My primary research interest is to model human diseases in small tropical fish known as zebrafish.
Approximately 70 percent of human disease genes have a counterpart in these tiny aquatic creatures. Zebrafish offer several advantages as a model system. One such benefit is the fact that the embryos are transparent and develop outside the body, allowing researchers to easily monitor different stages of embryonic development. Another is the maintenance of zebrafish is relatively easy and inexpensive compared to mice, making it possible to house thousands of animals in a laboratory environment.
My lab uses zebrafish to understand the disease pathology and mechanisms of human hearing loss. Hearing loss is one of the most common conditions affecting the population, particularly aging adults. According to the latest statistics from the National Institute on Deafness and Other Communication Disorders (NIDCD), approximately one in six American adults age 18 and older have some significant level of hearing loss.
The sensory macula involved in hearing and balance consists of two major cell types, the mechanosensory receptors called hair cells and another cell type called the supporting cells. Because zebrafish hair cells are functionally similar to the hair cells found in human inner ear—and their excellent genetic and genomic resources—zebrafish provide an excellent model system to understand the hearing loss disease pathology.
High-throughput Validation of Human Candidate Disease Genes in Zebrafish
Since the completion of the Human Genome Project and with the advent of low-cost whole-genome sequencing technologies, hundreds of genome-wide association studies (GWAS) and whole-exome sequencing projects have been revealing the full spectrum of mutations associated with human disease genes. Human geneticists are now facing the immense challenge of validating these candidate disease genes. For the vast majority of genes the clinical data is only correlative, i.e. it is insufficient to demonstrate causality of the disease state.
Our lab is focused on tackling the challenge of systematically validating the flood of human candidate disease genes identified in various genome- wide association (GWAS) and whole-exome sequencing studies with the additional goal of generating animal models for the validated disease genes.
We have developed and validated a high-throughput strategy for mutagenizing the zebrafish genome using CRISPR/Cas9 (Varshney et al. Gen Res 2015), making targeted screening of hundreds of gene knockouts possible with relatively modest resource investments.
Previous forward and reverse genetic screens in zebrafish, while labor intensive, have generated thousands of papers and tens to hundreds of useful disease models in zebrafish, but with the transformative development of gene targeting approaches using CRISPR/Cas9, completely new screening paradigms can be developed. We will use the high-throughput, targeted mutagenesis techniques that I developed at the National Human Genome Research Institute to systematically mutagenize genes in the zebrafish genome related to human deafness. As a proof-of-principle, we selected 50 human, non-syndromic deafness genes from the carefully curated list at hereditaryhearingloss.org, and have already generated knockouts of all 50 orthologs in zebrafish. We are systematically testing these knockouts for the phenotypic characteristics of deaf fish, e.g. failed startle responses, aberrant swimming or developmental defects in the ear or lateral line.
Development of Novel CRISPR-based Strategies for Functional Genomics in Zebrafish
We believe that it is essential to continually push technology development that will break open new avenues of research, so we are interested in developing new cutting edge techniques such as CRISPR/Cas9 based methods to integrate fluorescent tags (knock-in) to study the expression patterns and subcellular protein dynamics of candidate genes, or artificial transcription factors using modified Cas9 proteins. It is critical to develop the efficient techniques for “humanizing” zebrafish mutants (it is currently possible but difficult) to test the wide variety of missense alleles believed to result in altered gene function.
M.Sc., GB Pant University of Agriculture & Technology, Pantnagar, India, 1999
Ph.D., Umeå University, Umeå, Sweden, 2008
Postdoc, (Mentor: Shawn Burgess) National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, 2009-2016
Honors and Awards
2016 Genome Recognition of Employee Accomplishments and Talents Award (GREAT), NHGRI/NIH
2016 DeLill Nasser Award for Professional Development in Genetics, Genetics Society of America
2015 NHGRI Symposium Best Poster Award
2014 NHGRI Intramural Research Trainee Award
2014 NIH Fellows Award for Research Excellence (FARE)
2012 NIH Fellows Award for Research Excellence (FARE)
2012 Genome Recognition of Employee Accomplishments and Talents Award (GREAT), NHGRI/NIH
2007 Research Grant, JC Kempes Minnes Stipendiefond, Sweden
2007 Travel Award, Wallenberg Foundation, Sweden
1997-1999 Department of Biotechnology Fellowship, Government of India
Joined OMRF's Scientific Staff in 2017
Watkins-Chow DE, Varshney GK, Garrett LJ, Chen Z, Jimenez EA, Rivas C, Bishop KS, Sood R, Harper UL, Pavan WJ, Burgess SM. Highly Efficient Cpf1-Mediated Gene Targeting in Mice Following High Concentration Pronuclear Injection. G3 (Bethesda). 2017 Feb 9;7(2):719-22. PMCID:PMC5295614
Watkins-Chow DE*, Varshney GK*, Garrett LJ, Chen Z, Jimenez EA, Rivas C, Bishop KS, Sood R, Harper UL, Pavan WJ, Burgess SM. Highly Efficient Cpf1-Mediated Gene Targeting in Mice Following High Concentration Pronuclear Injection. G3(Bethesda). 2017 Feb 9;7(2):719-722. PMCID:PMC5295614* Joint first authors
Varshney GK, Carrington B, Pei W, Bishop K, Chen Z, Fan C, Xu L, Jones M, LaFave MC, Ledin J, Sood R, Burgess SM. A high-throughput functional genomics workflow based on CRISPR/Cas9-mediated targeted mutagenesis in zebrafish. Nat Protoc. 2016 Dec;11(12):2357-75. PMID: 27809318
Varshney GK, Burgess SM. DNA-guided genome editing using structure-guided endonucleases. Genome Biol. 2016 Sep 15;17(1):187. PMCID:PMC5025577
Pei W, Xu L, Varshney GK, Carrington B, Bishop K, Jones M, Huang SC, Idol J, Pretorius PR, Beirl A, Schimmenti LA, Kindt KS, Sood R, Burgess SM. Additive reductions in zebrafish PRPS1 activity result in a spectrum of deficiencies modeling several human PRPS1-associated diseases. Sci Rep. 2016 Jul18;6:29946. PMCID: PMC4947902
Carrington B, Varshney GK, Burgess SM, Sood R. CRISPR-STAT: an easy and reliable PCR-based method to evaluate target-specific sgRNA activity. Nucleic Acids Res. 2015 Dec 15;43(22):e157. PMCID:PMC4678847
Varshney GK, Zhang S, Pei W, Adomako-Ankomah A, Fohtung J, Schaffer K, Carrington B, Maskeri A, Slevin C, Wolfsberg T, Ledin J, Sood R, Burgess SM. CRISPRz: A database of validated sgRNAs in zebrafish, Nucleic Acids Research Oct. 4, 2015. PMCID:PMC4484386
Varshney GK, Pei W, LaFave MC, Idol J, Xu L, Gallardo V, Carrington B, Bishop K, Jones M, Li M, Harper U, Huang SC, Prakash A, Chen W, Sood R, Ledin J, Burgess SM. High-throughput gene targeting and phenotyping in zebrafish using CRISPR/Cas9. Genome Res. 2015 Jul;25(7):1030-42. PMCID:PMC4484386
Gallardo VE, Varshney GK, Lee M, Bupp S, Xu L, Shinn P, Crawford NP, Inglese J, Burgess SM. Phenotype-driven chemical screening in zebrafish for compounds that inhibit collective cell migration identifies multiple pathways potentially involved in metastatic invasion. Dis Model Mech. 2015 Jun;8(6):565-76.
Functional & Chemical Genomics Research Program, MS 46
Oklahoma Medical Research Foundation
825 N.E. 13th Street
Oklahoma City, OK 73104
Phone: (405) 271-2185
Fax: (405) 271-7128