curriculum vitae

B.Sc. east stroudsburg university 2015

• dual major in marine science, biology / chemistry minor

M.Sc. university of north carolina at wilmington 2020

• marine biology

• thesis: “phagotrophy in the diploid coccolithophore Schphosphaera apsteinii”

senior research scientist II

sept. 2023 – current

• designed and executed experiments related to nanopore sensing of bespoke biomolecules for the purposes of synthetic biology and proteomics.

• complete mastery of all upstream and downstream protein production techniques, from plasmid design to purification using his-tagged and novel synthetic proteins.

• regularly performed Gibson and Golden Gate Assembly, PCR, removal of affinity tags using TEV protease, and other common cloning methods in addition to confirmatory protocols such as capillary and gel electrophoresis.

• experience with performing and designing enzymatic post-translational modifications.

• use of Oxford Nanopore GridION and MinION to identify amino acid and nucleic acid sequences.

• developed techniques for in vitro N- and C-terminal tagging of amyloid beta with machinery for nanopore sequencing using a modified sortase and CuAAC click chemistry.

• performed conjugation of full-gene dsDNA onto gold electrode surfaces for modulating cell-free gene expression.

• atomic force microscopy (AFM) methods for live-imaging the above electrodes under varying charges and chemistries.

• responsible for regular laboratory maintenance and training of new students in the lab.

• served on the “Material Interfaces” committee at the National Science Foundation’s Molecular Programming Flight Plan, to collaboratively refine the field’s landmark research objectives, proposals, and industry partnerships.

environmental microbiology associate

may 2022 – july 2023

• research associate in the biomining division, specializing in non-model microbiology.

• setup, maintenance, and design of continuous cultivation chemostats for the growth of a variety of chemolithoautotrophic and heterotrophic extremophiles.

• assisted in development of Aspergillus niger isolated from environmental samples for oxalic acid production.

• responsible for routine maintenance of lab equipment, namely a flow cytometer system and analytical balances.

• performed a variety of metals testing, ranging from ferrous iron titration to preparing samples for ICP analysis.

• designing and executing a variety of bioleaching and toxicity experiments using model chemolithoautotrophic acidophiles, including conjugation and transformation of isolated axenic cultures.

• mutagenesis and adaptive laboratory evolution of mixed novel extremophilic communities for tolerance of a variety of environmental stressors.

research associate II

july 2020 – june 2022

• fermentation laboratory aide in the development of a variety of renewable biochemical products via the fermentation of anaerobic thermophiles in batch sizes ranging from 50 mL – 10 L.

• cell-free enzyme hydrolysis technology work independently expressing, extracting, quantifying, and purifying target proteins from E. coli using standard culturing and FPLC methods.

• enzyme kinetics performed independently using a variety of forward and reverse reactions to measure Vmax, Kcat, etc. using spectrophotometry.

• analytical chemistry experience collecting and analyzing results of a variety of biomass digestions using HPLC and ion-exclusion chromatography for the identification of small molecules such as sugars and alcohols.

• aseptic cell culture experience with E. coli and anaerobic thermophiles such as plating, cell counting, optical density measurements, etc.

• quantified, characterized, and identified proteins using standard methods such as western blots and SDS-PAGE.

• molecular techniques such as cloning and transforming E. coli cultures, PCR, etc.

• structural biochemistry including independent sample preparation and assisted data collection for negative grid-stained samples and Cryo-EM samples for electron microscopic imaging.

• responsible for laboratory management re: ordering, stocking, buffer and media prep, etc. 

research laboratory technician

aug. 2015 — apr. 2016

• process development technician in Flu Support Lab, working in sterile filtration techniques, instrumentation, etc. 

• utilized egg and aculovirus-insect cell expression systems to express target influenza hemagglutinin proteins.

• experience maintaining GMP in a fast-paced laboratory environment.

• designated trainer for zonal centrifugation techniques. 

aquaculture technician

february 2013 - may 2015

• responsible for maintaining a 700 gal. tank housing several native fish and plant species to the Northeast Appalachia region.

• maintained a variety of marine and freshwater tanks, including (but not limited to) planted, reef, fish-only and invertebrate setups.

• basic feeding, medicine administration, water quality testing, aquarium accessory maintenance.

• custom plumbing for bespoke aquarium systems, ranging use for canister filters and sump systems.

• response to emergency situations such as tank failure, contamination, etc.

graduate researcher

may 2016 - may 2020

A. R. Taylor lab

• developed protocols for multi-channel flow cytometry using the BD FACSCelesta Flow Cytometer, and analysis using FlowJo and Flowing software.

• non-model algaculture of several species of calcium-secreting haptophytes, both haploid and diploid stages of coccolithophores.

• mastery of multifluorescent confocal microscopy using the Leica SP6 system and LAS X software for intracellular imaging of stained vacuoles and fluorescent microspheres.

• co-authored methods paper for the high-throughput fluorescent tracking of coccolith development using flow cytometry.

J. R. Pawlik lab

• AAUS research certified diver working with sponge restoration and boring sponge feeding ecology.

• assisted in monitoring of the giant barrel sponge Xestospongia muta to test the sponge-loop hypothesis.

• co-developed a waterproof arduino-based flowmeter using a thermistor to monitor flow out of the small, benthic boring sponge Cliona delitrix.

• contributed to data collection for "A test of the sponge-loop hypothesis for emergent Caribbean reef sponges" published February 2018.

graduate teaching assistant

september 2016 - may 2020

biodiversity lab

• co-developed curriculum and testing to maintain rigorous academic standards.

• maintained excellent student reviews using university rubrics at the end of each semester.

• introduced students to fundamental skills for recognizing and identifying organisms from every tree of life.

• instructed students on laboratory safety for methods such as dissections and environmental microbiology samples.

• managed husbandry for several in-class organism displays, including hermit crabs, fish, and amphibians.

marine biology lab

• independently directed students in developing skills in marine biology field work and laboratory testing.

• instructed students in fundamental skills for field work in the salt marshes of Wilmington North Carolina.

teaching assistant, general chemistry

january - may 2015

• aid in classroom work, such as grading and test proctoring.

• tutoring responsibilities for both lab and lecture.

• experience leading groups of students in learning exercises.

• molecular techniques (plasmid design, PCR, transfection, cloning, Gibson/Golden Gate assembly, etc.)

• de novo recombinant protein design and expression optimization

• upstream and downstream recombinant protein expression, extraction, purification, etc.

• electrophoresis (SDS-PAGE, capillary, agarose, phos-gels, etc.)

• western blotting

• ÄKTA FPLC system (IEX, nickel, cobalt, etc.)

• Escherichia coli culture

• maxi and mini DNA preparations

• ONT MinION DNA and peptide sequencing

• product concentration via tangential flow filtration, centrifugal units, etc.

• electron microscopy sample preparation and analysis (Cryo-EM, negative staining TEM)

• atomic force microscopy

• analytical chemistry instrumentation (HPLC, MS, ICP)

• laboratory management (training, ordering, stocking, chemical hygiene, etc.)

• LIMS and digital notebook use (quartzy, confluence, labkey, benchling, etc)

• aseptic technique

• proficient wet lab skills (micropipette use, sterile techniques, GMP, ultracentrifuge use, etc.)

• Qubit and nanodrop concentration measurements

• SOP development