Davis Scientific Background and Research Accomplishments

Background

My Ph.D. training was in parasitology with an emphasis on parasitic worms (helminths).  I then undertook extensive post-graduate training in molecular biology and have applied this expertise to study the molecular biology of helminths with an emphasis on their RNA biology, genomics, and DNA elimination. Parasitic worms infect a third of the world’s population leading to death, long term morbidity, and they inhibit physical and cognitive development in young children.  They remain a significant problem in many parts of the world and have been described as “The Neglected Diseases of Poverty”.

A major focus in my lab has been to develop and use molecular methods to investigate atypical mechanisms of gene expression in helminths. The goal of our work was to provide insight into novel mechanisms of gene regulation that may be drug targets in helminths. The primary focus of this work was on spliced leader RNA trans-splicing and later programmed DNA elimination, a process where chromosomes break and portions of chromosomes are eliminated in somatic cells during early development. The research included investigating the contribution of small RNAs to programmed DNA elimination, the general role and function of the diversity of small RNAs present in parasitic worms, and a variety of studies aimed at characterizing the mechanism and function of DNA elimination. Ascaris is an interesting developmental biology model exhibiting unusual early transcription prior to pronuclear fusion and programmed DNA elimination. Over the years the lab developed a variety of molecular tools to carry out studies on Ascaris and was supported by funding from the National Institutes of Health, National Institute of Allergy and Infectious Disease.

Education and Training
BA, Lafayette College, Easton, PA (Biology)
MS, The University of Massachusetts, Amherst (Zoology/Parasitology)
PhD, The University of Massachusetts, Amherst (Zoology/Parasitology)
Additional Training: University of Colorado School of Medicine (Somatic Cell Genetics) and Case Western Reserve University (Molecular Biology)

Faculty Appointment
Professor Emeritus
Department of Biochemistry and Molecular Genetics
RNA Bioscience Initiative
University of Colorado School of Medicine


Research Accomplishments

Analysis of programmed DNA elimination in Ascaris

Genomes rarely change. However, a few organisms undergo a wholesale genome change called DNA elimination, a programmed process that eliminates specific DNA sequences from the genome. We have shown that 18% of the genome of the parasitic nematode Ascaris is eliminated in somatic cell lineages during the third through fifth cleavages (4 to 16 cell stage), while the germline genome remains intact. Both specific repetitive and unique sequences (including ~1000 genes) are lost during the elimination process that forms the somatic genome. The DNA elimination events are identical in all individuals and in five distinct cell lineages. The eliminated genes are primarily expressed in the Ascaris germline and early embryo. Comparative genome analysis of DNA elimination in several other parasitic nematodes (A. lumbricoides, Toxocara, and Parascaris) demonstrated that they also eliminate 1,000-2,000 genes (5-10% of their genes). Overall, our data suggest that DNA elimination in parasitic nematodes is an essential, irreversible mechanism for silencing a subset of germline and early embryo expressed genes in somatic tissues. Comprehensive chromosome analysis of DNA elimination in Ascaris demonstrated that 72 DNA breaks occur and all 24 germline chromosome ends undergo remodeling through subtelomeric DNA breaks, loss of terminal sequences, and healing of new chromosome ends by de novo telomere healing. Eliminated DNA is incorporated into micronuclei during telophase of a DNA elimination mitosis and the DNA is eventually degraded in autophagosomes. Worm specific Argonautes are on condensed chromosomes during elimination mitoses and are present either on chromosomes that will be retained (WAGO-2) or eliminated (WAGO-3). We hypothesize that the Argonautes and/or their small RNAs contribute to DNA elimination. Ascaris has holocentric chromosomes with multiple centromeric/kinetochore regions distributed along the length of the chromosome in the gametogenic germline. Ascaris holocentric chromosomes are dynamic. During DNA elimination, only those chromosome regions to be retained exhibit extensive deposition of the centromeric histone CENP-A and are holocentric; chromosome regions destined for elimination have lost CENP-A and are not segregated.  Thus, both specific sites for DNA breaks and changes in CENP-A localization on the chromosomes are involved in a concerted mechanism to define portions of chromosomes for retention or elimination in programmed DNA elimination.

Selected Publications

a). Wang, J., Czech, B. Crunk, A., Wallace, A., Mitreva, M., Hannon, G., and R.E. Davis. 2011. Deep small RNA sequencing from the nematode Ascaris reveals conservation, functional diversification, and novel developmental profiles. Genome Research 21:1462–1477.
b). Wang, J., Mitreva, M.,Berriman, M., Thorne, A., Magrini, V., Koutsovoulos, G., Kumar, S., Blaxter, M.L., and R.E. Davis. 2012. Germline gene silencing by DNA elimination in the nematode Ascaris. Developmental Cell 23:1072–1080.
c). Kang, Y., Wang, J., Neff, A., Kratzer, S., Kimura, H., and R.E. Davis. 2016. Differential chromosomal localization of centromeric histone CENP-A contributes to nematode programmed DNA elimination. Cell Reports. 16:2308-2316.
d). Wang, J., Gao, S., Mostovoy, Y. Y. Kang, Y., Zagoskin, M., Sun, Y., Zhang, Y, Zhang, B. While, L.K., Eaton, A., Nutman, T., Kwok, P.-Y., Hu, S., Nielsen, M.K. and R.E. Davis. 2017. Comparative genome analysis of programmed DNA elimination in nematodes. Genome Research. 27:2001-2014.
e). Wang, J., Veronezi, G.M.B., Kang, Y., Zagoskin, M., O’Toole, E.T., and R.E. Davis. 2020. Comprehensive chromosome end remodeling during programmed DNA elimination. Current Biology, 30, 3397–3413.


Genome, transcriptome, and small RNA resources for Ascaris

Neither genomic nor transcriptome resources were available for our studies on Ascaris.  Therefore, we first developed and refined a variety of resources including genome (somatic and germline), mRNA transcriptome, and small RNA profiles.

Our analysis of small RNAs and Argonaute proteins in Ascaris demonstrated several unusual features. miRNAs are synthesized immediately after fertilization in utero, before pronuclear fusion, and before the first cleavage of the zygote. This is the earliest expression of small RNAs ever described, and at a developmental stage long thought to be transcriptionally quiescent. Strikingly, piRNAs, Piwi-clade Argonautes, and other proteins associated with the piRNA pathway have been lost in Ascaris. A comparison of the two classes of Ascaris endo-siRNAs identified, 22G-RNAs and 26G-RNAs, with those present in C. elegans, suggests great diversification and plasticity in the use of small RNA pathways during spermatogenesis in different nematodes. Our studies significantly expand our understanding of the conservation, divergence, and flexibility of nematode Argonautes and small RNA pathways. Nematode Argonautes and small RNA pathways may have initially evolved for specific types of gene regulation leading to better fitness, but over time may have been co-opted for other functions in diverse nematodes.

Ascaris early development is very slow and synchronous. We took advantage of this feature and RNA-seq and ribosome profiling to obtain a comprehensive time course of mRNA expression, degradation, and translation during A. suum early development. Unexpectedly, we found that A. suum maternal mRNAs are degraded right after fertilization, and ~4,000 genes are transcribed prior to pronuclear fusion and in the 1-4 cell embryos. Importantly, our data challenge a long-standing view of early development that transcription is silenced in the oocyte and major transcription only occurs after the 2-cell stage or later. Our data suggest that major transcription can occur before pronuclear fusion and in the 1-2-cell stages. Our data define the earliest and most complex transcription described in any metazoan and raise the hypothesis that cell cycle length is a key factor that determines the timing and amount of transcription during early development. Early development in Ascaris and C. elegans is identical. However, our data provide novel insights into re-wiring of early gene expression programs in the two nematodes as this regulation occurs at the level of transcription in Ascaris whereas it occurs post-transcriptionally in C. elegans. 

Selected Publications

a). Wang, J., Czech, B. Crunk, A., Wallace, A., Mitreva, M., Hannon, G., and R.E. Davis. 2011. Deep small RNA sequencing from the nematode Ascaris reveals conservation, functional diversification, and novel developmental profiles. Genome Research 21:1462–1477.
b). Wang, J., Mitreva, M.,Berriman, M., Thorne, A., Magrini, V., Koutsovoulos, G., Kumar, S., Blaxter, M.L., and R.E. Davis. 2012. Germline gene silencing by DNA elimination in the nematode Ascaris. Developmental Cell 23:1072–1080.
c). Wang, J., Garrey, J., and R.E. Davis. 2014. Transcription in pronuclei and one- to four-cell embryos drives early development in a nematode. Current Biology. 24:124-133.
d). Easton, A.V., Gao, S., Lawton, S.P., Bennuru, A., Khan, A., Dahlstrom, E., Oliveira, R.G., Kepha, S.F., Webster, J.P., Anderson, R.M., Grigg, M.E., Davis*, R.E., Wang. J., and T.B. Nutman. 2020. Molecular evidence of hybridization between pig and human Ascaris indicates an interbred species complex infecting humans. eLife, 9:e61562.
e). Zagoskin, M.V., Wang, J., Neff, A.T., Veronezi, G.M.B., and R.E. Davis. 2022. Small RNA pathways in the nematode Ascaris in the absence of piRNAs. Nature Communications, 13:837.


Discovery and analysis of spliced leader RNA trans-splicing in schistosomes

A major focus of my lab over the years was to investigate the functional significance, phylogenetic distribution, and cell and molecular adaptations to spliced leader RNA trans-splicing.  Following our discovery of trans-splicing in schistosomes, we carried out studies to examine the function, conserved features, and phylogenetic distribution of trans-splicing in flatworms.  We found that 15% of mRNAs are trans-spliced in schistosomes and the substrate spliced leader RNA is expressed in all tissues. The proteins encoded by trans-spliced mRNAs are not associated with any particular types of proteins or pathways, and there appear to be no differentiating characteristics of the splice acceptor sites of trans-spliced genes (results that have been corroborated by more recent genome studies on schistosomes).  We then developed methods to identify trans-splicing in several other parasitic flatworms and carried out a phylogenetic comparison of their spliced leader RNAs. These studies demonstrated that flatworm spliced leader RNAs exhibit significant flexibility in their structures and particularly in the spliced leader sequence in comparison with other trans-splicing metazoa. We demonstrated that one function of trans-splicing in flatworms is to resolve polycistronic transcripts into monocistronic mRNAs. Overall, these studies defined unique and conserved aspects of trans-splicing in flatworms providing novel insight into the sequence, structure, and function of spliced leader RNA trans-splicing in these helminths as well as general insights into spliced leader RNA trans-splicing in metazoa.

Selected Publications

a). A. Rajkovic, R. E. Davis, J. N. Simonsen, F. M. Rottman. 1990. A spliced leader is present on a subset of mRNAs from the human parasite Schistosoma mansoni. Proc Natl Acad Sci U S A 87:8879-8883.
b). R. E. Davis, C. Hardwick, P. Tavernier, S. Hodgson, H. Singh. 1995. RNA trans-splicing in flatworms. Analysis of trans-spliced mRNAs and genes in the human parasite, Schistosoma mansoni. J Biol Chem 270:21813-21819.
c). R. E. Davis. 1997. Surprising diversity and distribution of spliced leader RNAs in flatworms. Mol Biochem Parasitol 87:29-48.
d). R. E. Davis, S. Hodgson. 1997. Gene linkage and steady state RNAs suggest trans-splicing may be associated with a polycistronic transcript in Schistosoma mansoni. Mol Biochem Parasitol 89:25-39.


Key insights into translation of trans-spliced mRNAs

Unlike most groups of organisms with trans-splicing, the spliced leader sequence is highly divergent in different flatworms. However, a conserved feature is that the 3’ terminal nucleotides are invariably AUG.  This raised the interesting question of whether a purpose of trans-splicing in flatworms is to provide mRNA open reading frames with a translation initiation codon.  Answering this question required development of new technology for transfecting parasitic helminths. We developed and optimized particle bombardment to transfect parasitic helminths. We used these methods and bioinformatic analyses to show that the flatworm spliced leader AUG codon is in fact used as a translation initiator methionine in schistosomes despite the fact that it is not in a context known to be functional for translation in other metazoans.  These studies demonstrated that flatworm trans-splicing provides some mRNAs with an initiator methionine for translation. Spliced leaders have an atypical cap that is a trimethylguanosine cap in metazoans. The presence of an atypical cap and conserved sequence at the 5’ end of trans-spliced mRNAs led to the speculation that trans-splicing might serve to enhance mRNA translation or stability of recipient mRNAs. We developed several new methods/tools including 1) RNA transfection and 2) a cap-dependent translation system in Ascaris embryos to address these longstanding questions. Overall, our data suggest that trans-splicing does not improve mRNA translation or stability, and that efficient translation of TMG-capped mRNAs requires the downstream spliced leader sequence. We defined a core set of nucleotides and a stem-loop within the 22-nucleotide nematode SL that stimulate translation of mRNAs with a TMG cap. In addition, we identified unique adaptations in the nematode eukaryotic translation initiation factor 4E/G (eIF4E/G) complex that enables efficient translation of the TMG-SL RNAs. Our results demonstrated that the mRNA 5’ untranslated region (UTR) can play a positive and novel role in translation initiation through interaction with the eIF4E/G complex in nematodes and raised the issue of whether eIF4E/G-RNA interactions play a role in the translation of other eukaryotic mRNAs.  Overall, a major contribution of our studies was to demonstrate that the 5’ UTR can enhance translation (previous work has historically viewed the 5’ UTR as a largely negative regulatory element) and provided key insights into how trans-spliced mRNAs are translated. 

Selected Publications

a). R. E. Davis, A. Parra, P. T. LoVerde, E. Ribeiro, G. Glorioso, S. Hodgson. 1999. Transient expression of DNA and RNA in parasitic helminths by using particle bombardment. Proc Natl Acad Sci U S A 96: 8687-8692.
b). G. Cheng, L. Cohen, D. Ndegwa, R. E. Davis. 2006. The flatworm spliced leader 3'-terminal AUG as a translation initiator methionine. J Biol Chem 281:733-743.
c). S. Lall, C. C. Friedman, M. Jankowska-Anyszka, J. Stepinski, E. Darzynkiewicz, R. E. Davis. 2004. Contribution of trans-splicing, 5' -leader length, cap-poly(A) synergism, and initiation factors to nematode translation in an Ascaris suum embryo cell-free system. J Biol Chem 279:45573-45585.
d). A. Wallace, M. E. Filbin, B. Veo, C. McFarland, J. Stepinski, M. Jankowska-Anyszka, E. Darzynkiewicz, R. E. Davis. 2010. The nematode eukaryotic translation initiation factor 4E/G complex works with a trans-spliced leader stem-loop to enable efficient translation of trimethylguanosine-capped RNAs. Mol Cell Biol 30:1958-1970.


Biochemical and structural studies on atypical cap-binding proteins

Metazoan spliced leader trans-splicing generates mRNA 5’ ends through addition of both a nucleotide sequence and an atypical, trimethylguanosine cap that has two additional methyl groups at the N2 position.  When we began our studies, the consensus in the field was that the eukaryotic cap-binding protein, eIF4E, was not able to efficiently recognize this atypical cap.  Similarly, one would predict that other cap-interacting proteins, such as decapping proteins, would require adaptation in trans-splicing helminths to recognize and act on the atypical TMG cap.  Using an in vitro system from Ascaris embryos we developed, we identified and characterized two decapping activities in Ascaris embryo extracts: Dcp2 for RNA decapping and DcpS for dinucleotide decapping. We showed that nematode DcpS has a novel adaptation as it hydrolyzes both m7GpppG and m2,2,7GpppG dinucleoside triphosphates, whereas human DcpS can act only on m7GpppG. We found that nematode Dcp2 RNA activity is influenced by both 5’ end sequence and its context. The trans-spliced leader sequence on mRNAs reduces Dcp2 RNA decapping activity ~10-fold, suggesting that 5’-to-3’ turnover of trans-spliced RNAs might be regulated. Nematode Dcp2 decaps both m7GpppG- and m2,2,7GpppG-capped RNAs. Surprisingly, we showed that budding yeast and human Dcp2 are also active on m2,2,7GpppG-capped RNAs indicating that the ability to act on trimethylguanosine substrates is not a unique adaptation in nematodes to trans-splicing. Additional kinetic, biochemical, and functional studies were also carried out on these proteins.

Nematodes including Ascaris have multiple isoforms of the key and rate-limiting translation initiation factor that binds the cap, eIF4E. We demonstrated that a single Ascaris eIF4E protein is able to efficiently translate both TMG-capped trans-spliced mRNAs and monomethyl non-trans-spliced mRNAs.  X-ray crystal structure and Nuclear Magnetic Resonance (NMR) data indicate that Ascaris eIF4E binds the two different caps in a similar manner (except for the loss of a single hydrogen bond with the m2,2,7G-cap). Surprisingly, both nematode and mammalian eIF4E both have a low affinity for m2,2,7G-cap. Nematode eIF4E binding to the monomethyl cap, the trimethyl cap, and the trimethyl capped SL 22-nt RNA leads to distinct eIF4E conformational changes. We proposed that interactions between Ascaris eIF4E and the SL allosterically modifiy eIF4G function and represent a unique nematode adaptation that is required to contribute to the efficient translation initiation of trans-spliced mRNAs. Our studies represent the most comprehensive and in depth biochemical characterization and analysis of translation and decay of trans-spliced mRNAs in any system to date.

Selected Publications

a). L. S. Cohen, C. Mikhli, C. Friedman, M. Jankowska-Anyszka, J. Stepinski, E. Darzynkiewicz, R. E. Davis. 2004. Nematode m7GpppG and m3(2,2,7)GpppG decapping: activities in Ascaris embryos and characterization of C. elegans scavenger DcpS. RNA 10:1609-1624.
b). L. S. Cohen, C. Mikhli, X. Jiao, M. Kiledjian, G. Kunkel, R. E. Davis. 2005. Dcp2 Decaps m2,2,7GpppN-capped RNAs, and its activity is sequence and context dependent. Mol Cell Biol 25:8779-8791.
c). S. Lall, F. Piano, R. E. Davis. 2005. Caenorhabditis elegans decapping proteins: localization and functional analysis of Dcp1, Dcp2, and DcpS during embryogenesis. Mol Biol Cell 16:5880-5890.
d). W. Liu, M. Jankowska-Anyszka, K. Piecyk, L. Dickson, A. Wallace, A. Niedzwiecka, J. Stepinski, R. Stolarski, E. Darzynkiewicz, J. Kieft, R. Zhao, D. N. Jones, R. E. Davis. 2011. Structural Basis for Nematode eIF4E binding an m2,2,7G-Cap and its Implications for Translation Initiation. Nucleic Acids Research 39:8820–8832.


Summary

A major focus of our research has been to develop and use new tools and systems for in depth molecular studies in parasitic worms. In many cases, we were the first to develop and use these methods in parasitic worms. Notably, our tool set in Ascaris remains one of the most robust and diverse set of tools developed in a parasitic helminth.  Furthermore, in spite of Ascaris not being a model system, our development and use of these tools enabled us to address long-standing questions and generate data that have provided unique and informative insight into trans-splicing and DNA elimination that could not be addressed in more conventional model systems.


Complete List of Published Work in MyBibliography at the National Library of Medicine

Google Scholar Citations


For information on Ascaris, See the Current Biology "Quick Guide for Ascaris"

For information on Parasitology, See Parasitology Information


Richard E. Davis
Professor Emeritus
RNA Bioscience Initiative
Department of Biochemistry and Molecular Genetics
University of Colorado School of Medicine
Anschutz Medical Campus