ligation efficiencies. Students had to design PCR primers and optimize them for generating clean products for sequencing reactions. Transfections of plasmids into mammalian cells can take much troubleshooting with regards to the ratio of DNA to lipofectamine. The phenotype analysis could be different or require different levels of optimization depending on the gene being mutated and its expected phenotype.

Using CURE Data

All data produced by students was recorded in their laboratory notebooks and digital data was stored on shared cloud storage space managed by the instructors. All data collected was vetted for accuracy and reproducibility by the instructors.

Resources

General reviews on Ciliogenesis:
Ishikawa, H.; Marshall, W. F. Ciliogenesis: Building the Cell's Antenna. Nat. Rev. Mol. Cell Biol. 2011, 12 (4), 222–234.
Oh, E. C.; Katsanis, N. Cilia in Vertebrate Development and Disease. Development 2012, 139 (3), 443–448.

Techniques: CRISPR/Cas9:
Mali, P.; Yang, L.; Esvelt, K. M.; Aach, J.; Guell, M.; DiCarlo, J. E.; Norville, J. E.; Church, G. M. RNA-Guided Human Genome Engineering via Cas9. Science 2013, 339 (6121), 823–826.
Liu, L.; Fan, X.-D. CRISPR–Cas System: A Powerful Tool for Genome Engineering. Plant Mol. Biol. 2014, 85 (3), 209–218.
Sampson, T. R.; Weiss, D. S. Exploiting CRISPR/Cas Systems for Biotechnology. Bioessays 2014, 36 (1), 34–38.

Techniques: Imaging:
Besschetnova, T. Y.; Roy, B.; Shah, J. V. Imaging Intraflagellar Transport in Mammalian Primary Cilia. Methods Cell Biol. 2009, 93 (08), 331–346.
Ott, C.; Lippincott-schwartz, J. Visualization of Live Primary Cilia Dynamics Using Fluorescence Microscopy. 2013.
MARTIN-FERNANDEZ, M. L.; TYNAN, C. J.; WEBB, S. E. D. A 'Pocket Guide' to Total Internal Reflection Fluorescence. J. Microsc. 2013, 252 (1), 16–22.

Publications on specific genes:
Huangfu, D.; Liu, A.; Rakeman, A. S.; Murcia, N. S.; Niswander, L.; Anderson, K. V. Hedgehog Signalling in the Mouse Requires Intraflagellar Transport Proteins. Nature 2003, 426 (6962), 83–87.
Walczak-Sztulpa, J.; Eggenschwiler, J.; Osborn, D.; Brown, D. A.; Emma, F.; Klingenberg, C.; Hennekam, R. C.; Torre, G.; Garshasbi, M.; Tzschach, A.; et al. Cranioectodermal Dysplasia, Sensenbrenner Syndrome, Is a Ciliopathy Caused by Mutations in the IFT122 Gene. Am. J. Hum. Genet. 2010, 86 (6), 949–956.
Bujakowska, K. M.; Zhang, Q.; Siemiatkowska, A. M.; Liu, Q.; Place, E.; Falk, M. J.; Consugar, M.; Lancelot, M.-E.; Antonio, A.; Lonjou, C.; et al. Mutations in IFT172 Cause Isolated Retinal Degeneration and Bardet–Biedl Syndrome. Hum. Mol. Genet. 2015, 24 (1), 230–242.
Tam, L.-W.; Wilson, N. F.; Lefebvre, P. a. A CDK-Related Kinase Regulates the Length and Assembly of Flagella in Chlamydomonas. J. Cell Biol. 2007, 176 (6), 819–829.
Yang, Y.; Roine, N.; Mäkelä, T. P. CCRK Depletion Inhibits Glioblastoma Cell Proliferation in a Cilium-Dependent Manner. EMBO Rep. 2013, 14 (8), 741–747.
Ko, H. W.; Norman, R. X.; Tran, J.; Fuller, K. P.; Fukuda, M.; Eggenschwiler, J. T. Broad-Minded Links Cell Cycle-Related Kinase to Cilia Assembly and Hedgehog Signal Transduction. Dev. Cell 2010, 18 (2), 237–247.
Moon, H.; Song, J.; Shin, J.-O.; Lee, H.; Kim, H.-K.; Eggenschwiller, J. T.; Bok, J.; Ko, H. W. Intestinal Cell Kinase, a Protein Associated with Endocrine-Cerebro-Osteodysplasia Syndrome, Is a Key Regulator of Cilia Length and Hedgehog Signaling. Proc. Natl. Acad. Sci. U. S. A. 2014, 111 (23), 8541–8546.
Berman, S. A.; Wilson, N. F.; Haas, N. A.; Lefebvre, P. A.; Paul, S. A Novel MAP Kinase Regulates Flagellar Length in Chlamydomonas. 2003, 13 (Figure 1), 1145–1149.
Du Toit, A. KIF7 Organizes Cilia. Nat. Rev. Mol. Cell Biol. 2014, 15 (8), 498–499. Hilton, L. K.; Gunawardane, K.; Kim, J. W.; Schwarz, M. C.; Quarmby, L. M. The Kinases LF4 and CNK2 Control Ciliary Length by Feedback Regulation of Assembly and Disassembly Rates. Curr. Biol. 2013, 23 (22), 2208–2214.
Hernandez-Hernandez, V.; Pravincumar, P.; Diaz-Font, A.; May-Simera, H.; Jenkins, D.; Knight, M.; Beales, P. L. Bardet-Biedl Syndrome Proteins Control the Cilia Length through Regulation of Actin Polymerization. Hum. Mol. Genet. 2013, 22 (19), 3858–3868.
Shida, T.; Cueva, J. G.; Xu, Z.; Goodman, M. B.; Nachury, M. V. The Major Alpha-Tubulin 
K40 Acetyltransferase AlphaTAT1 Promotes Rapid Ciliogenesis and Efficient Mechanosensation. Proc. Natl. Acad. Sci. U. S. A. 2010, 107 (50), 21517–21522.
Szyk, A.; Deaconescu, A. M.; Spector, J.; Goodman, B.; Valenstein, M. L.; Ziolkowska, N. E.; Kormendi, V.; Grigorieff, N.; Roll-Mecak, A. Molecular Basis for Age-Dependent Microtubule Acetylation by Tubulin Acetyltransferase. Cell 2014, 157 (6), 1405–1415.
Kalebic, N.; Sorrentino, S.; Perlas, E.; Bolasco, G.; Martinez, C.; Heppenstall, P. A. ΑTAT1 Is the Major α-Tubulin Acetyltransferase in Mice. Nat. Commun. 2013, 4 (1), 1962.
Mykytyn, K.; Sheffield, V. C. Establishing a Connection between Cilia and Bardet-Biedl Syndrome. Trends Mol. Med. 2004, 10 (3), 106–109.
Tadenev, A. L. D.; Kulaga, H. M.; May-Simera, H. L.; Kelley, M. W.; Katsanis, N.; Reed, R. R. Loss of Bardet-Biedl Syndrome Protein-8 (BBS8) Perturbs Olfactory Function, Protein Localization, and Axon Targeting. Proc. Natl. Acad. Sci. U. S. A. 2011, 108 (25), 10320–10325.
Dafinger, C.; Liebau, M. C.; Elsayed, S. M.; Hellenbroich, Y.; Boltshauser, E.; Korenke, G. C.; Fabretti, F.; Janecke, A. R.; Ebermann, I.; Nürnberg, G.; et al. Mutations in KIF7 Link Joubert Syndrome with Sonic Hedgehog Signaling and Microtubule Dynamics. J. Clin. Invest. 2011, 121 (7), 2662–2667.