Progress 05/01/21 to 04/30/24
Outputs
Target Audience:Our target audience, consisting of scientists and professionals involved in nematology and related fields, greatly benefited from our project's activities. We presented our data at various small nematology community meetings and in small groups to collaborators and researchers, which facilitated valuable exchanges of information with other scientists and individuals engaged in similar efforts. This exchange not only enriched the collective knowledge but also helped in avoiding the repetition of efforts across different research groups. Additionally, we are currently working on writing publications of our work for more targeted journals, ensuring that our findings reach and benefit the broader scientific community involved in nematode research and management. Changes/Problems:Due to the unforeseen challenges associated with the COVID-19 pandemic, the execution of our project experienced slight delays. Additionally, our Postdoc scientist received a compelling job offer in the industry that aligned well with his future career aspirations. I wholeheartedly encouraged him to pursue this opportunity, recognizing its significance to his professional growth. However, this transition inevitably introduced further slight delays to our project timeline. The optimization of microinjection techniques also took longer than expected, and completing the chromosome-level genome was more time-consuming than projected. These factors have resulted in us being slightly behind the timeline, but we are confident that this project has provided much-needed advancements toward the genome editing of plant-parasitic nematodes. Despite these challenges, we have made significant progress, and we currently have three publications in the pipeline. The first is focused on the genome of Meloidogyne hapla, the second on the transient expression of mRNA and CRISPR knockout in M. hapla, and the third on the development of selection markers in root-knot nematodes. These upcoming publications will contribute to the scientific community's understanding and will be a key outcome of our project's efforts. What opportunities for training and professional development has the project provided? During this period, Graduate Student Ching-Jung Lin was trained to carry out reporter assays (fluorescence, luciferase, GUS) and the use of microinjection apparatus. This hands-on experience has been crucial in maintaining the momentum of our project and ensuring that critical techniques are mastered within our team. How have the results been disseminated to communities of interest?The broader public benefited from our project's activities through targeted outreach efforts aimed at raising awareness about the nematode problem. We participated in UC Davis Biodiversity Day in both 2022 and 2023, where we introduced the public to the challenges posed by plant-parasitic nematodes. More than 1,800 people, including home gardeners, growers, and children, visited our stall during these events. Additionally, we set up a nematode introductory stall at the UC Davis Picnic Day, which attracted around 2,000 attendees. These events provided an excellent platform to educate the public about nematode issues, emphasizing their impact on agriculture and the importance of research in this area. What do you plan to do during the next reporting period to accomplish the goals?
Nothing Reported
Impacts
What was accomplished under these goals?
Previously, we introduced fluorescent nucleotides, specifically Cy3-dUTP, into the germline of M. hapla males via microinjection. In the lifecycle of root-knot nematodes, females remain attached to the root, while males become mobile in search of females, making males more accessible for injections and offering higher survival rates from the procedure. In this project, we have focused on refining the delivery of macromolecules into the gonads of male M. hapla using microinjection. The nematode's resilient cuticle has posed challenges, often resulting in needle breakage. To address this, we have been adjusting various aspects of the injection needle--such as its pull, material, and bevel--and protocol parameters like angle, pressure, immobilizing oil, and agarose pad. These refinements have led to improvement in nematode survival, motility, and the effective incorporation of macromolecules into the germline. Additionally, we have developed a method to sustain females on agar plates for extended periods, enabling them to survive for several weeks and lay eggs, even though they are typically rooted in their natural habitat. We are currently microinject these females. Our next steps involve introducing our CRISPR target into both male and female nematodes, marking significant progress toward our overarching goal of developing a gene-editing toolkit for agriculturally important plant-parasitic nematodes.
Publications
- Type:
Conference Papers and Presentations
Status:
Accepted
Year Published:
2023
Citation:
W4186 Multi-state Nematology Meeting - Nov 15th, Davis
- Type:
Conference Papers and Presentations
Status:
Accepted
Year Published:
2024
Citation:
The Bay Area Worm Meeting, April 22, Davis
- Type:
Conference Papers and Presentations
Status:
Accepted
Year Published:
2023
Citation:
Research Seminar Umea Plant Science Centre, Mar 13th, Umea, Sweden
- Type:
Conference Papers and Presentations
Status:
Accepted
Year Published:
2023
Citation:
W4186 Multi-state Nematology Meeting - Nov 14th, Santa Fe
- Type:
Conference Papers and Presentations
Status:
Accepted
Year Published:
2023
Citation:
35th Symposium of the European Society of Nematologists, Cordoba, Spain
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Progress 05/01/22 to 04/30/23
Outputs
Target Audience:The results of the project were presented at the following events: 1) W4186 Multi-state Nematology Meeting - Nov 15th, Davis 2) The Bay Area Worm Meeting, April 22, Davis 3) Research Seminar Umea Plant Science Centre, Mar 13th, Umea, Sweden Changes/Problems:Due to the unforeseen challenges associated with the COVID pandemic, the execution of our project experienced slight delays. Additionally, our Postdoc scientist received a compelling job offer in the industry that aligned well with his future career aspirations. I wholeheartedly encouraged him to pursue this opportunity, recognizing its significance to his professional growth. However, this transition inevitably introduced further slight delays to our project timeline. On a brighter note, our graduate student, Ching-Jung, has taken charge of the current tasks and has already made significant achievements, showcasing her dedication and capability. What opportunities for training and professional development has the project provided?Graduate Student Ching-Jung Lin was trained to carry out reporter assays (fluorescence, luciferase, GUS). She is currently being trained to use microinjection apparatus. How have the results been disseminated to communities of interest?
Nothing Reported
What do you plan to do during the next reporting period to accomplish the goals?We have already optimized methodology to deliver and translate mRNA in M. hapla J2. We will use this platform now to optimize CRISPR/Cas9 non-homologous end joining in vivo. we will design 5 guides for ben-1 gene (using ATUM gRNA Design Tool), and the minimum number of primer pairs to amplify the edited region/s in amplicons of ≤400 bp. We will package Cas9-encoding mRNA with gRNAs (synthesized by Twist Bioscience) in liposomes and deliver them to 2,000 J2s, each in biological triplicate. We will extract genomic DNA using the DNA kit (Macherey-Nagel) according to manufacturer's instructions and amplify across the target region/s with the relevant primers. Amplicons will be purified using a QIAquick PCR purification kit (Qiagen) and sequenced using 250-bp paired-end Illumina amplicon sequencing (Genewiz). Individual overlapping 250-bp read pairs will be re-capitulated into the <400 bp product, and analysed for indel frequency in the expected region (within 20 bp of PAM). Once an edit is detected, we will optimize the entire procedure for time, temperature, guides, and Cas enzymes for a particular gene. Finally, we will deliver optimized Cas9-mRNA and gRNAs into male gonads of M. hapla. Genomic DNA will be extracted from single injected males and edits will be detected using amplicon sequencing. Once editing is successfully demonstrated, injected males will be placed on young females on tomato root culture plates. These females will be of a different strain (LM) with DNA polymorphisms compared to the male parent (VW9). After 2 weeks, we will collect egg masses, which each contain the brood (~100 -1,000 eggs) of a single female (F0), from the host plant roots. We will extract DNA from half of the pool of eggs and assess it by PCR to confirm that progeny from the male parent are present. The remaining eggs in positive pools will be allowed to hatch and infect tomato plants, where they will develop into F1 adult females under conditions favoring parthenogenic reproduction. The egg masses produced by these females will contain F2 eggs that, due to the recombination mechanism of M. hapla, will resemble Recombinant Inbred Lines and segregate at 1:1 for the parental traits. Parthenogenic reproduction will result in individuals that are homozygous for the introduced disruption.
Impacts
What was accomplished under these goals?
Following progress has been made to accomplish the goals of the project, Objective 1: Optimize CRISPR/Cas9 experiments in somatic cells A successful CRISPR experiment requires three steps, the absence of any one of which will result in failure: 1) delivery of components into cells, 2) targeting an amenable gene to CRISPR, including guides that work in vivo, and 3) detection of possibly rare events. It seems prudent to optimize as many of these as possible, before adding the constraints of germline delivery and long life-cycle. Our preliminary data show that it is possible to deliver exogenous mRNA encoding reporter genes to cyst nematode Heterodera schachtii second stage juvenile (J2) somatic cells, and that the delivered mRNA is translated in sufficient quantity, for a sufficient duration, that we can detect it by multiple means. During the current project period, we have been working to transfer this technology to root-knot nematode M. hapla, and further optimizing transient expression to get the highest output system to serve future objectives. We synthesized capped and polyadenylated mRNA encoding luciferase, eGFP, mCherry, and β-gal (GUS). Nematodes were then soaked in solutions containing one of these mRNAs. Following this, we established detection assays using confocal microscopy and a microplate reader. For comparison, we utilized C. elegans transgenic lines that express fluorescent proteins like luciferase, eGFP, and mCherry. Our results showed that GUS staining in nematodes was undetectable, and due to high autofluorescence in nematodes, eGFP was considered unsuitable. However, consistent detections were made for mCherry and luciferase through microplate reader. In addition, we were also able to visualize nematodes expressing mCherry via confocal microscope. Presently, we are validating our results via western blot, employing monoclonal antibodies. Having successfully optimized transient expression using mRNA, we are ready for the next phase: gene editing using CRISPR. A significant challenge we face is the absence of a selectable target for CRISPR in plant-parasitic nematodes. After an extensive literature review and consultations with experts, we have settled on the beta-tubulin gene as our first CRISPR target (ben-1 locus). Notably, in C. elegans, mutations in the ben-1 locus confer resistance to the drug benzimidazoles, a characteristic that can be harnessed for selection. We have successfully sequenced the genome of M. hapla and identified ben-1 homologs within it. Currently, we are in the process of synthesizing the CRISPR construct and concurrently determining the optimal benzimidazoles concentrations for selection assays on M. hapla. Objective 2: Demonstrate heritable genome editing We have previously introduced fluorescent nucleotides, specifically Cy3-dUTP, into the germline of M. hapla males via microinjection. In the lifecycle of root-knot nematodes, females remain attached to the root, whereas males become mobile in their quest to locate females. This mobility makes males more accessible for injections, and they also tend to have a higher survival rate from this procedure than females. In the current project phase, our focus has been on refining the delivery of macromolecules into the gonads of male M. hapla using microinjection. We observed that the nematode's cuticle is particularly resilient, often resulting in needle breakage. As a result, we're adjusting various aspects of the injection needle --including its pull, material, and bevel --as well as protocol parameters like angle, pressure, immobilizing oil, and agarose pad. These refinements are intended to improve nematode survival, motility, and the effective incorporation of macromolecules into the germline. Beyond our work with males, we have also developed a method to sustain females on agar plates for extended periods, even though in their natural habitat, they stay rooted. These females survive for several weeks, during which they lay eggs. We plan to microinject these females in the near future. In our subsequent steps, we aim to introduce our CRISPR target into both male and female nematodes.
Publications
|
Progress 05/01/21 to 04/30/22
Outputs
Target Audience:The results of the project were presented in a Multi-state Nematology Meeting (W4186) held in Hawaii between November 15th-16th. Changes/Problems:
Nothing Reported
What opportunities for training and professional development has the project provided?Postdoctoral scientist Henok Yimer was trained to use microinjection apparatus. Graduate Student Ching-Jung Lin is being trained to carry out reporter assays (fluorescence, luciferase, GUS). How have the results been disseminated to communities of interest?
Nothing Reported
What do you plan to do during the next reporting period to accomplish the goals?Once we optimize methodology to deliver and translate mRNA in M. hapla J2, we will use this platform to optimize CRISPR/Cas9 non-homologous end joining in vivo. We will select up to 10 genes, prioritized based on those highly expressed throughout the lifecycle and therefore likely to be present in regions of open chromatin. For each gene we will design 5 guides (using ATUM gRNA Design Tool), and the minimum number of primer pairs to amplify the edited region/s in amplicons of ≤400 bp. For each gene, we will package Cas9-encoding mRNA (synthesized as in SO1.1) with gRNAs (synthesized by Twist Bioscience) in liposomes and deliver them to 2,000 J2s, each in biological triplicate. We will extract genomic DNA using the DNA kit (Macherey-Nagel) according to manufacturer's instructions and amplify across the target region/s with the relevant primers. Amplicons will be purified using a QIAquick PCR purification kit (Qiagen) and sequenced using 250-bp paired-end Illumina amplicon sequencing (Genewiz). Individual overlapping 250-bp read pairs will be re-capitulated into the <400 bp product, and analysed for indel frequency in the expected region (within 20 bp of PAM). Once an edit is detected, we will optimize the entire procedure for time, temperature, guides, and Cas enzymes for a particular gene. Finally, we will deliver optimized Cas9-mRNA and gRNAs into male gonads of M. hapla. Genomic DNA will be extracted from single injected males and edits will be detected using amplicon sequencing. Once editing is successfully demonstrated, injected males will be placed on young females on tomato root culture plates. These females will be of a different strain (LM) with DNA polymorphisms compared to the male parent (VW9). After 2 weeks, we will collect egg masses, which each contain the brood (~100 -1,000 eggs) of a single female (F0), from the host plant roots. We will extract DNA from half of the pool of eggs and assess it by PCR to confirm that progeny from the male parent are present. The remaining eggs in positive pools will be allowed to hatch and infect tomato plants, where they will develop into F1 adult females under conditions favoring parthenogenic reproduction. The egg masses produced by these females will contain F2 eggs that, due to the recombination mechanism of M. hapla, will resemble Recombinant Inbred Lines and segregate at 1:1 for the parental traits. Parthenogenic reproduction will result in individuals that are homozygous for the introduced disruption.
Impacts
What was accomplished under these goals?
Following progress has been made to accomplish the goals of the project, Objective 1: Optimize CRISPR/Cas9 experiments in somatic cells A successful CRISPR experiment requires three steps, the absence of any one of which will result in failure: 1) delivery of components into cells, 2) targeting an amenable gene to CRISPR, including guides that work in vivo, and 3) detection of possibly rare events. It seems prudent to optimize as many of these as possible, before adding the constraints of germline delivery and long life-cycle. Our preliminary data show that it is possible to deliver exogenous mRNA encoding reporter genes to cyst nematode Heterodera schachtii second stage juvenile (J2) somatic cells, and that the delivered mRNA is translated in sufficient quantity, for a sufficient duration, that we can detect it by multiple means. During the current project period, we have been working to transfer this technology to root-knot nematode M. hapla, and further optimizing transient expression to get the highest output system to serve future objectives. Capped and polyadenylated mRNA encoding luciferase, eGFP, and β-gal have been synthesized and assays for detection are currently being established and optimized. We think that optimization of these platforms will allow us to rapidly prototype CRISPR experiments in vivo before deploying them to generate heritable mutations. Objective 2: Demonstrate heritable genome editing We previously introduced fluorescent nucleotides, (Cy3-dUTP) by microinjection into the germline of M. hapla males. Males are more likely to survive the process than females and are technically easier to manipulate on the injection apparatus. During the current project period, we have been optimizing the delivery of macromolecules into the gonads of male M. hapla using microinjection. We found that cuticle of nematodes is very tough leading to the breaking of injection needle. The injection needle (pulling, material, bevel) and protocol (angle, pressure, immobilizing oil, agarose pad) is therefore currently being fine-tuned to enhance nematode survival, motility and macromolecule incorporation into the germline. Delivery formulations such as using lipofectamine will be tested to optimize delivery in the next steps.
Publications
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